A Model of Reverse Vascular Remodeling in Pulmonary Hypertension Due to Left Heart Disease by Aortic Debanding in Rats

Pengchao Sang; Mariya M. Kucherenko; Juquan Yao; Qiuhua Li; Szandor Simmons; Wolfgang M. Kuebler; Christoph Knosalla

doi:10.3791/63502

JoVE Journal > Medicine

Medicine

A Model of Reverse Vascular Remodeling in Pulmonary Hypertension Due to Left Heart Disease by Aortic Debanding in Rats

Published: March 01, 2022

doi:

10.3791/63502

Pengchao Sang^2,3, Mariya M. Kucherenko^2,3, Juquan Yao, Qiuhua Li, Szandor Simmons³, Wolfgang M. Kuebler*^2,3, Christoph Knosalla*^1,3,4

¹Department of Cardiothoracic and Vascular Surgery,German Heart Center Berlin (DHZB), ²Institute of Physiology,Charité – Universitätsmedizin Berlin, corporate member of Freie Universität Berlin, Humboldt-Universität zu Berlin, and Berlin Institute of Health, ³DZHK (German Centre for Cardiovascular Research), Partner Site Berlin, ⁴Charité – Universitätsmedizin Berlin, corporate member of Freie Universität Berlin, Humboldt-Universität zu Berlin, and Berlin Institute of Health

Summary

The present protocol describes a surgical procedure to remove ascending-aortic banding in a rat model of pulmonary hypertension due to left heart disease. This technique studies endogenous mechanisms of reverse remodeling in the pulmonary circulation and the right heart, thus informing strategies to reverse pulmonary hypertension and/or right ventricular dysfunction.

Abstract

Pulmonary hypertension due to left heart disease (PH-LHD) is the most common form of PH, yet its pathophysiology is poorly characterized than pulmonary arterial hypertension (PAH). As a result, approved therapeutic interventions for the treatment or prevention of PH-LHD are missing. Medications used to treat PH in PAH patients are not recommended for treatment of PH-LHD, as reduced pulmonary vascular resistance (PVR) and increased pulmonary blood flow in the presence of increased left-sided filling pressures may cause left heart decompensation and pulmonary edema. New strategies need to be developed to reverse PH in LHD patients. In contrast to PAH, PH-LHD develops due to increased mechanical load caused by congestion of blood into the lung circulation during left heart failure. Clinically, mechanical unloading of the left ventricle (LV) by aortic valve replacement in aortic stenosis patients or by implantation of LV assist devices in end-stage heart failure patients normalizes not only pulmonary arterial and right ventricular (RV) pressures but also PVR, thus providing indirect evidence for reverse remodeling in the pulmonary vasculature. Using an established rat model of PH-LHD due to left heart failure triggered by pressure overload with subsequent development of PH, a model is developed to study the molecular and cellular mechanisms of this physiological reverse remodeling process. Specifically, an aortic debanding surgery was performed, which resulted in reverse remodeling of the LV myocardium and its unloading. In parallel, complete normalization of RV systolic pressure and significant but incomplete reversal of RV hypertrophy was detectable. This model may present a valuable tool to study the mechanisms of physiological reverse remodeling in the pulmonary circulation and the RV, aiming to develop therapeutic strategies for treating PH-LHD and other forms of PH.

Introduction

Heart failure is the leading cause of death in developed countries and is expected to increase by 25% over the next decade. Pulmonary hypertension (PH) – a pathological increase of blood pressure in the pulmonary circulation – affects approximately 70% of patients with end-stage heart failure; the World Health Organization classifies PH as pulmonary hypertension due to left heart disease (PH-LHD)¹. PH-LHD is initiated by impaired systolic and/or diastolic left ventricular (LV) function that results in elevated filling pressure and passive congestion of blood into the pulmonary circulation². Albeit initially reversible, PH-LHD gradually becomes fixed due to active pulmonary vascular remodeling in all compartments of the pulmonary circulation, i.e., arteries, capillaries, and veins³^,⁴. Both reversible and fixed PH increase RV afterload, initially driving adaptative myocardial hypertrophy but ultimately causing RV dilatation, hypokinesis, fibrosis, and decompensation that progressively lead to RV failure¹^,²^,⁵^,⁶. As such, PH accelerates disease progression in heart failure patients and increases mortality, particularly in patients undergoing surgical treatment by implantation of left ventricular assist devices (LVAD) and/or heart transplantation⁷^,⁸^,⁹. Currently, no curative therapies exist that could reverse the process of pulmonary vascular remodeling, so fundamental mechanistic research in appropriate model systems is needed.

Importantly, clinical studies show that PH-LHD as a frequent complication in patients with aortic stenosis can improve rapidly in the early post-operative period following aortic valve replacement¹⁰. Analogously, high (>3 Wood Units) pre-operative pulmonary vascular resistance (PVR) that was, however, reversible on nitroprusside was sustainably normalized after heart transplantation in a 5-year follow-up study¹¹. Similarly, an adequate reduction of both reversible and fixed PVR and improvement of RV function in LHD patients could be realized within several months by unloading the left ventricle using implantable pulsatile and non-pulsatile ventricular assist devices¹²^,¹³^,¹⁴. Currently, the cellular and molecular mechanisms that drive reverse remodeling in the pulmonary circulation and RV myocardium are unclear. Yet, their understanding may provide important insight into physiological pathways that may be therapeutically exploited to reverse lung vascular and RV remodeling in PH-LHD and other forms of PH.

A suitable preclinical model that adequately replicates the pathophysiological and molecular features of PH-LHD can be used for translational studies in pressure overload-induced congestive heart failure due to surgical aortic banding (AoB) in rats⁴^,¹⁵^,¹⁶. In comparison to similar heart failure due to pressure overload in the murine model of transverse aortic constriction (TAC)¹⁷, banding of the ascending aorta above the aortic root in AoB rats does not produce hypertension in the left carotid artery as the banding site is proximal of the outflow of the left carotid artery from the aorta. As a result, AoB does not cause left-sided neuronal injury in the cortex as is characteristic for TAC¹⁸, and which may affect the study outcome. Compared to other rodent models of surgically induced PH-LHD, rat models in general, and AoB in particular, prove to be more robust, reproducible and replicate the remodeling of the pulmonary circulation characteristic for PH-LHD patients. At the same time, perioperative lethality is low¹⁹. Increased LV pressures and LV dysfunction in AoB rats induce PH-LHD development, resulting in elevated RV pressures and RV remodeling. As such, the AoB rat model has proven extremely useful in a series of previous studies by independent groups, including ourselves, to identify pathomechanisms of pulmonary vascular remodeling and test potential treatment strategies for PH-LHD⁴^,¹⁵^,²⁰^,²¹^,²²^,²³^,²⁴^,²⁵.

In the present study, the AoB rat model was utilized to establish a surgical procedure of aortic debanding to study mechanisms of reverse remodeling in the pulmonary vasculature and the RV. Previously, myocardial reverse remodeling models such as aortic debanding in mice²⁶ and rats²⁷ have been developed to investigate the cellular and molecular mechanisms regulating the regression of left ventricular hypertrophy and test potential therapeutic options to promote myocardial recovery. Moreover, a limited number of earlier studies have explored the effects of aortic debanding on PH-LHD in rats and showed that aortic debanding might reverse medial hypertrophy in pulmonary arterioles, normalize the expression of pre-pro-endothelin 1 and improve pulmonary hemodynamics²⁷^,²⁸, providing evidence for the reversibility of PH in rats with heart failure. Here, the technical procedures of the debanding surgery are optimized and standardized, e.g., by applying a tracheotomy instead of endotracheal intubation or by using titanium clips of a defined inner diameter for aortic banding instead of polypropylene sutures with a blunt needle²⁶^,²⁷, thus providing for better control of the surgical procedures, increased reproducibility of the model and an improved survival rate.

From a scientific perspective, the significance of the PH-LHD debanding model does not solely lie in demonstrating the reversibility of the cardiovascular and pulmonary phenotype in heart failure, but more importantly, in the identification of molecular drivers that trigger and/or sustain reverse remodeling in pulmonary arteries as promising candidates for future therapeutic targeting.

Protocol

All procedures were performed following the "Guide for the Care and Use of Laboratory Animals" (Institute of Laboratory Animal Resources, 8th edition 2011) and approved by the local governmental animal care and use committee of the German State Office for Health and Social Affairs (Landesamt für Gesundheit und Soziales (LaGeSO), Berlin; protocol no. G0030/18). First, congestive heart failure was surgically induced in juvenile Sprague-Dawley rats ~100 g body weight (bw) (see Table of Materials) by placing a titanium clip with a 0.8 mm inner diameter on the ascending aorta (aortic banding, AoB) as described previously²⁹^,³⁰. At week 3 after AoB (Figure 1), debanding (Deb) surgery was performed to remove the clip from the aorta. The surgical procedures and validation of PH reversal in AoB rats performed are schematically depicted in Figure 1.

1. Surgical preparations

Sterilize the required surgical instruments (Figure 2) by autoclaving.
Inject rat with carprofen (5 mg/kg bw) (see Table of Materials) intraperitoneally (i.p.) for analgesia 30 min prior to surgery.
Anesthetize rat by i.p. injection of ketamine (87 mg/kg bw) and xylazine (13 mg/kg bw).
Remove the hair from the animal's neckline and chest using an electric shaver.
Apply a drop of eye ointment to protect the eyes during surgery.
Place the rat in a supine position on a sterilized surgical table. Carefully fix the animal's abdomen and limbs with adhesive tape.
NOTE: To maintain body temperature, place a 37 °C heating mat under the surgical table. Avoid heating of the head region to prevent drying of eyes.
Disinfect animal skin with povidone-iodine/iodophor solution. Note scars and sutures from the primary AoB surgery and drape the surgical field.
Ensure adequate depth of anesthesia by toe pinching.
NOTE: Depth of anesthesia needs to be controlled regularly during surgery.

2. Tracheotomy and mechanical ventilation

NOTE: Throughout the surgery, change gloves after handling non-sterile equipment.

With fine scissors (Figure 2A), make a 7-10 mm long cervical midline incision (Figure 3A).
With the help of a pair of blunt forceps (Figure 2B'), dissect the cervical soft tissue to expose the infrahyoid muscles. Split muscles in the midline to visualize the trachea. Cut and remove the suture from the primary AoB surgery.
Make ~2 mm trachea incision between two cartilaginous rings using angled Noyes spring scissors (Figure 2C,3B). Insert the tracheal cannula of outer diameter 2 mm (Figure 2D) into the trachea and secure it with a 4-0 silk suture (Figure 2E,3C).
Connect the tracheal cannula to a mechanical ventilator (see Table of Materials) while keeping dead space to a minimum (Figure 3D-E). Keep perioperative lung ventilation at a respiratory rate of 90 breaths/min at a tidal volume (Vt) of 8.5 mL/kg bw.

3. Aortic debanding

Make an ~20 mm long skin incision between the second and third ribs using fine scissors (Figure 3F).
With the help of smaller surgical scissors (Figure 2F), carefully spread muscles and cut them layer by layer (Figure 3G). Make a 10 mm lateral incision along the intercostal space between the second and third rib.
NOTE: The midsternal line needs to be carefully approached to avoid bleeding.
Use a rib spreader (Figure 2G) to expand the intercostal space between the second and third rib to create a surgical window (Figure 3H).
With the help of blunt forceps (Figure 2B,B'), carefully separate the thymus from the heart and conduit arteries to visualize the aorta with the clip (Figure 4A).
Hold the clip with the help of the forceps and carefully remove the connective tissue around the clip to expose it.
NOTE: Avoid holding or lifting the aorta with the forceps, as it may injure the aorta resulting in bleeding and a lethal outcome.
With the help of a needle holder (Figure 2H), open the clip (Figure 4B) and remove it from the thoracic cavity.
Before closing the chest, open up lung atelectasis, ensure adequate lung recruitment without overdistension, continue mechanical ventilation with a Vt of 9.5 mL/kg bw for another 10 min, and return to a Vt of 8.5 mL/kg bw to recruit the lungs and resolve a possible pneumothorax.
Close the deep muscles by a simple interrupted suture using 4-0 silk. Then connect the upper muscles and the skin with a simple continuous suture (Figure 5A,B).

4. Tracheal extubation

Disconnect the tracheal cannula from the ventilation machine. Attentively observe the rat until spontaneous breathing is re-established. If the animal fails to breathe spontaneously upon disconnection, reconnect the ventilator and continue ventilating for an additional 5 min. Then repeat the procedure.
After spontaneous breathing is re-established, remove the cannula from the trachea and clean the liquid around the trachea with sponge points (Figure 2I) (see Table of Materials).
Close the trachea with a simple suture using 6-0 prolene (Figure 2E' and Figure 5C). Then close infrahyoid muscles in a simple interrupted suture using 4-0 silk (Figure 5D), and connect the skin in a simple continuous suture (Figure 5E). Clean and disinfect the muscles and the skin during the process with povidone-iodine/iodophor solution.

5. Post-operative care

After completing the surgical procedure, carefully move the animal to a recovery cage with supplemental oxygen and an infra-red lamp to keep animals warm and sufficiently oxygenated during the recovery phase. Place the oxygen mask close to the rat's snout. Only keep one animal per recovery cage at any time.
After the animal wakes up, carefully move it to a regular cage supplied with water and food. For the next 12 h, control the health status of the operated animal in 2 h intervals.
After completing the surgical procedure, apply analgesia daily by i.p. injection of carprofen (5 mg/kg bw) for one week.
To avoid bacterial infection, administer amoxicillin (500 mg/L) in the drinking water for one week postoperatively.

Representative Results

First, successful aortic debanding was confirmed by transthoracic echocardiography performed before and after the debanding procedure in AoB animals (Figure 6). To this end, the aortic arch was assessed in parasternal long axis (PLAX) B-mode view. The position of the clip on the ascending aorta in AoB animals and its absence after the Deb surgery was visualized (Figure 6A,B). Next, aortic blood flow was evaluated by pulsed-wave Doppler imaging (Figure 6C-F). Peak blood flow velocity in AoB animals measured before and after the clip was 733.24 ± 17.39 mm/s and 5215.08 ± 48.05 mm/s (n = 8 animals), respectively (Figure 6C,E), demonstrating a steep gradient across the AoB site. After clip removal, peak blood flow velocity was 1093.79 ± 28.97 mm/s and 2578.73 ± 42.27 mm/s, respectively, at corresponding aortic locations, showing a marked attenuation of the gradient in line with functional debanding (Figure 6D,F). To probe for reversal of left heart failure by aortic debanding, the expression levels of brain natriuretic peptide (BNP), a clinical routine parameter for assessing heart disease³¹, were accessed in the LV myocardium. At weeks 3 and 5 after aortic banding, AoB animals showed a significantly increased production of BNP in comparison to sham-operated controls. In contrast, Deb rats at week 5 expressed BNP at levels comparable to sham animals, indicating the reversal of LV failure by aortic debanding (Figure 7A-C). In parallel, evaluation of LV function by transthoracic echocardiography revealed an increased LV ejection fraction and LV volume in Deb animals compared to AoB rats (Figure 7D-E). While LV ejection fraction in Deb animals was comparable to sham rats, LV volume in Deb rats failed to fully normalize to values seen in the sham group, indicating that reversal of LV function is incomplete.

To probe whether Deb animals may serve as a preclinical model to study reverse pulmonary vascular and right ventricular (RV) remodeling in PH-LHD, left ventricular systolic pressure (LVSP), and right ventricular systolic pressure (RVSP) was assessed using a microtip Millar catheter. In brief, rats were again anesthetized with ketamine (87 mg/kg bw) and xylazine (13 mg/kg bw), tracheotomized, and ventilated as described above. Cardiac catheterization was performed after median thoracotomy³² through the apex of (first) the left and (second) the right ventricle, respectively, as direct catheterization of the left ventricle via the vascular route is prevented by the aortic band in AoB animals. Following euthanasia by an overdose of ketamine/xylazine, the heart was excised, and ventricular hypertrophy was assessed as the weight of the left ventricle including septum (LV+S) or the right ventricle (RV) normalized to body weight (BW). In accordance with AoB rats as an established model for PH-LHD, AoB animals showed a significantly increased LVSP and RVSP and LV and RV hypertrophy compared to sham-operated animals at 3 weeks post-surgery (Figure 8A-F). Debanding (Deb) surgery performed at week 3 after AoB resulted in a significant reduction of both LVSP and LV hypertrophy in comparison to AoB animals without Deb at week 3 and week 5 post-AoB, demonstrating that normalization of LV hemodynamics following clip removal from the aorta reversed LV remodeling (Figure 8C,D). Compared to AoB rats at week 3 and week 5, Deb animals also showed a significant reduction in RVSP and RV/BW, demonstrating successful reversal of PH-LHD (Figure 8E,F). Notably, RVSP in Deb rats was comparable to values measured in sham animals, indicating a complete normalization of RV hemodynamics. In contrast, RV hypertrophy in Deb animals was only partially reversed with RV/BW, remaining significantly increased compared to sham controls (Figure 8E,F).

Figure 1: Schematic representation of the surgical procedures and validation of PH reversal in AoB rats. The schematic depicts the different experimental groups used in the present study to test whether debanding surgery reverses PH-LHD. Sham, sham-operated controls; AoB, aortic banding; Deb, debanding. Triangles mark the time point of surgical interventions: primary operation (sham or AoB; red) at week 0 and secondary operation (Deb; green) at week 3. Circles mark the end-point analyses at which PH-LHD was assessed by LV and RV pressures and hypertrophy measurements, respectively. Please click here to view a larger version of this figure.

Figure 2: Surgical instruments. (A) Fine scissors Tungsten carbide. (B) Moria Iris forceps and (B') Serrated Graefe forceps. The forceps' tips are shown enlarged. (C) Noyes spring scissors. (D) Tracheal cannula. (E, E') 4-0 and 6-0 sutures, respectively. (F) Fine scissors Tungsten carbide. (G) Rib spreader. (H) Mathieu needle holder. (I) Sponge points tissue. Please click here to view a larger version of this figure.

Figure 3: Tracheotomy and thoracotomy. Images illustrate the surgical steps for the tracheotomy. (A) Cervical midline incision. (B) Incision of the trachea between two cartilaginous rings. (C) Tracheal cannula inserted into the trachea and secured with a suture. (D) The tracheal cannula is connected to a mechanical ventilator. (E) Images illustrate the surgical steps for the thoracotomy. (F) Skin incision between the second and third ribs. (G) Cutting of muscles. (H) Creation of a thoracic surgical window by spreading the second and the third ribs. Please click here to view a larger version of this figure.

Figure 4: Visualization of the aortic-constricting clip in vivo and ex vivo. (A) The image shows the thoracic cavity of an AoB rat with a titanium clip placed on the ascending aorta. (B) The image shows closed and opened clip ex vivo. Asterisk marks the part of the clip that the needle holder compresses in vivo to open the clip. Please click here to view a larger version of this figure.

Figure 5: Wound closure. Images illustrate the closing of the upper thoracic muscles (A) and the skin (B) with a simple continuous suture. The trachea (C) and the infrahyoid muscles (D) are closed by a simple suture and the skin on the neck (E) by a simple continuous suture. Please click here to view a larger version of this figure.

Figure 6: Aortic blood flow prior to and after debanding surgery. (A-B) Visualization of the ascending aorta in a rat with aortic banding (AoB, left) and a rat after debanding surgery (Deb, right) by transthoracic echocardiography. The arrow shows the titanium clip on the aorta in (A) absent in (B). (C,D) Pulsed-wave Doppler echocardiographic images show blood flow before the clip in an AoB rat (C) and blood flow in the corresponding aortic segment in a Deb rat (D) taken one day prior and one day after the aortic debanding surgery, respectively. (E,F) Analogously, images show blood flow in the aortic segment after the clip in an AoB rat (E) and in the corresponding aortic segment in a Deb rat (F) taken one day prior and one day after the aortic debanding surgery, respectively. Turquois vertical lines illustrate measurements of peak aortic flow velocity. Please click here to view a larger version of this figure.

Figure 7: Normalization of left ventricular function by aortic debanding. (A) Representative Western blots show protein levels of BNP and with GAPDH as loading control in left ventricles (LV) of AoB rats at week 3 after aortic banding (n = 5) and in corresponding sham controls (n = 5). (B) Representative Western blots show BNP and GAPDH in left ventricles (LV) of AoB rats at week 5 after aortic banding (n = 4), in Deb rats at week 5 (n = 5), and in sham controls at the corresponding time after primary surgery (n = 4). (C) Box and whisker plots show quantification of BNP expression normalized to GAPDH and sham control at the corresponding time after primary surgery. Boxes show median, 25, and 75 percentiles, respectively; whiskers indicate the minimum and maximum values. For statistical analyses, Student's t-test³³ was used. *p-value < 0.05. (D) Bar graphs (mean ± standard deviation) show LV ejection fraction and volume in sham (n = 4), AoB (n = 9), and Deb (n = 7) animals at week 5 as measured by echocardiography from M- and B-mode images. (E) Representative M-mode echocardiographic images show changes in LV fractional shortening in sham, AoB, and Deb animals at week 5. For statistical analyses Mann-Whitney U test³³ was used. *p-value < 0.05. Please click here to view a larger version of this figure.

Figure 8: Ventricular hemodynamics are normalized, and cardiac hypertrophy is reversed by aortic debanding. (A) Representative measurements of left ventricular (LV) and right ventricular (RV) blood pressure in a rat 3 weeks after aortic banding (AoB) as compared to corresponding sham control. (B) Representative images show cardiac hypertrophy in an AoB rat 3 weeks after aortic banding compared to sham control. (C-F) Box and whisker plots show left ventricular systolic pressure (LVSP; C), LV hypertrophy ([LV+S]/BW; D), right ventricular systolic pressure (RVSP; E), and RV hypertrophy (RV/BW; F) in sham and AoB animals at 3 and 5-weeks post-surgery, and normalized parameters (in comparison to 3- and 5-week AoB groups) in Deb rats. Boxes show median, 25, and 75 percentiles, respectively; whiskers indicate the minimum and maximum values. n = 9-12 animals were analyzed for hemodynamic measurements, and heart weight was measured in n = 7-12 rats per group. For statistical analyses Mann-Whitney U test was used. *p-value < 0.05. Please click here to view a larger version of this figure.

Discussion

Here, a detailed surgical technique for aortic debanding in a rat AoB model is reported that can be utilized to investigate the reversibility of PH-LHD and the cellular and molecular mechanisms that drive reverse remodeling in the pulmonary vasculature and the RV. Three weeks of aortic constriction in juvenile rats results in PH-LHD evident as increased LV pressures, LV hypertrophy, and concomitantly increased RV pressures and RV hypertrophy. Aortic debanding at week 3 post-AoB was able to unload the LV and fully reverse LV hypertrophy within two weeks after Deb. In parallel, aortic debanding also caused a full normalization of RV pressures and a partial reversal of RV hypertrophy.

The present model thus mimics critical aspects of the clinical scenario where mechanical unloading of the LV by an implantable non-pulsatile LVAD with continuous flow properties has previously been found to reverse PH in patients with heart failure³⁴^,³⁵. In a retrospective analysis, LVAD support was shown to reduce pulmonary artery pressure to similar degrees in heart failure patients with either reversible or fixed PH, with fixed PH defined as mean pulmonary arterial pressure >25 mm Hg, pulmonary vascular resistance >2.5 Wood Unit and a mean pressure transpulmonary gradient >12 mm Hg despite pharmacological treatment³⁵. Importantly, these findings³⁴^,³⁵ provide indirect evidence that left ventricular unloading not only decreases passive pulmonary congestion and secondary changes in lung vascular tone but triggers reverse remodeling of the pulmonary vasculature by “physiological” mechanisms, i.e., by adaptation to altered hemodynamics. In-depth, multi-omics analyses of the cellular and molecular processes that drive reverse remodeling in the pulmonary vasculature could open up new avenues for identifying novel therapeutic options for treating PH in heart failure patients and potentially also in other forms of PH including PAH. The present model of debanding in AoB rats provides a unique possibility for such analyses as complete normalization of RVSP confirms effective reversal of PH, thus allowing for mechanistic studies to identify pathways with the ability to restore homeostatic processes in the diseased pulmonary vasculature.

With a similar rationale, the present model may further be utilized to study intra- and intercellular processes that drive reverse remodeling of the RV. RV function has recently been recognized as a significant predictor of prognosis for morbidity and mortality in cardiovascular diseases. Yet no therapies have been clinically approved to improve RV function³⁶. As such, the ability to study reverse remodeling processes in the RV myocardium in an animal model provides a unique opportunity to address a significant knowledge gap and a critical medical need.

The success of the technically challenging aortic debanding procedure in AoB rats depends on surgical skills and precise perioperative strategies. Outlined in the following are critical surgical procedure steps that may cause perioperative lethality by excessive bleeding (1-5) or insufficient respiration (6) and recommendations on avoiding these complications.

(1) During a thoracotomy, the midsternal line needs to be approached carefully with scissors to avoid injury to the internal mammary artery. (2) To visualize the heart and the conduit arteries, the thymus should be mobilized and carefully relocated in the cranial direction. In the debanding surgery, the thymus tissue is often connected to the heart and arteries via post-operative adhesions from the original AoB surgery. These adhesions should be carefully separated with a pair of blunt forceps to avoid injury to the cardiovascular structures. (3) In the debanding surgery, the aorta with the clip is frequently embedded in connective tissue. This connective tissue must be gently dissected with blunt forceps to visualize the clip. Here, the transthoracic echocardiography performed before the surgery is a helpful preparation step, allowing to identify whether the clip is located close to the aortic root, in the middle of the ascending aorta, or close to the brachiocephalic artery. This knowledge saves precious time for clip allocation during the surgery. (4) The orientation of the clip is a critical step that must be considered carefully during the initial aortic banding surgery. To facilitate optimal assessment and rapid opening of the clip during aortic debanding, the part that needs to be compressed by the needle holder (Figure 4B) should be oriented ventrally. Clip reorientation during debanding surgery is feasible, although at the risk of injury to the aorta. For clip reorientation, clips need to be held by forceps while surrounding connective tissue is carefully removed, then the clip should be mobilized and turned. Holding the aorta with the forceps is to be avoided. (5) For debanding, the clip should be held by a forceps with one hand and opened with a needle holder with the other hand. The aorta need not be lifted ventrally. (6) After completing the debanding procedure, extubated PH-LHD rats are at considerable risk of respiratory failure, with animals commonly dying within 10-20 min after surgery while still under the anesthesia. Atelectasis of the left lung is the most common cause of death in this period, and prolonged mechanical ventilation before chest closure helps recruit the lung and warrant sufficient respiration after surgery.

We also suggest that compared to endotracheal intubation as performed in previous studies²⁶^,²⁷, tracheostomy provides better control of appropriate ventilation during surgical procedures, which is specifically relevant during aortic debanding. This notion is based on the following rationale: (1) Tracheostomy, routinely performed in our lab for perioperative lung ventilation, is a straightforward and safe technique with no perioperative or post-operative complications. (2) Tracheostomy eliminates the risk of esophageal intubation or tracheal injury; it enables precise positioning and fixation of the tracheal cannula and constant visual control of the cannula during all steps of the surgical procedure. (3) At the time of aortic debanding, AoB animals are already in heart failure and are more sensitive to additional stress; as a result, the potential risks that come with endotracheal intubation may add to increased lethality. (4) Finally, when the operated animal is weaned from the ventilator but fails to develop spontaneous breathing, a tracheostomy allows for rapid reintubation and reconnection with the ventilator, thus potentially saving lives due to the ability for prolonged post-operative ventilation.

The present study reports an aortic debanding technique performed 3 weeks after initial aortic banding in rats. For studies aiming to compare reverse remodeling of the pulmonary vasculature and the RV at different PH stages, the described procedures may also be performed at later time points after aortic banding. Yet, caution is warranted as scar and connective tissue surrounding the aorta will likely become more abundant with time, further complicating the procedure and necessitating additional troubleshooting and refinement. At the same time, the basic principles of the reported protocol still apply.

Disclosures

The authors have nothing to disclose.

Acknowledgements

This research was supported by grants of the DZHK (German Centre for Cardiovascular Research) to CK and WMK, the BMBF (German Ministry of Education and Research) to CK in the framework of VasBio, and to WMK in the framework of VasBio, SYMPATH, and PROVID, and the German Research Foundation (DFG) to WMK (SFB-TR84 A2, SFB-TR84 C9, SFB 1449 B1, SFB 1470 A4, KU1218/9-1, and KU1218/11-1).

Materials

Amoxicillin	Ratiopharm	PC: 04150075615985	Antibiotic
Anti-BNP antibody	Abcam	ab239510	Western Blotting
Aquasonic 100 Ultrasound gel	Parker Laboratories	BT-025-0037L	Echocardiography consumables
Bepanthen	Bayer	6029009.00.00	Eye ointment eye ointment
Carprosol (Carprofen)	CP-Pharma	401808.00.00	Analgesic
Clip holder	Weck stainless USA	523140S	Surgical instruments
Fine scissors Tungsten carbide	Fine Science Tools	14568-12	Surgical scissors
Fine scissors Tungsten carbide	Fine Science Tools	14568-09	Surgical scissors
High-resolution imaging system	FUJIFILM VisualSonics, Amsterdam, Netherlands	VeVo 3100	Echocardiography machine. Images were acquired with pulse-wave Doppler mode, M-mode and B-mode
Isoflurane	CP-Pharma	400806.00.00	Anesthetic
Ketamine	CP-Pharma	401650.00.00	Anesthetic
Mathieu needle holder	Fine Science Tools	12010-14	Surgical instruments
Mechanical ventilator (Rodent ventilator)	UGO Basile S.R.L.	7025	Volume controlled respirator
Metal clip	Hemoclip	523735	Surgical consumables
Microscope	Leica	M651	Manual surgical microscope for microsurgical procedures
Millar Mikro-Tip pressure catheters	ADInstruments	SPR-671	Hemodynamics assessment
Moria Iris forceps	Fine Science Tools	11373-12	Surgical forceps
Noyes spring scissors	Fine Science Tools	15013-12	Surgical scissors
Povidone iodine/iodophor solution	B/Braun	16332M01	Disinfection
PowerLab	ADInstruments	4_35	Hemodynamics assessment
Prolene Suture, 4-0	Ethicon	EH7830	Surgical consumables
Rib spreader (Alm selfretaining retractor blunt, 70 mm, 2 3/4″)	Austos	AE-BV010R	Surgical instruments
Serrated Graefe forceps	Fine Science Tools	11052-10	Surgical forceps
Silk Suture, 4-0	Ethicon	K871	Surgical consumables
Skin disinfiction solution (colored)	B/Braun	19412M07	Disinfection
Spectra 360 Elektrode gel	Parker Laboratories	TB-250-0241H	Echocardiography consumables
Sponge points tissue	Sugi	REF 30601	Surgical consumables
Sprague-Dawley rat	Janvier Labs, Le Genest-Saint-Isle, France		Study animals
Tracheal cannula			Outer diameter 2 mm
Xylazin	CP-Pharma	401510.00.00	Anesthetic

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Sang, P., Kucherenko, M. M., Yao, J., Li, Q., Simmons, S., Kuebler, W. M., Knosalla, C. A Model of Reverse Vascular Remodeling in Pulmonary Hypertension Due to Left Heart Disease by Aortic Debanding in Rats. J. Vis. Exp. (181), e63502, doi:10.3791/63502 (2022).