Measurement of Pulmonary Artery Pressure in Rats Using Right Heart Catheterization

Currently, pulmonary artery pressure measured by right heart catheterization is the gold standard for diagnosing pulmonary hypertension¹. In humans and large animals, the floating catheter is mostly used for right heart catheterization². However, in small animals (such as rats), due to the lack of suitable floating catheters, the application of this technique is limited, which brings inconvenience to scientific research on replicating pulmonary hypertension models using rats³. Rats typically used for modeling pulmonary hypertension generally have a weight range of 300-400 g. A smaller body size means they have finer blood vessels and thinner heart walls, which makes catheterization difficult. Although there have been literature reports on the method of making a right heart catheter with a polyethylene (PE) tube with an outer diameter of 0.9 mm and an inner diameter of 0.5 mm, and the intubation technique^3,4,5, the pressure waveform of the pulmonary artery pressure has been successfully monitored. But in actual operation, there are still many problems: the catheter is likely to enter the inferior vena cava by mistake or stay in the right ventricle for a long time and fail to enter the pulmonary artery. Prolonged catheter insertion time can also lead to a decrease in the heart rate of the rat and distorted pressure measurement. The catheter may even puncture the right atrium due to improper operation, resulting in the failure of the experiment^4,5,6. Analyzing the reasons, it may be that the toughness and hardness of the catheter affect the curvature of the catheter tip, consequently making it difficult to enter the pulmonary artery.

To solve this problem, in this study, a plasticizer-added polyvinyl chloride tube (Outer diameter 1.6 mm, inner diameter 1.0 mm, length 25 cm, suitable for jugular veins of rats over 150 g) with greater elasticity and toughness was selected as the material for making the catheter. We proposed a set of systematic operation schemes, including the making of the pigtail at the catheter tip, the dissection of the external jugular vein, the insertion of the catheter, and the determination of the catheter's position. Through improvements to the prior methods, we are able to achieve easier passage to the pulmonary artery and a higher success rate^7,8. Therefore, we can provide more reliable technical support for the research on pulmonary hypertension models in small animals.

All procedures were conducted according to the guidelines of the Institutional Animal Care Committee of the Army Medical University. The procedures followed were performed in accordance with institutional guidelines. Due to the invasiveness of the right heart catheterization, the animals should be euthanized immediately after data acquisition. Euthanasia should be performed according to the institution's animal studies guidelines. Healthy male Sprague-Dawley (SD) rats, specific pathogen-free (SPF) grade, body weight 300 ± 20 g were used here.

1. Preparation of the catheter

1. Select a venous infusion needle (PVC tube) with an outer diameter of 1.6 mm, an inner diameter of 1.0 mm, and a length of 25 cm. Cut off the needle tip. Thread a copper wire with a diameter smaller than 1.0 mm into the tube.
2. Make the tail end into a circle with a diameter of 7 mm (Figure 1A). The experimenter can use a cylindrical object of appropriate size (such as a pencil) to assist in bending the catheter. Immerse the circular end in water at 100 °C for 10 min and then take it out. Remove the copper wire. The tail end is similar to a pigtail, and the other end can be connected to a three-way stopcock (Figure 1B).

Figure 1: Fabrication of the catheter. (A) Thread a copper wire through the PVC tube and bend the end of the tube into a 7mm diameter circle. (B) After immersing the circle end in 100 ℃ water for 10 minutes, remove the copper wire. The tube is shaped, resembling the shape of a pigtail. Please click here to view a larger version of this figure.

2. Animal preparation

1. Temperature control: Turn on the heating pad and set the temperature to 37 °C. Keep the temperature at 37°C throughout the surgery.
2. Anesthetize the rat by intraperitoneal injection of 0.3 mL of 2% Sodium Pentobarbital per 100 g of body weight. To check that the animal is anesthetized, check for non-responsiveness of the paw or tail pinch. Avoid situations where animals wake up during surgery due to insufficient anesthesia, as well as deaths or other adverse effects caused by excessive anesthesia. During the surgery, continuously monitor whether the rat's heart rate and breathing are normal.
3. Place the rat in a supine position on the heating pad of a rat operating table and secure its incisors and hind limbs to the operating table using rubber bands.

3. Operative procedure

1. Shave the right cervical area and disinfect the area with a cotton swab soaked in rubbing alcohol. Use tissue scissors to make a 2 cm incision in the skin on the right side of the neck. Use curved ophthalmic forceps to bluntly dissect the external jugular vein on the right side.
2. Free about 1 cm of the vein. Ligate the distal end of the vein with a 5-0 suture thread. Tie a slipknot at the proximal end. Clamp the suture thread at the distal end with a hemostat and gently pull it towards the head to moderately tighten the blood vessel.
3. For venotomy, bend the prepared No. 7 needle 45° in the opposite direction of the needle tip (Figure 2A), and puncture the external jugular vein towards the proximal end (Figure 2B).
   CAUTION: Needles are considered sharp instruments. Avoid direct contact with the pointed end when using them.

Figure 2: Tool of Venotomy. (A) Bend the No. 7 needle 45° in the opposite direction of the needle tip. (B) Puncture the external jugular vein towards the proximal end. Due to the previous bending of the needle, the experimenter's operation will be more convenient. Please click here to view a larger version of this figure.

4. Fill the cardiac catheter connected to the three-way stopcock with heparin sodium solution and close the three-way stopcock.
   NOTE: Heparin, as an anticoagulant, can prevent blood clots from forming during catheterization. Prepare 500 mL of heparin sodium solution at 1000 IU/mL by mixing sodium heparin and normal saline.
5. Insert the bent tip of the self-made No. 7 needle into the vein, lift it up, and insert the curved ophthalmic forceps into the vein along the way to open it up (Figure 3A). Withdraw the needle. Insert the catheter into the vein through the gap opened by the forceps (Figure 3B).
6. Withdraw the curved ophthalmic forceps. Untie the slipknot, push the catheter forward, and then tie a new slipknot to fix the catheter to prevent it from falling off (Figure 3C). Do not tie the slipknot too tightly, ensuring that there are no blood leakage and the catheter can be pushed freely at the same time.

Figure 3: Catheterization. (A) Insert the curved ophthalmic forceps into the vein to open the vein up. (B) Insert the catheter into the vein through the gap opened by the forceps. (C) Tie a new slipknot to fix the catheter to prevent it from falling off. Ensure that there is no blood leakage and the catheter can be pushed freely at the same time. Please click here to view a larger version of this figure.

7. Turn on the polygraph, connect the pressure transducer, and empty the pressure transducer with normal saline to ensure there are no air bubbles.
8. Open the recording software, zero it by exposing it to the atmosphere, and then perform two-point calibration with a mercury sphygmomanometer. In the software, set the filter type to low pass, the cutoff frequency to 40 Hz, and the sampling rate to 1000 SPS.
9. Waveform measurement and position judgment: Connect the three-way stopcock connected to the catheter to the pressure transducer, ensuring there are no air bubbles in the three-way stopcock. Then open and rotate the switch of the three-way stopcock to make the catheter communicate with the pressure transducer, and the venous waveform can be seen (Figure 4A).
10. Rotate the catheter while pushing it forward. If there is resistance, do not force it to advance. Raise the pad (made with a surgical blade), pull the catheter out a little, and then rotate and push it again. When a sense of emptiness is felt, the ventricular waveform can be seen (Figure 4B).
11. Push it forward again to reach the pulmonary artery. The sliding of the catheter tip against the heart wall can be felt during advancement. The waveform of the pulmonary artery will appear (Figure 4C).
12. If the ventricular waveform shown in Figure 4D appears, it may mean that it is difficult for the catheter to enter the pulmonary artery at this position. Withdraw the catheter a little, rotate it, and push it forward until the ventricular waveform shown in Figure 4B appears, and then it is very easy to enter the pulmonary artery. At the same time, beginners can also refer to Sarkar et al. for a better understanding of waveforms at different stages⁹.

Figure 4: Waveform of Pressure. (A) The venous waveform. (B) The ventricular waveform. (C) The waveform of the pulmonary artery. (D) The abnormal ventricular waveform. Please click here to view a larger version of this figure.

4. Data collection

1. Once the characteristic waveform of the pulmonary artery pressure is confirmed and the waveform remains stable for 1 min, gently place the catheter and keep it steady. Now, measure the PAP.
2. Record a stable waveform for at least 10 consecutive cardiac cycles. If the experiment requires long-term continuous measurement of PAP, place gauze pads over the exposed skin to prevent dryness or adverse reactions caused by prolonged exposure to the environment.
3. Set the pressures of the right atrium, right ventricle, and pulmonary artery at 2-6 mmHg, 0-25 mmHg, and 10-25 mmHg, respectively, each with its characteristic peaks^{7,8,9,10,11,12,13}. When the right ventricular pressure waveform's systolic peak has a notch, the catheter is more likely to enter the pulmonary artery.
4. After data collection is completed, euthanize the animals according to the institution's animal studies guidelines. Clean the catheters after all data collection is finished.
   NOTE: All waste generated from the procedures should be properly managed. Used needles and other sharps should be immediately placed in dedicated sharps disposal containers. Biological tissues (including animal corpses) should be rendered harmless. Used chemical reagents should be poured into waste liquid tanks and discharged after centralized purification.

We have published the data on the mean pulmonary arterial pressure of rats with chronic hypoxia-induced pulmonary hypertension and normal control rats from the plains measured using this method7,8. The results are basically consistent with the data reported in the literature9,10,11,12. Using our method for experiments, the success rate can reach 95%.

In this study, we established two models of hypoxic pulmonary hypertension through the plateau environment simulation chamber (Kept the rats in a plateau environment simulation chamber simulating 5,800 m above sea level for 28 days) and chemical methods (subcutaneously injecting 1 mL of 60% monocrotaline (MCT) solution per 100 g of body weight into the neck for 28 days). Using this technology, the right ventricular systolic pressure (RVSP) and mean pulmonary arterial pressure (mPAP) measured were significantly higher than those in the control group (Table 1). The RVSP values of rats obtained in our laboratory are similar to those reported by Neelakantan et al. and Mendiola et al.11,12, while the mPAP values were similar to those reported by Sarkar et al. and Neelakantan et al.9,11. Due to factors such as modeling methods, animal strains, and depth of anesthesia, the pressure values may vary. However, the rationality of the measured pressures can be comprehensively assessed by comparing them with the control group and analyzing the pressure waveforms.

Table 1: Comparison between Chronic hypobaric hypoxia and MCT model. Chronic hypobaric hypoxia model: SD rats were kept in a plateau environment simulation chamber simulating an altitude of 5800 m for 28 days. MCT model: SD rats were subcutaneously injected in the neck with 1 mL of 60% MCT solution per 100 g of body weight and kept for 28 days.

In 1970, Swan and Ganz reported the use of a floating catheter to measure pulmonary artery pressure¹³, establishing invasive right heart catheterization as the gold standard for diagnosing pulmonary hypertension^14,15. However, in studies involving small animals such as rats, the size of the catheter limits the feasibility of closed-chest catheterization. In 1984, Bo et al.³described a method using a self-made right heart catheter with a 0.9 mm outer diameter PE tube to measure pulmonary artery pressure in rats. In recent years, multiple studies have reported improvements and optimizations in the fabrication and insertion techniques of right heart catheters^4,5,6, but several challenges remain in practical applications.

In this study, we share our experience and techniques in this field by providing detailed descriptions of catheter material selection, fabrication, isolation of the external jugular vein, catheter insertion, and catheter position determination. The detected waveform of the pulmonary artery pressure is similar to that reported in the literature^5,16. We aim to offer more reliable technical support for research on pulmonary hypertension models in small animals.

First, the choice of catheter material is critical. Previous studies used 0.9 mm PE tubes, which are thin and easy to insert into venous vessels^3,4,5. However, the slight curvature at the catheter tip often makes it difficult to pass through the clavicular segment of the jugular vein into the right atrium, leading to misplacement into the inferior vena cava or atrial perforation, resulting in experimental failure. This issue may be attributed to the catheter's flexibility and elasticity, which affect the curvature of the tip and hinder its advancement into the pulmonary artery.

In this study, we used intravenous infusion needles made of polyvinyl chloride (PVC) with the addition of a plasticizer, di(2-ethylhexyl) phthalate (DEHP). Compared to PE tubes, these catheters are softer in hardness and also possess memory properties. Softer hardness means that the tip of the catheter has some cushioning when pressed against the heart wall, making it less likely to get stuck and pierce the cardiac wall. The memory property is reflected in its ease of shaping. During the catheter manufacturing process, after the copper wire is removed, the tip of the catheter still retains a curve. During catheterization, despite pulling, the tip of the catheter consistently tends to maintain this curve, making it easier for the catheter to enter the pulmonary artery. The catheter tip maintains a straight shape in the vessel and easily passes through the clavicular segment of the jugular vein. When reaching the right ventricle, the catheter forms a curve (previously shaped), and the tip can sway with the blood flow, facilitating its smooth entry into the pulmonary artery. The outer diameter of the catheter is 1.6 mm, which may seem relatively large to beginners, making venous insertion challenging. However, using the methods described in this study, the catheter can be successfully inserted into the jugular vein of rats weighing over 150 g. Alternatively, a polyurethane catheter or a silicone tube (outer diameter: 1.4 mm, inner diameter: 0.8 mm) can also be chosen. The pigtail tip of the catheter is shaped as described in this study, while the other end is connected to a blunt 12 G needle and then to a three-way stopcock. The pigtail tip is formed into a 7 mm diameter circle by soaking in hot water for 10 min. After removing the copper wire, the catheter's curvature is suitable for rats weighing 200-400 g.

In the literature, ophthalmic scissors are often used to create a V-shaped incision in the vessel, which can easily lead to vessel transection, especially for beginners. In this study, a curved 7-gauge needle is used to puncture the vessel. Before puncturing, a pad is placed beneath the vessel to facilitate the puncture. The needle is then lifted upward, creating an incision in the vessel. An ophthalmic curved forceps (with a smooth, polished tip) is inserted into the vessel to widen the opening, allowing the catheter to be inserted.

Regarding catheter position determination, many studies mark the catheter with scales, which can be helpful for beginners^4,5,9. However, in this study, this step is not considered mandatory. Right heart catheterization is essentially a blind insertion technique, and the position is primarily determined by pressure waveform analysis.

Due to the experimenter's error, a series of injuries may occur. Here, we propose methods to detect the occurrence of these injuries. On the one hand, from a tactile perspective, if resistance is felt but the catheter is still forcibly advanced, and then there is a sudden loss of resistance with the catheter unexpectedly advancing deeply, the heart wall or the blood vessel may have been pierced. On the other hand, from a waveform perspective, the appearance of negative pressure may indicate damage to the heart and the blood vessels. After an injury occurs, the animal is likely to die quickly from bleeding. When the catheter is in the right ventricle but has not yet reached the pulmonary artery, with the prolongation of catheter insertion time, the displayed right ventricular waveform shows a decrease in systolic pressure. If the rat's heart rate and breathing are normal, the catheter tip may be blocked by a blood clot. The beginners can use a three-way stopcock to push about 0.5 mL of heparin sodium saline to clear the catheter.

In summary, right heart catheterization is a complex and technically demanding experimental procedure. Through systematic learning and practice, beginners can gradually master this technique and improve experimental success rates. We hope that the insights shared in this study will provide valuable references for researchers in related fields and contribute to advancements in the field of pulmonary hypertension research.