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

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 veins^3,4. 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 failure^1,2,5,6. 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 transplantation^7,8,9. 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 devices^12,13,14. 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 rats^4,15,16. 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-LHD^{4,15,20,21,22,23,24,25}.

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 hemodynamics^27,28, 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 needle^26,27, 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 previously^29,30. 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

1. Sterilize the required surgical instruments (Figure 2) by autoclaving.
2. Inject rat with carprofen (5 mg/kg bw) (see Table of Materials) intraperitoneally (i.p.) for analgesia 30 min prior to surgery.
3. Anesthetize rat by i.p. injection of ketamine (87 mg/kg bw) and xylazine (13 mg/kg bw).
4. Remove the hair from the animal's neckline and chest using an electric shaver.
5. Apply a drop of eye ointment to protect the eyes during surgery.
6. 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.
7. Disinfect animal skin with povidone-iodine/iodophor solution. Note scars and sutures from the primary AoB surgery and drape the surgical field.
8. 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.

1. With fine scissors (Figure 2A), make a 7-10 mm long cervical midline incision (Figure 3A).
2. 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.
3. 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).
4. 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

1. Make an ~20 mm long skin incision between the second and third ribs using fine scissors (Figure 3F).
2. 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.
3. Use a rib spreader (Figure 2G) to expand the intercostal space between the second and third rib to create a surgical window (Figure 3H).
4. 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).
5. 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.
6. With the help of a needle holder (Figure 2H), open the clip (Figure 4B) and remove it from the thoracic cavity.
7. 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.
8. 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

1. 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.
2. 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).
3. 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

1. 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.
2. 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.
3. After completing the surgical procedure, apply analgesia daily by i.p. injection of carprofen (5 mg/kg bw) for one week.
4. To avoid bacterial infection, administer amoxicillin (500 mg/L) in the drinking water for one week postoperatively.

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 (...

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
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
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