A Murine Model of Pressure Overload-Induced Right Ventricular Hypertrophy and Failure by Pulmonary Trunk Banding

Abstract

Right ventricular (RV) failure caused by pressure overload is strongly associated with morbidity and mortality in a number of cardiovascular and pulmonary diseases. The pathogenesis of RV failure is complex and remains inadequately understood. To identify new therapeutic strategies for the treatment of RV failure, robust and reproducible animal models are essential. Models of pulmonary trunk banding (PTB) have gained popularity, as RV function can be assessed independently of changes in the pulmonary vasculature.

In this paper, we present a murine model of RV pressure overload induced by PTB in 5-week-old mice. The model can be used to induce different degrees of RV pathology, ranging from mild RV hypertrophy to decompensated RV failure. Detailed protocols for intubation, PTB surgery, and phenotyping by echocardiography are included in the paper. Furthermore, instructions for customizing instruments for intubation and PTB surgery are given, enabling fast and inexpensive reproduction of the PTB model.

Titanium ligating clips were used to constrict the pulmonary trunk, ensuring a highly reproducible and operator-independent degree of pulmonary trunk constriction. The severity of PTB was graded by using different inner ligating clip diameters (mild: 450 µm and severe: 250 µm). This resulted in RV pathology ranging from hypertrophy with preserved RV function to decompensated RV failure with reduced cardiac output and extracardiac manifestations. RV function was assessed by echocardiography at 1 week and 3 weeks after surgery. Examples of echocardiographic images and results are presented here. Furthermore, results from right heart catheterization and histological analyses of cardiac tissue are shown.

Introduction

Right ventricular (RV) failure is a clinical syndrome with symptoms of heart failure and signs of systemic congestion resulting from RV dysfunction¹. RV dysfunction is strongly associated with morbidity and mortality in a number of cardiovascular and pulmonary diseases². The etiology of RV dysfunction is complex, and its underlying signaling pathways and regulation remain inadequately elucidated.

Observations from current therapies show that improved RV function correlates closely to afterload reduction, suggesting pulmonary vasculature as the primary treatment target³. This indicates that current therapies only have a minimal direct effect on RV function, which can deteriorate even after the improvement of pulmonary vascular resistance³. Further research into improving RV function independently of afterload reduction is thus highly needed.

Robust and reproducible animal models are essential in the search for new therapeutic agents. In most models of chronic RV failure, the underlying cause is pulmonary hypertension induced by structural alteration of the pulmonary vasculature^4,5,6. Well-characterized models include the chronic hypoxia model^7,8, the Sugen-hypoxia model^9,10,11, and the monocrotaline model^12,13. Because the RV failure is secondary to pulmonary hypertension in these models, it is impossible to differentiate the effects of interventions on the pulmonary vasculature from the direct effects on the RV⁶.

To study the RV independently from the pulmonary vasculature, the pulmonary trunk banding (PTB) model has gained popularity and has been described in several animal species, including mice, rats, rabbits, dogs, sheep, and pigs^{6,14,15,16,17,18,19,20,21,22,23,24,25,26,27}. In PTB models, constriction of the pulmonary trunk is achieved surgically, causing an increase in RV pressure⁶. Different approaches to the application of PTB exist, including constriction of the vessel with a ligature or with a metal ligating clip^18,28. In models using ligatures, the pulmonary trunk is tied to a needle, and the needle is retracted, leaving the ligature in place. This results in a constriction of the vessel that depends on the needle size and the tension of the knot^18,29. In models employing metal ligating clips, the degree of pulmonary trunk constriction may be more reproducible. Modified ligating clip appliers are used to close the ligating clips to a predefined and constant diameter. This makes the method operator-independent and reduces PTB-related variability in the disease phenotype^15,27,28.

Murine PTB models have been shown to induce RV hypertrophy and failure^18,28. One major challenge when using the PTB model is choosing the appropriate PTB diameter to achieve the desired degree of RV pathology. This is especially challenging when attempting to model decompensated RV failure. For this, the constriction needs to be tight enough to induce chronic RV failure without leading to acute RV failure and death shortly after surgery⁶. One approach to solving this challenge is to use weanlings or juvenile animals^6,15. A PTB model has been used successfully to study different stages of RV failure using Wistar rat weanlings^15,30. To achieve this, juvenile rats with remaining growth potential underwent PTB surgery with the application of titanium ligating clips. When the rats grew, the pulmonary stenosis gradually became more severe and resulted in RV hypertrophy or chronic RV failure, depending on the severity of PTB^15,30. Inspired by this model, we hypothesized that different stages of RV pathology could be produced in a murine PTB model using juvenile mice. Studying a broad spectrum of RV pathology from mild to severe disease may help elucidate our understanding of disease progression and the transition from RV hypertrophy to RV failure.

Here, we present a murine model of RV pressure overload induced by PTB in juvenile mice. With this model, different degrees of RV pathology can be produced, ranging from RV hypertrophy to decompensated RV failure. This study includes detailed protocols for intubation, PTB surgery, and phenotyping by echocardiography.

Protocol

The study was approved by the Danish Animal Experiments Inspectorate (authorization number: 2021-15-0201-00928) and was performed in accordance with the national laboratory animal legislation. This study used 5-week-old male C57BL/6N mice.

1. Customization of instruments for intubation and surgery (Figure 1)

NOTE: This section details the most important steps in the preparation of custom-made instruments for intubation and PTB surgery from inexpensive and readily available materials.

1. Prepare the endotracheal tube (Figure 1A).
   Common intravenous (IV) catheters can be used for intubation and ventilation of mice.
   1. Choose the appropriate size of intravascular catheter corresponding to the size of the mice. A 22G catheter is recommended for 5-week-old mice (17-20 g) and a 23G catheter for larger mice (>20 g).
   2. Pull out the needle and detach the locking mechanism. Insert the needle back into the catheter and cut the tip of the needle at a 45° angle, approximately 2 mm from the tip of the catheter.
   3. Use sandpaper to blunt the tip to prevent injury to the vocal cords of the mice.
   4. Cut off one wing of the catheter for a better view during intubation.
2. Prepare the thoracic retractor (Figure 1B).
   1. Use a needle holder to bend an approximately 10 cm piece of flexible metal wire in the middle at a 30° angle.
   2. Use the needle holder to carefully create atraumatic hooks of 5 mm width on both ends of the wire.
3. Prepare the intubation stand (Figure 1C).
   NOTE: The use of an intubation stand enables continuous delivery of anesthesia via a nasal tube during intubation. This allows for controlled and safe intubation under visual guidance, which reduces the risk of injury to the vocal cords and trachea as well as the risk of misplacement of the tube in the esophagus. Any metal or plastic frame can be used as a frame for the intubation stand. A slightly bent kitchen role holder was used in this study.
   1. Cut a 3 cm piece of rubber tubing that fits the snout of a mouse and connect it to an IV catheter valve. Inhalant anesthesia can be connected via the valve prior to intubation.
   2. Make a loop using a 1-0 braided suture approximately 5 mm from the opening of the tube. This will be used to secure the murine snout in the tube.
   3. Cut a hole near the top of a flexible plastic sheet and place the tube in the hole. The sheet is used to support the mouse in the intubation stand.
   4. Use tape to hold the individual parts together as shown in Figure 1C.
4. Prepare the guidance cannula (Figure 1D, E).
   1. Pull a 6-0 monofilament suture through a blunt 27G cannula and tie a knot on the suture. Use this knot later to grasp the suture during PTB surgery.
   2. Use a needle holder to bend the tip of the cannula to an 80° angle.
5. Prepare the ligating clip applier.
   ​NOTE: A ligating clip applier with angled jaws is modified with an adjustable stop mechanism (Figure 1F), which stops the compression of the ligating clip when the jaws are at a precisely predetermined distance from each other. The modified ligating clip applier is used for the application of titanium ligating clips on the pulmonary trunk.
   1. Fix a custom-cut piece of brass to the handles of the ligating clip applier with two screws. Mount an adjustable screw in the center (Figure 1F, white arrow) that determines the exact distance between the compressed handles of the ligating clip applier, which corresponds to a precise distance between the jaws.

Figure 1: Instruments for intubation and PTB surgery. (A) Endotracheal tube made from an IV catheter. (B) Thoracic retractor. (C) Intubation stand and mouse placed in intubation stand receiving anesthesia on a nasal tube. (D) Surgical instruments and modified ligating clip applier used for PTB surgery. (E) Guidance cannula. (F) Custom-made adjustable stop-mechanism. Please click here to view a larger version of this figure.

2. Adjustment of the ligating clip applier

1. Choose the internal diameter of the ligating clip based on the desired severity of RV failure. For 5-week-old male C57BL/6N mice (17-20 g), use a clip diameter of 250 µm for severe and 450 µm for mild RV pressure overload. Use intermediate clip diameters to induce moderate pressure overload.
2. Use metal wire or needles to adjust the ligating clip applier. Ensure that the diameter of the wire corresponds to the desired clip diameter. 
3. Load the clip applier with a ligating clip and place the adjustment wire in the center of the ligating clip. While compressing the clip applier, turn the screw until the jaws of the clip fit tightly around the wire. Ensure the clip stays in place on the adjustment wire once the clip applier is released.
4. With the clip applier adjusted, place another ligating clip on the adjustment wire to validate the set diameter.

3. Preparations for surgery

1. Induce anesthesia in an induction chamber using 7% sevoflurane in 0.6 L/min 100% oxygen. Confirm sufficient anesthesia by toe pinch prior to intubation.
   NOTE: Isoflurane may also be used. Be aware that different concentrations should be used.
2. Intubate the mouse with a 22G IV catheter. Perform intubation under visual guidance, using a surgical microscope and an intubation stand, allowing for proper alignment for visualization of the vocal cords and continuous delivery of inhalant anesthetics on a nasal tube (Figure 1C).
3. Ventilate the mouse at 175 strokes/min and a tidal volume of 300 µL/stroke.
   NOTE: Tidal volumes of 8-10 µL/g are also recommended, and the ideal tidal volume depends on possible leakage and dead space in the ventilation system.
4. Place the intubated mouse on a covered heating pad (37 °C) and apply moisturizing ointment to the eyes of the mouse.
5. Maintain anesthesia (3.5% sevoflurane in 0.6 L/min 100% oxygen) and administer 0.1 mg/kg buprenorphine and 5 mg/kg carprofen subcutaneously for perioperative analgesia. Remove all hair from the chest using depilatory cream and disinfect the skin with disinfectant wipes.

4. PTB surgery

1. Make a 10 mm incision in the skin above the second intercostal space from the sternal angle to the left anterior axillary line. Split the major and minor pectoral muscles by blunt dissection.
2. Cut the intercostal muscles in the second intercostal space and bluntly dissect the thymus to expose the heart, pulmonary trunk, and aorta. Place a thoracic retractor in the intercostal space to keep the operating field accessible.
   NOTE: Great caution must be used when cutting the intercostal muscles, as the left internal mammary artery runs just 1-2 mm laterally of the sternum. Injury to this artery may result in significant blood loss.
3. Separate the pulmonary trunk from the ascending aorta by bluntly removing the connective tissue between the vessels using microscopic forceps. Improve the exposure of the pulmonary trunk further by rotating the lower body of the mouse (left leg over right leg).
4. Pass the guidance cannula through the transverse pericardial sinus posteriorly of the pulmonary trunk. Use forceps to grasp the knot on the tip of the guidance cannula and pull the suture through the guidance cannula. Carefully remove the guidance cannula while the suture stays in place around the pulmonary trunk.
5. Load the ligating clip applier and use the suture to guide the pulmonary trunk into the jaws of the ligating clip and compress the clip. Release the suture immediately after placement of the clip and observe the filling of the pulmonary trunk.
   NOTE: Bradycardia may be observed in the first few seconds after application of the ligating clip.
6. Place a 6-0 monofilament absorbable suture around the second and third costae and close the intercostal space. Evacuate as much air as possible from the thoracic cavity by applying gentle pressure on the chest while tightening the suture.
7. Lastly, suture the skin with a 6-0 monofilament absorbable suture.
8. Perform the same procedure, except for placing the ligating clip (step 4.5), during sham surgery.

5. Echocardiography

1. After induction of anesthesia in an induction chamber (1-2 min, 6% sevoflurane in 100% oxygen), maintain anesthesia using a nasal tube (3.5% sevoflurane in 100% oxygen). Remove all hair from the chest and abdomen using depilatory cream and place the mouse on a heating pad. Apply moisturizing ointment to the eyes and ultrasound gel to the mouse's chest.
   NOTE: Adjust 2D gain, focus depth, and image depth to improve image quality in all echocardiographic measurements.
2. Adjust the ultrasonic probe to find the parasternal long axis (PLAX) view (Figure 2). In PLAX, measure the pulmonary trunk inner diameter and velocity time integral (VTI) in the pulmonary trunk.
   1. Select B-mode and carefully move the heating pad in the x-, y- and z-axis to identify the pulmonary trunk in the center of the image. Use color to identify the largest pulmonary trunk diameter. Use cine store to capture a sequence for measurement of the pulmonary trunk diameter.
   2. Select color and pulse wave (PW) Doppler and place the cursor in the center of the pulmonary trunk. Adjust the PW angle until the dotted lines are parallel to the blood flow in the vessel.
   3. Press cine store to measure VTI. Place the cursor near both walls of the pulmonary trunk and press cine store again to acquire flow near the vessel wall.
      NOTE: Right ventricular (RV) inner diameter and RV free wall thickness may also be assessed in the PLAX view.
3. Find the parasternal short axis (PSAX) view (Figure 3) to measure left ventricular (LV) inner diameters, which can be used to assess septal bulging (D-configuration).
   1. Select B-mode and turn the probe 90° anti-clockwise. Angle the probe 20-30 degrees laterally to avoid shadowing of the RV by the sternum, and tilt the probe 20-30 degrees cranially until the left ventricle is as round as possible. Then, slide the probe in the craniocaudal direction to identify the level of the papillary muscles with the largest ventricular diameters and press cine store.
      NOTE: PSAX at the midpapillary level may also be used to assess RV fractional area change. PSAX at the level of the aortic valve may be used to measure RV fractional shortening or RV free wall thickness.
4. Use the apical 4-chamber (A4CH) view (Figure 4) to measure the tricuspid annular plane systolic excursion (TAPSE) and assess for tricuspid regurgitation.
   1. Position the probe as shown in Figure 4. Once the heart is identified, make small adjustments to the probe using only the wrist and fingers to identify all four chambers of the heart and the tricuspid valve.
      1. Identifying and maintaining a good A4CH view is challenging. Allow the probe-operating hand to rest on the heating pad for stability. Slide, tilt, and turn the probe slightly until the appropriate image is found.
      2. Only move the probe in one dimension at a time: E.g., slide craniocaudal to find the heart, then tilt the probe to identify all four chambers and finally turn the probe until all four chambers and the tricuspid valve are in the frame. If necessary, repeat all three steps several times before obtaining the ideal image.
   2. Once the tricuspid valve is identified, select M-mode and place the dotted line in the tricuspid annulus of the free wall. Press cine store to save the measurements.
   3. Select color to assess the tricuspid valve for regurgitation. If regurgitation is present, a jet of retrograde flow from the RV to the right atrium (RA) will be seen in systole (Figure 5).
      NOTE: In the A4CH view, tissue Doppler in the RV free wall and RV inflow velocity may also be measured.

Figure 2: Parasternal long axis view (PLAX). (A-D) Positioning of the ultrasonic probe. (E, F) The normal murine heart in PLAX. (G, H) RV dilation and hypertrophy after PTB. Abbreviations: LV: left ventricle, RV: right ventricle, PV: pulmonary valve, PT: pulmonary trunk, Ao: aorta. Please click here to view a larger version of this figure.

Figure 3: Parasternal short axis view (PSAX). (A-D) Positioning of the ultrasonic probe. (E, F) The normal murine heart in PSAX. (G, H) PSAX after PTB. Abbreviations: LV: left ventricle, RV: right ventricle, PM: papillary muscle. Please click here to view a larger version of this figure.

Figure 4: Apical 4-chamber view (A4CH). (A-D) Positioning of the ultrasonic probe. (E, F) The normal murine heart in the A4CH view. (G, H) RV and RA dilatation after PTB. Abbreviations: LV: left ventricle, RV: right ventricle, RA: right atrium, LA: left atrium. Please click here to view a larger version of this figure.

Figure 5: Tricuspid regurgitation visualized by color Doppler in the A4CH-view. (A) In diastole, flow from the RA to the RV is observed (arrow). (B) During systole, a thin jet of flow from the RV to the RA is visible (arrow). Abbreviations: LV: left ventricle, RV: right ventricle, RA: right atrium, LA: left atrium. Please click here to view a larger version of this figure.

6. Data analyses

1. Measure the pulmonary trunk diameter in three cardiac cycles in PLAX and use the mean pulmonary trunk diameter for further data analyses. Measure VTI in three cardiac cycles in PLAX for each of the three stored cine loops (in the center of the pulmonary trunk and near the walls of the vessel). Use mean VTI of all VTI measurements for further analysis. Use the following formula to calculate CO:
   
   : radius of the pulmonary trunk, HR: heart rate
2. Measure LVEI in PSAX at midpapillary level. Use a measuring tool to measure the largest LV inner diameter (LVid₁) from mid-septum to the free wall. Then, measure the LV inner diameter orthogonal to the first measurement (LVid₂). Repeat these measurements in three cardiac cycles and calculate LVEI by using the mean LV inner diameters and the following formula:
   
3. Measure TAPSE in three cardiac cycles in the A4CH view and use mean TAPSE for further data analyses.

7. Right heart catheterization

1. Measure the right ventricular (RV) pressure and volume by right heart catheterization with a 1.4F micro-tip catheter 3 weeks after PTB surgery.
2. Anesthetize and intubate the mouse as described in steps 3.1-3.4. Place the mouse on a covered heating pad (37 °C) and maintain anesthesia (3.5% sevoflurane in 0.6 L/min 100% oxygen).
3. Administer 2000 IU of Heparin (intramuscular [i.m.]) and 0.5 mL of NaCl (subcutaneous [s.c.]).
4. Using surgical scissors, cut the abdominal wall just caudally to the xiphoid process and gain access to the thoracic cavity by carefully cutting the diaphragm along its incision in the thoracic wall. Cut the diaphragm and costae until sufficient access to the heart has been obtained.
5. Place a ligature around the inferior vena cava. Use this for occluding the vessel to reduce preload for the recording of pressure-volume measurements later in the protocol.
6. Use a 26G needle to carefully poke a small hole in the RV. Ensure this is as near to the apex as possible and the needle does not penetrate the ventricle entirely but merely acts as a guide for insertion of the conductance catheter. If bleeding occurs, apply gentle pressure with a small cotton swab to minimize blood loss.
7. Identify the small hole in the ventricular wall and insert the catheter by penetrating the tissue.
   NOTE: When inserting the catheter, be careful not to damage the inner ventricular wall.
8. Changes in right ventricular pressure (RVP) are often observed for several minutes after insertion of the catheter. Wait until RVP stabilizes to obtain representative steady-state measurements.
9. To obtain pressure-volume loops, use the ligature previously placed around the inferior vena cava. Carefully pull the ligature to occlude the vessel, thereby gradually reducing preload.
10. Once representative pressure-volume loops have been recorded, extract the catheter, and euthanize the mouse by excision of the heart. At this time, collect blood and tissue samples for further analysis.

Representative Results

C57BL/6N mice (male, 5-week old, 17-20 g) were randomized to either severe PTB (sPTB, 250 µm, n = 12), mild PTB (mPTB, 450 µm, n = 9), or sham surgery (sham, n = 15). Evaluation of cardiac function was performed by echocardiography 1 week and 3 weeks after surgery. Right heart catheterization with subsequent euthanasia was performed 3 weeks post-surgery. Organs were weighed, and cardiac tissue was prepared for histological analyses.

Echocardiography 1 week after surgery revealed signs of elevated RV pressure and RV dysfunction in sPTB compared with mPTB and sham groups. RV dysfunction was evident by decreased cardiac output (CO) and TAPSE (Figure 6A,B). Decreased left ventricular eccentricity index (LVEI) indicated increased RV pressure (Figure 6C). Therefore, reversal interventions may be investigated already after 1 week in severe PTB models. The mPTB group showed echocardiographic signs of elevated RV pressure compared with sham (decreased LVEI) after 1 week but no signs of RV dysfunction as assessed by echocardiography (Figure 6).

Figure 6: Echocardiographic measurements 1 week after surgery. (A) Cardiac output (CO). (B) Tricuspid annular plane systolic excursion (TAPSE). (C) Left ventricular eccentricity index (LVEI). Abbreviations: mPTB: mild pulmonary trunk banding (PTB); sPTB: severe PTB. Data are presented by scatterplots with means ± SD. *p < 0.05, **p < 0.001. Please click here to view a larger version of this figure.

After 3 weeks, the effects of the PTB surgery were accentuated in sPTB with a further reduction in cardiac output and a consistent decrease in TAPSE compared with sham (Figure 7A,B). No difference in echocardiographic parameters was observed between mPTB and sham groups after 3 weeks (Figure 7A,B).

Right heart catheterization revealed increased RV systolic pressure (RVSP) in both PTB groups compared with sham 3 weeks after surgery (Figure 7C). RV diastolic function was affected in the sPTB group, indicated by an increase in RV end-diastolic elastance (Eed), a load-independent measure of RV stiffness (Figure 7D). RV contractility, as assessed by end-systolic elastance (Ees), was also increased in the sPTB group compared with the sham, whereas the mPTB group showed a non-significant increase (Figure 7E).

Anatomical measurements revealed increasing RV hypertrophy with increased severity of PTB expressed by a stepwise increase in RV weight to body weight (RV/BW) ratio (Figure 7F). RVSP was linearly proportional to the degree of RV hypertrophy (Figure 7G). RV cardiomyocyte cross-sectional area (CSA) confirmed a 40% increase in RV hypertrophy in sPTB compared with sham, while there was a non-significant increase in mPTB compared with sham (Figure 8A-D). The degree of fibrosis in the RV and right atrium (RA) was seven-fold higher in sPTB compared with sham. No increase in RV fibrosis was observed in mPTB (Figure 8E-H).

Tricuspid regurgitation (TR) was observed in 64% in the sPTB group after 1 week and in 91% at 3 weeks, whereas no TR was found in the sham and mPTB groups (data not shown). Varying degrees of liver discoloration, a sign of extracardiac decompensation, were seen in the sPTB group (Figure 9). No discoloration was seen in the mPTB and sham groups.

In summary, we show a correlation between banding diameter and disease severity. The mPTB group mimics an early disease stage with mild RV hypertrophy without RV dysfunction, whereas the sPTB group shows signs of RV dysfunction and failure.

Figure 7: Echocardiographic, invasive pressure volume, and anatomical measurements 3 weeks after surgery. (A) Cardiac output (CO). (B) Tricuspid annular plane systolic excursion (TAPSE). (C) Right ventricular systolic pressure (RVSP). (D) End-diastolic elastance (Eed). (E) End-systolic elastance (Ees). (F) Ratio of right ventricular weight and body weight (RV/BW). (G) Correlation between right ventricular systolic pressure (RVSP) and the weight of the RV/BW at 3 weeks post-surgery. A linear regression analysis was performed and is presented in the plot. Adjusted R² = 0.75, p < 0.001. Abbreviations: mPTB: mild pulmonary trunk banding (PTB); sPTB: severe PTB. Normally distributed data are presented by scatterplots with means ± SD. Non-normally distributed data are presented by boxplots with medians with IQR and with whiskers representing values within 1.5 x IQR from Q1 and Q3, respectively *p < 0.05, **p < 0.001. Please click here to view a larger version of this figure.

Figure 8: Histological analyses of RV tissue. (A) RV cross-sectional area. (B-D) HE stained RV tissue from (B) sham, (C) mPTB, and (D) sPTB. (E) Percentage of interstitial fibrosis relative to healthy tissue in RV. (F-H) Picrosirius stained RV tissue from (F) sham, (G) mPTB, and (H) sPTB. Scalebars: (B-D) = 50 µm, (F-H) = 250 µm. Data are presented by boxplots with medians, IQR, and whiskers representing values within 1.5 x IQR from Q1 and Q3, respectively. *p < 0.05, **p < 0.001. Please click here to view a larger version of this figure.

Figure 9: Different degrees of liver discoloration. Discoloration was rated 0-3. The rating was based on the degree of speckled appearance and color of liver tissue. Pictures were taken through a microscope, making the tissue appear lighter than in ambient lighting. (A) 0: normal liver. (B) 1: Light speckles without discoloration. (C) 2: moderate speckles and light discoloration. (D) 3: Severe speckles and discoloration (nutmeg liver). Please click here to view a larger version of this figure.

Discussion

In this paper, we present a murine model of pressure overload-induced RV hypertrophy and failure. We demonstrate that: (i) PTB in juvenile mice can induce varying degrees of RV pathology, ranging from mild RV hypertrophy to RV failure with extracardiac signs of decompensation and histologically confirmed RV fibrosis. (ii) Signs of RV dysfunction can be observed and quantified by echocardiography at 1 and 3 weeks after PTB surgery. (iii) The degree of RV hypertrophy is proportional to the severity of PTB and the resulting increase in RV pressure. This model may help elucidate the pathogenesis of RV failure secondary to RV pressure overload, as we show that the PTB model is suited to study different stages of RV pathology.

To ensure success and reproducibility of the murine PTB model, great attention should be paid to the following steps. (i) Choosing the appropriate ligating clip diameter. The right clip diameter depends on several factors, such as the desired degree of RV pressure overload and follow-up time. The age and body weight of the mice are also important factors to consider. Accordingly, the protocol can be adjusted depending on the current scientific question. For this reason, we recommend performing pilot surgeries and echocardiography at relevant time points after surgery to evaluate the suitability of the selected clip diameters. (ii) Precise adjustment of the clip applier is essential to ensure reproducible pathology. We recommend using metal wire or cannulas with a well-defined outer diameter as a guide to adjust the ligating clip applier. (iii) Blood loss was the most frequent severe complication during the PTB surgery and the most common cause of peri- and postoperative mortality. To limit blood loss, use blunt microsurgical instruments and blunt dissection whenever possible. Only use surgical scissors when indicated in the protocol. (iv) Another cause of perioperative mortality is cardiac arrythmias. Short episodes of bradycardia can occur during the dissection of the pulmonary trunk and application of the ligating clip. To reduce the risk of cardiac arrest, the time of manipulation of the pulmonary trunk should be limited. When bradycardia is observed, pause manipulation of the pulmonary trunk until the heart rate has normalized. (v) Reproducible echocardiographic assessment can be challenging and requires training before reproducible results are achieved. Due to the small size of the RV, it is especially challenging to achieve good visualization of the RV in sham-operated mice in PLAX and A4CH view. (vi) Heart rate may vary significantly during echocardiographic assessment. To reduce variability, monitor the heart rate closely and use low doses of anesthesia while ensuring sufficient anesthesia.

A limitation of the murine PTB model is the need for surgical equipment and training. Pilot surgeries should always be performed for training, and the methods should be adjusted to locally available equipment. We used titanium ligating clips, which have been found to be a highly reproducible method for the induction of RV pressure overload¹⁵. Additionally, the use of ligating clips has been shown to result in lower peri-surgical mortality and better post-surgical recovery compared with surgical ligatures³¹.

Echocardiography for assessment of RV function has some limitations, as it is operator-dependent, and inter- and intraoperator variability may influence the results of echocardiography. In the present study, echocardiography was performed by two operators. To avoid bias from interobserver variation, both operators performed echocardiography on an equal number of mice from all groups. Furthermore, echocardiography in mice has its challenges. This is especially true in small and relatively healthy mice, as evidenced by the larger CO and TAPSE measurements variation at 1 week (Figure 5) compared with 3 weeks (Figure 6). To achieve a more accurate measurement of CO, VTI in the pulmonary trunk was measured in three cardiac cycles in three different locations (in the center of the pulmonary trunk and near the walls of the vessel). Further, echocardiography has been shown to overestimate CO compared with MRI in rats subjected to PTB³². MRI is the gold standard for measuring CO in mice and may therefore be considered when using the present model³². Another challenge in mice with little or no elevation of RV pressure, is the small size of the RV, complicating a good A4CH-view to measure, e.g., TAPSE. This may result in an underestimation of TAPSE in some mice in the sham and mPTB groups. Despite these challenges, statistically significant results were achieved for CO and TAPSE in sPTB compared with sham and mPTB. This shows that CO and TAPSE are useful markers of RV function in the present model, but also highlights the need for extensive training of investigators prior to echocardiographic assessment of mice.

An inherent limitation to all animal models is that results cannot be directly extrapolated to humans due to genetic and physiologic differences.

To our knowledge, this is the only PTB model in juvenile mice. This approach has been used in rats, where a gradual and slow increase in afterload develops as the rats grow. This slower disease induction better mimics the development of chronic RV failure¹⁵.

The current protocol includes two distinct groups of mice with different degrees of RV dysfunction. To our knowledge, no other published murine PTB models include a group with only mild RV hypertrophy without signs of RV dysfunction, hemodynamic impairment, or fibrosis. This early stage of the disease is thus largely overlooked. Studying this early stage of disease development may increase our understanding of the gradual progression from RV hypertrophy to RV failure seen in patients with chronic PAH¹⁷.

In this model, the pathology of chronic RV failure develops within 3 weeks after surgery, making it one of the most time-efficient models of chronic RV failure⁶. Other murine PTB models report study durations of 1 to 8 weeks^{17,18,33,34,35}. We observed signs of RV dysfunction already 1 week post-surgery in sPTB. This time-efficient approach with 3 weeks follow-up, however, comes with limitations. Mice are not fully grown at 8 weeks and a longer follow-up might result in an increasingly severe pulmonary trunk stenosis and an even more severe RV failure.

Genetically modified mice are readily available, and new genetically modified strains can be produced in mice relatively fast and time-efficiently³⁶. This constitutes a major advantage of murine PTB models in contrast to models using rats or larger mammals.

In conclusion, we present a reproducible murine model of pressure overload-induced RV hypertrophy and failure in juvenile mice. Detailed protocols for intubation, surgery, and phenotyping by echocardiography are included in the paper. Custom-made instruments are used for intubation and surgery, allowing for fast and inexpensive reproduction of the model. The model may be used to identify mechanisms that govern the RV adaptation to pressure overload, as well as processes underlying RV pathology ultimately resulting in RV failure. Finally, the model can be used to test new therapeutic targets for the treatment of RV failure.

Materials

Name	Company	Catalog Number	Comments
Biosyn 6-0, monofilament, absorbable suture	Covidien	UM-986
Blunt cannula, 27G 0.4x0.25,	Sterican	292832
Bupaq Multidose vet 0,3 mg/ml (Buprenorphinum)	Salfarm Danmark	VNR 472318
C57BL/6NTac mice	Taconic Biosciences	C57BL/6NTac
Dagrofil 1, braided, non-absorbable suture	B Braun	C0842273
Depilatory cream	Veet	3132000
Disposable scalpels, size 11	Swann-Morton	11708353
Dräger Vapor 2000 Sevoflurane	Dräger	M35054
Eye oinment neutral, "Ophta"	Actavis	MTnr.: 07586 Vnr: 53 96 68
Horizon ligating clips	Teleflex Medical	5200 (IPN914931)
Horizon Open Ligating Clips applier, curved, 6" (15 cm)	Teleflex Medical	537061
Kitchen roll holder	n.a.	n.a.
Metal wire of different thickness	n.a.	n.a.
Microsurgical instruments set	Thompson	n.a.
MiniVent Ventilator	Hugo Sachs	Type 845
MS505S transducer	Visual sonics	n.a.
Rimadyl Bovis vet. 50 mg/ml (Carprofen)	Zoetis	MTnr: 34547, Vnr: 10 27 99,
Sevoflurane Baxter 100 %	Baxter Medical	MTnr: 35015
Silicone tubing	n.a.	n.a.
Soft plastic sheet	n.a.	n.a.
Stereomicroscope, "Opmi Pico"	Carl Zeiss Surgicals GmbH	n.a.
Ultrasonic probe holder/rail	Visual Sonics	11277
Varming plate	Visual sonics	11437
Venflon ProSafety, 22G, 0,9 x 25mm	Becton Dickinson	393222