Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove


Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice

doi: 10.3791/60090 Published: October 24, 2019


Here, we present a protocol to perform an invasive hemodynamic assessment of the right ventricle and pulmonary artery in mice using an open-chest surgery approach.


Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. However, the evaluation of right ventricular pressure (RVP) and pulmonary artery pressure (PAP) remains technically challenging in mice. RVP and PAP are more difficult to measure than left ventricular pressure because of the anatomical differences between the left and right heart systems. In this paper, we describe a stable right heart hemodynamic measurement method and its validation using healthy and PAH mice. This method is based on open-chest surgery and mechanical ventilation support. It is a complicated procedure compared to closed chest procedures. While a well-trained surgeon is required for this surgery, the advantage of this procedure is that it can generate both RVP and PAP parameters at the same time, so it is a preferable procedure for the evaluation of PAH models.


Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder with elevation in pulmonary artery pressure (PAP) and right ventricular pressure (RVP) that is caused by cellular proliferation and fibrosis of small pulmonary arteries1. Pulmonary artery catheters, also called Swan-Ganz catheters2, are commonly used in the clinical monitoring of RVP and PAP. Furthermore, a wireless PAP monitoring system has been used clinically3,4,5. To mimic the disease for study in mice, a hypoxic environment is used to simulate human clinical manifestations of PAH6. In the evaluation of PAP in animals, large animals are relatively easy to monitor through pulmonary artery catheters using the same technique as for human subjects, but small animals such as rats and mice are difficult to assess because of their small body size. Hemodynamic measurement of the right ventricular system in mice is possible with an ultrasmall size 1 Fr catheter7. A method for measuring RVP and PAP in mice has been reported in the literature8,9, but the methodology lacks a detailed description. RVP and PAP are more challenging to measure than left ventricular pressure because of the anatomical differences between the left and right heart systems.

To get both PAP and RVP parameters in the same mouse, we describe an open-chest surgery-based approach for right heart hemodynamic measurements, its validation with healthy and PAH mice, and how to avoid generating artificial data during the complicated open-chest surgery. Although this technique is best performed by a well-trained surgeon, it has the advantage of being able to assess PAP and RVP in the same mouse.

Subscription Required. Please recommend JoVE to your librarian.


The animal protocol was reviewed and approved by the Institutional Animal Care and Use Committee at Fuwai Hospital, Chinese Academy of Medical Science, Peking Union Medical College (NO.0000287). The experimental animals were housed and fed according to the guidelines of animal welfare in China.

NOTE: Eight- to 12-week-old male C57BL mice were housed in an environment with a 12 h dark/ 12 h light cycle. The PAH mice were housed for 4 weeks under an oxygen concentration of 10%, maintained by an oxygen-controlled hypoxia chamber to induce pulmonary hypertension, and control mice were housed in room air (21% oxygen) under identical conditions. RVP and PAP measurements were performed at the end of the 4 weeks of hypoxia challenge.

1. Preoperative preparation

  1. Soak the pressure transducer catheter (size: 1 Fr) in 0.9% saline at room temperature for at least 30 min before the hemodynamic experiment.
  2. Filter 2,2,2-Tribromoethanol solution with 0.22 μm filter and store in 4-degree refrigerator.
  3. Prepare cleaned surgery tools and supplies such as gloves for surgery.
  4. Prepare 10 mL of 1.0% digestive enzyme solution for catheter cleaning.
  5. Connect the pressure transducer catheter to a pressure-volume system.
  6. Calibrate the pressure transducer prior to obtaining pressure measurements for each mouse.
    1. Turn the calibration knob to 0 mmHg and 25 mmHg to send a verification pressure signal to the data acquisition software and configure the calibration setting in the software.
    2. Turn the knob to Transducer and adjust the Balance knob to zero baseline.
  7. Set up a standard stereomicroscope and a temperature-controlled small animal surgical table for body temperature maintenance during the surgery.
  8. Set up a light illumination system for microsurgery to provide enough light over the surgical area.

2. Open-Chest surgery and hemodynamic measurement

  1. Anesthetize mice with 250 mg/kg of 2,2,2-Tribromoethanol via intraperitoneal (i.p.) injection. If needed, repeat supplemental doses at 1/3 to 1/2 of the original dose during the procedure.
  2. Remove chest and neck fur using a shaver and hair removal lotion (Figure 1A, 2A).
  3. Secure each mouse in the supine position on a temperature-controlled small animal surgical table to help maintain body temperature (37 °C) during surgery.
  4. Clean the surgical site with 70% ethanal.
  5. Once anesthesia is in effect, confirm adequate anesthesia induction using a toe pinch.
  6. Make a midline incision on the neck skin (Figure 1A).
  7. Dissect the skeletal muscle using curved forceps and expose the trachea (Figure 1B, 1C).
  8. Perform intubation through the mouth using a modified 22 G intravenous sheath catheter. Confirm that the tubing is in the trachea using forceps (Figure 1D).
  9. Connect the tubing to a small animal ventilator. Calculate and set respiration rate and tidal volume based on body weight according to the ventilator user manual10. For example, set respiration rate to 133/min and tidal volume to 180 μL for a 30 g mouse based on the described calculation.
  10. Secure the tubing for ventilation using tape.
  11. Confirm adequate anesthesia induction using a toe pinch.
  12. Make a midline incision on the chest skin and carefully dissect the chest muscles using a cautery tool (Figure 2B, 2C).
  13. Cut the sternum using scissors across the middle and expose the thoracic cavity (Figure 2D).
  14. Prevent any bleeding using the cautery tool during the open-chest surgery procedure.
  15. Expose the right ventricle with retractors (Figure 2E).
  16. Insert the saline-soaked pressure transducer catheter through a small tunnel created with a 25 G needle into the right ventricle to measure RVP (Figure 2F and Figure 3A, 3C).
  17. Hold the catheter cable and cross the pulmonary valve in a coaxial manner with the pulmonary artery. Observe the pressure waveform and obtain a stable PAP signal (Figure 3B, 3D).
  18. Record hemodynamic data using the data acquisition system and software.
  19. After the final measurements, euthanize mice humanely through i.p. injection of an excess dose of 2,2,2-Tribromoethanol solution.
  20. Carefully remove catheter from the right heart system and place into a 1 mL syringe containing 1% digestive enzyme solution.
  21. Use distilled water to continuously flush the catheter carefully and store it in the original box.

3. Data analysis for hemodynamics

NOTE: The hemodynamic data were recorded and analyzed using analysis software11 (Table of Materials).

  1. For each mouse, select at least 10 continuous and stable heartbeat cycles without noise to obtain the average data of RVP or PAP data for each parameter.
  2. Use Student’s t-test to compare the normal air control and hypoxia groups. NOTE: p < 0.05 was considered statistically significant. Data are presented as the mean ± SD.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

The pressure transducer catheter was inserted into the right ventricle (Figure 3A) through a tunnel expanded by a 25 G needle, and a typical RVP waveform (Figure 3C) was obtained. The catheter was continually adjusted and slowly advanced and kept in the same axis as the pulmonary artery while passing through the pulmonary valve (Figure 3B). When the pressure sensor was successfully inserted into the pulmonary artery, a typical PAP waveform with a characteristic dicrotic notch appeared (Figure 3D). To avoid the generation of artificial data, we observed whether the waveform had noise (Figure 4) or whether the zero level of the catheter had drifted (Figure 5). If this occurred, corrections were made, and these segments with noise were excluded from data analysis.

PAH is characterized by a sustained elevation in PAP and RVP, caused by increased resistance in small pulmonary arteries. PAH is defined by a mean PAP of ≥25 mmHg at rest, measured during right heart catheterization in the clinic12. We measured the RVP and PAP in the mice with the induced chronic hypoxia (kept at 10% oxygen for 4 weeks) or a control group (kept in normal air). The results are shown in Figure 6. Compared to those in the normal air control group, systolic PAP (Figure 6A), diastolic PAP (Figure 6B), mean PAP (Figure 6C), and right ventricular systolic pressure (Figure 6D) were all significantly increased in the chronic hypoxia group. Investigators have also reported that compared with hypoxia alone, a combination of a VEGFR inhibitor with chronic hypoxia for 3 weeks to induce severe PAH in mice can result in significantly increased RVP13,18.

Figure 1
Figure 1: Intubation for mechanical ventilation support in mice. (A) The neck fur is removed using hair removal lotion to obtain a clean area for surgery. A midline incision is made on the skin of the neck. (B) The skeletal muscle covering the trachea is exposed. (C) The skeletal muscles are bluntly dissected to expose the trachea. The yellow arrow indicates the trachea. (D) The tubing (modified using a 22 G intravenous catheter) is inserted into the airway, with placement confirmed using forceps. The yellow arrow indicates the tubing inside the trachea. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Open-chest surgery for hemodynamic measurement at the right ventricular system. (A) The chest fur is removed using hair removal lotion to obtain a clean area for surgery. (B) A midline incision is made to expose chest skeletal muscles and the sternum. (C) A cautery tool is used to minimize bleeding during chest opening (the arrow indicates the cautery tip). (D) The sternum is cut along the midline (the yellow dash line). (E) Two retractors are used to expose the heart (the upper arrow indicates the right atrial wall, and the lower arrow indicates the right ventricular free wall). (F) A pressure transducer catheter (the lower arrow) is inserted into the right ventricular chamber using a puncture tool (25 G size needle, the upper arrow) to produce a small tunnel on the right ventricular free wall. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Representative RVP and PAP curves. The pressure transducer catheter is inserted into the right ventricular chamber (A) to obtain the RVP waveform (C). The pressure transducer catheter goes through the pulmonary valve and then stays in the pulmonary artery (B) to generate the PAP waveform. The arrows indicate the characteristic dicrotic notch on the PAP waveform (D), which is a sign of a pulmonary valve closure. RA = right atrium, RV = right ventricle, PA = pulmonary artery, LV = left ventricle. Please click here to view a larger version of this figure.

Figure 4
Figure 4: RVP waveform noise caused by touching of the pressure sensor surface to the ventricular wall. The arrow point shows a sharp increase in pressure on the RVP curve (the upper channel), which simultaneously produces an artificial change in dP/dt (the lower channel). dP/dt is calculated from RVP. The dashed lines indicate dP/dt noise. If the noise is constantly present, adjustment of the catheter sensor position in the ventricle can prevent noise. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Zero drift of pressure transducer during RVP measurement. The left window shows artificially slightly elevated end-diastolic RVP. The right expanded window shows increased end-diastolic RVP (arrows indicate end-diastolic RVP). Please click here to view a larger version of this figure.

Figure 6
Figure 6: Hypoxia-induced pulmonary artery hypertension in C57BL mice. (A) Systolic PAP (sPAP). (B) Diastolic PAP (dPAP). (C) Mean PAP (mPAP). (D) Right ventricular systolic pressure (RVSP). (E) and (F), Representative PAP waveforms for control and PAH mice respectively *p < 0.05; Student's t-test; control group n = 10; hypoxia group n = 3. Data are presented as the mean ± SD. PAP = pulmonary artery pressure, RVP = right ventricular pressure. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.


Tracheal intubation is the first important step for open-chest surgeries. The classic method of tracheal intubation for small animals, such as rats or mice, involves making a T-shaped incision on the trachea and directly inserting Y-type tracheal tubing into the trachea. In practice, we find that this method is not easy during operation. The Y-type tracheal tubing is too large for small animals and forms an angle with the trachea. Thus, it is difficult to fix the tubing in place. Additionally, once the intubation tubing accidentally comes out from the airway during open-chest surgery, it usually results in animal death because of loss of mechanical ventilation support. Therefore, we modified the method of endotracheal intubation14 by making an incision on the skin, separating the muscle layer to expose the trachea (Figure 1C), and directly inserting the tracheal tubing into the airway through the animal's mouth. The placement of tubing in the trachea can be conveniently confirmed by clamping the trachea using forceps (Figure 1D). After removing the guide needle and only using the sheath catheter, a 22 G intravenous catheter is used as the intubation tubing. The tubing can be easily secured after intubation. This is a safe way to manage intubation during surgery and can significantly improve the success rate of small animal open-chest surgery. However, this method requires training and practice.

The closed-chest approach for right heart hemodynamic measurement has been described in detail15,16. One limitation of the closed-chest method is that it can be used to only evaluate RVP, because the catheter cannot access the pulmonary artery in mice. We use a midline chest incision where the right ventricular free wall is located, just below the sternum (Figure 2D). After right ventricular catheterization to obtain RVP, it is easy to insert the catheter in a coaxial manner with the pulmonary artery to get PAP (Figure 2E). When the sternum is cut during the open-chest surgery, an electrocoagulation tool is used to avoid sternal cutting section bleeding to prevent artificial blood pressure decrease caused by blood loss (Figure 2C). It is optional for this open-chest surgery to use a P-V loop catheter to get both RVP and volume information17. However, it is best to not use it to obtain PAP because of its bigger size. Although this method is best performed by a well-trained surgeon, it is preferable to the closed-chest approach because it allows for the maintenance of tracheal intubation and prevention of bleeding during open-chest surgery to avoid animal death.

Additionally, the right ventricular free wall is punctured with a 25 G or smaller needle to reduce resistance during the insertion of the catheter into the ventricle. During catheterization,  the pressure sensor surface must not deviate from the bevel of the needle to prevent accidental damage to the catheter sensor by the sharp metal surface. It is preferable to not use a large needle to puncture the ventricular free wall, as it usually causes further bleeding, and the insufficient blood volume in circulation also causes artificial pressure data.

Because of the small volume of the ventricle and the irregular size of the right ventricular chamber in mice, the pressure sensor of the catheter easily touches the right ventricular free wall during the high heartbeat rate. This generates noise on the ventricular pressure curve (Figure 4), directly affecting ventricular pressure analysis. In this case, the angle and depth of the catheter should be adjusted until the noise disappears to obtain a smooth ventricular pressure waveform again.

The small size of the 1 Fr pressure transducer catheter7 makes it a very precise, accurate pressure transducer. Zero drift is generally not experienced during a standard catheter test in saline solution in vitro unless the catheter is faulty or damaged. However, in the presence of body blood, blood components adhering to the pressure sensor surface may cause the catheter to undergo zero drift during an in vivo experiment (Figure 5). To address this issue, we do the following: temporarily remove the catheter out from the ventricular chamber and place the sensor tip of the catheter into warm 1.0% digestive enzyme solution; incubate it to digest the blood components attached to the sensor surface; and after gently wiping the catheter with saline-soaked gauze, insert the catheter back to the ventricular chamber to obtain a stable, non-zero drift ventricular pressure waveform.

The preparation of a pressure transducer catheter is also essential to obtain stable data. The pressure sensor tip of the catheter must be soaked for at least 30 min in 0.9% saline at room temperature before the in vivo procedure to maintain the stability of the catheter. In this way, the electrical characteristics of the pressure transducer catheter can be optimally stabilized.

Finally, the hypoxia period is viable from 3 to 4 weeks for the hypoxia-induced hypertension model in mice6,14,17,18. Our data showed that 4 weeks of hypoxia can induce a stable pulmonary hypertension model in C57BL mice, and the PAP and RVP levels are comparable with the literature. Further study is needed to address how long the PAH model can be maintained if the mice are put back in normoxic conditions for different hypoxia protocols.

Subscription Required. Please recommend JoVE to your librarian.


The authors have nothing to disclose.


This research is supported by the Postgraduate Education and Teaching Reform Project of Peking Union Medical College (10023-2016-002-03), the Fuwai Hospital Youth Fund (2018-F09), and the Director Fund of Beijing Key Laboratory of Pre-clinical Research and Evaluation for Cardiovascular Implant Materials (2018-PT2-ZR05).


Name Company Catalog Number Comments
2,2,2-Tribromoethanol Sigma-Aldrich T48402-5G For anesthesia
Animal temperature controller Physitemp Instruments, Inc. TCAT-2LV For temperature control
Dissection forceps Fine Science Tools, Inc. 11274-20 For surgery
Gemini Cautery System Gemini GEM 5917 For surgery
Intravenous catheter (22G) BD angiocath 381123 For intubation
LabChart 7.3 ADInstruments For data analysis
Light illumination system Olympus For surgery
Mikro-Tip catheter Millar Instruments, Houston, TX SPR-1000 For pressure measurement
Millar Pressure-Volume Systems Millar Instruments, Houston, TX MVPS-300 For pressure measurement
O2 Controller and Hypoxia chamber Biospherix ProOx 110 For chronic hypoxia
PowerLab Data Acquisition System ADInstruments PowerLab 16/30 For data recording
Scissors Fine Science Tools, Inc. 14084-08 For surgery
Small animal ventilator Harvard Apparatus Mini-Vent 845 For surgery
Stereomicroscope Olympus SZ61 For surgery
Surgery tape 3M For surgery
Terg-a-zyme enzyme Sigma-Aldrich Z273287-1EA For catheter cleaning



  1. Humbert, M., et al. Advances in therapeutic interventions for patients with pulmonary arterial hypertension. Circulation. 130, (24), 2189-2208 (2014).
  2. Chatterjee, K. The Swan-Ganz catheters: past, present, and future: a viewpoint. Circulation. 119, (1), 147-152 (2009).
  3. Adamson, P. B., et al. CHAMPION trial rationale and design: the long-term safety and clinical efficacy of a wireless pulmonary artery pressure monitoring system. Journal of Cardiac Failure. 17, (1), 3-10 (2011).
  4. Abraham, W. T., et al. Wireless pulmonary artery haemodynamic monitoring in chronic heart failure: a randomised controlled trial. The Lancet. 377, (9766), 658-666 (2011).
  5. Adamson, P. B., et al. Wireless pulmonary artery pressure monitoring guides management to reduce decompensation in heart failure with preserved ejection fraction. Circulation: Heart Failure. 7, (6), 935-944 (2014).
  6. Shatat, M. A., et al. Endothelial Kruppel-like Factor 4 modulates pulmonary arterial hypertension. American Journal of Respiratory Cell and Molecular Biology. 50, (3), 647-653 (2014).
  7. SPR-1000 Mouse Pressure Catheter. Available from: https://millar.com/products/research/pressure/single-pressure-no-lumen/spr-1000 (2019).
  8. Tabima, D. M., Hacker, T. A., Chesler, N. C. Measuring right ventricular function in the normal and hypertensive mouse hearts using admittance-derived pressure-volume loops. American Journal of Physiology Heart and Circulatory Physiology. 299, (6), 2069-2075 (2010).
  9. Skuli, N., et al. Endothelial deletion of hypoxia-inducible factor-2alpha (HIF-2alpha) alters vascular function and tumor angiogenesis. Blood. 114, (2), 469-477 (2009).
  10. Harvard Inspira Advanced Safety Ventilator User's Manual. Available from: http://www.harvardapparatus.com/media/harvard/pdf/Inspira_557058_9.pdf. (2019).
  11. LabChart. Available from: https://www.adinstruments.com/products/labchart?creative=290739105773_keyword=labchart_matchtype=e_network=g_device=_gclid=CjwKCAjwxrzoBRBBEiwAbtX1n42I2S06KmccVncUHkmExU8KKOXXREyzx8bvTrxYMSze-ooE0atcbRoCliwQAvD_BwE (2019).
  12. Marius, M. H., et al. Definitions and diagnosis of pulmonary hypertension. Journal of the American College of Cardiology. 62, (25), 42-50 (2013).
  13. Ciuclan, L., et al. A novel murine model of severe pulmonary arterial hypertension. American Journal of Respiratory and Critical Care Medicine. 184, (10), 1171-1182 (2011).
  14. Brown, R. H., Walters, D. M., Greenberg, R. S., Mitzner, W. A. A method of endotracheal intubation and pulmonary functional assessment for repeated studies in mice. Journal of Applied Physiology. 87, (6), 2362-2365 (1999).
  15. Chen, W. C., et al. Right ventricular systolic pressure measurements in combination with harvest of lung and immune tissue samples in mice. Journal of Visualized Experiments. (71), 50023 (2013).
  16. Ma, Z., Mao, L., Rajagopal, S. Hemodynamic characterization of rodent models of pulmonary arterial hypertension. Journal of Visualized Experiments. (110), 53335 (2016).
  17. Chen, M. Berberine attenuates hypoxia-induced pulmonary arterial hypertension via bone morphogenetic protein and transforming growth factor-β signaling. Journal of Cellular Physiology. (2019).
  18. Bueno-Beti, C., Hadri, L., Hajjar, R. J., Sassi, Y. The Sugen 5416/Hypoxia mouse model of pulmonary arterial hypertension. Experimental Models of Cardiovascular Diseases. Methods in Molecular Biology. vol 1816. Ishikawa, K. Humana Press. New York, NY. (2018).
Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice
Play Video

Cite this Article

Luo, F., Wang, X., Luo, X., Li, B., Zhu, D., Sun, H., Tang, Y. Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice. J. Vis. Exp. (152), e60090, doi:10.3791/60090 (2019).More

Luo, F., Wang, X., Luo, X., Li, B., Zhu, D., Sun, H., Tang, Y. Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice. J. Vis. Exp. (152), e60090, doi:10.3791/60090 (2019).

Copy Citation Download Citation Reprints and Permissions
View Video

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
simple hit counter