Multiple Intravenous Bolus Dosing and Invasive Hemodynamic Assessment in a Hypoxia-Induced Mouse Pulmonary Artery Hypertension Model

Summary

This protocol provides a step-by-step procedure for executing multiple intravenous bolus dose administration and invasive hemodynamic monitoring in mice. Investigators can use this protocol for future therapeutic compound screening for pulmonary artery hypertension.

Abstract

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, there is no cure for PAH. It is important to discover new compounds that can be used to treat PAH. The mouse hypoxia-induced PAH model is a widely used model for PAH research. This model recapitulates human clinical manifestations of PAH Group 3 disease and is an important research tool to evaluate the effectiveness of new experimental therapies for PAH. Research using this model often requires the administration of compounds in mice. For a compound that needs to be given directly into the bloodstream, optimizing intravenous (IV) administration is a key part of the experimental procedures. Ideally, the IV injection system should permit multiple injections over a set time course. Although the mouse hypoxia-induced PAH model is very popular in many laboratories, it is technically challenging to perform multiple IV bolus dosing and invasive hemodynamic assessment in this model. In this protocol, we present step-by-step instructions on how to carry out multiple IV bolus dosing via mouse jugular vein and perform arterial and right ventricle catheterization for hemodynamic assessment in mouse hypoxia-induced PAH model.

Introduction

Pulmonary artery hypertension (PAH) is defined by a mean pulmonary artery systolic pressure greater than 20 mmHg at rest^1,2. It is a progressive and fatal disease characterized by a sustained elevation in pulmonary arterial pressure, leading to right ventricle overload and ultimately death due to right ventricular failure¹. Currently, there is no cure for PAH.

The use of animal models of pulmonary hypertension is important for testing the effectiveness of experimental PAH therapies. Among those models, the mouse hypoxia-induced PAH model has provided key insights into human PAH group 3 disease development^3,4. Research using this model often requires the administration of compounds in mice to evaluate the novel compound's effectiveness and safety. Therefore, investigators need a detailed experimental procedure for compound dosing and hemodynamic measurements to ensure injection consistency and blood pressure measurement reproducibility from the beginning to the end.

Methods for intravenous (IV) injection and blood pressure measuring have been reported in the literature^5,6. However, the methodology lacks visual illustration and detailed description. Here we illustrate the key steps for a successful IV bolus injection and accurate measurement and recording of systemic and right ventricle blood pressure. The procedures presented here are an important resource for investigators interested in the IV route of compound administration platform to develop a treatment for PAH.

Protocol

All animal procedures were performed under protocols approved by Yale University Institutional Animal Care and Use Committees.

1. Preparation of animals, tools, blood pressure measuring equipment, and hypoxia chamber

1. Animal acclimation.
   NOTE: Experimental animals used for this study were male, 8-week-old C57BL/6 mice weighing 25-27 g. Several factors should be considered when estimating the number of animals required for the experiment, including surgery-associated mortality, unexpected surgical complications, and sudden unexpected death. Use at least 10 mice per group to reach statistical power and avoid underpowered studies.
   1. Upon reception, house the animals in ventilated rodent cages (groups of five animals per cage) provided with appropriate bedding, rodent chow, and water. Let the animals acclimate to the new environment (12 h light-dark cycle at 18-20 °C) for at least 3 days.
   2. Randomly assign them to the following groups: Normoxia (Group 1), hypoxia (Group 2), and hypoxia + 7C1/let-7 miRNA (Group 3).
2. Surgical tools and blood pressure measuring equipment preparation.
   1. Sterilize all surgical tools by autoclaving (Figure 1A).
   2. Prepare a makeshift injection platform with a homemade anesthesia nose cone (Figure 1B), suture packs (Figure 1C), and equipment for the PAH procedure (Figure 1D-F).
3. Experimental setting for pulmonary artery hypertension (PAH) induction.
   1. Set the N₂ tank, the oxygen sensor, and the semisealable hypoxia chamber (Figure 2A).
   2. Establish a set point of 10% O₂ in the oxygen sensor and let the system reach the steady-state (Figure 2B, C).
   3. Keep hypoxia (Group 2) and hypoxia + 7C1/let-7 miRNA (Group 3) animals in hypoxia (10% O₂) for 3 weeks. After 3 weeks of hypoxia, place the animals under normoxic conditions for 1 week (Figure 2D). The normoxic group (Group 1) stays in normoxia for 4 weeks.
      NOTE: (1) 3 weeks of hypoxia followed by 1 week of normoxia is a well-established method for developing PAH and right ventricle heart failure⁷. The oxygen sensor detects O₂ concentration inside the semisealable hypoxia chamber and corrects it by infusing N₂ gas through the gas infusion tube.
   4. Inspect the animals daily for the entire duration of the experiment (3 weeks). Consult a veterinarian if the animals exhibit signs of distress such as dramatic weight loss and difficulty breathing. If euthanasia is necessary for the animals in severe distress, exclude the animal from the study.
      NOTE: Hypoxia exposure causes mouse body weight loss. A loss of 10% body weight is typically used as a reliable indication of PAH development.
   5. Avoid extensive opening of the hypoxia chamber. For cage cleaning, replenishing food, changing water bottles, and compound administration, open the chambers for not more than 1 hour per week.

2. Intravenous bolus injection  via the jugular vein

1. Mouse preparation and anesthesia.
   1. Take out hypoxia (Group 2) and hypoxia + 7C1/let-7 miRNA (Group 3) mouse cages from the hypoxia chamber and gently remove the animal from the cage.
      NOTE: The dosage regimen for 7C1/let-7 miRNA (1.5 mg/kg IV/dose) is twice per week for 4 weeks of treatment. It is recommended that investigators take both hypoxia and hypoxia + compound treatment cages out of the hypoxia chamber during IV injection to make sure all the animals receive the same magnitude of hypoxia exposure per time interval.
   2. Weigh the mouse using a precision scale and record its weight (Figure 3A).
   3. Place the mouse into an anesthesia induction chamber connected to the anesthetic vaporizer and close it (Figure 3B). Provide thermal support and apply eye lubricant to both eyes to prevent drying while anesthetizing. Expose the mouse to 3% isoflurane until unconscious (Figure 3C-D).
   4. Remove the mouse from the chamber and shave the fur from the jaw cranially to the middle of the sternum caudally. Laterally, shave the fur from the jaw angles, through the sides of the neck, and toward the shoulders (Figure 3E).
   5. Place the isoflurane-anesthetized mouse in a supine position (belly side face-up) on an injection platform underneath a dissection microscope. Maintain the anesthesia via a nose cone with 1.5% isoflurane and gently restrain the four legs with adhesive tape to immobilize the body (Figure 3F).
   6. Apply a noxious stimulus (i.e., toe pinch) with straight forceps to ensure an adequate level of anesthesia. The anesthetized mouse should not respond to the stimulation prior to and throughout the surgical procedure.
2. Injection agent preparation.
   1. Prepare a single-dose injection compound at a dose of 1.5 mg/kg under sterile conditions.
      NOTE: Warm the injection compound to room temperature (RT) since injection of cold substances can cause discomfort and a drop in mouse body temperature (if this does not damage the compound). The optimal dose and duration of compound 7C1/let-7 miRNA used in this study are based on previous publications^8,9.
   2. Load the sterile single-use syringe with the volume to be injected. Hold the syringe upright and advance the plunger to expel the air from the syringe. Do not reuse the syringe.
   3. Limit the injection volume to 200 μL in a 25 g mouse to reduce the incidence of hemodilution and abnormal cardiac effects on the animals. If a larger volume is required, divide the injection compound into two injections with an interval of 10 min.
3. Prepare mouse for IV injection.
   1. Gently scrub the surgical area three times with three alternating rounds of povidone-iodine solution and 70% ethanol. Administer buprenorphine (0.05 mg/kg, SQ) 30 min prior to the surgical procedure.
   2. Make a 0.5 cm longitudinal cut slightly to the right of the midline of the neck using a scalpel blade (Figure 3G).
   3. Use forceps to separate the muscle and the fat tissues to locate the right external jugular vein (Figure 3H).
      NOTE: Rotate injection sites each time to avoid scar formation.
   4. Use a high-power objective lens to allow easy visualization of the injection area (Figure 3I).
4. IV injection
   1. Insert a 28 G sterile needle into the jugular vein with the needle bevel facing up (Figure 3J, K).
      NOTE: Tail vein injection is an alternative to jugular vein injection. However, this technique is difficult to carry out repeat-dosing due to the variability in vein depth, mice tail skin color, and skin hardness.
   2. Slowly press the syringe plunger to inject the compound into the vein. Allow the needle to remain within the vein for an additional 10 s to prevent backflow of the injectant (Figure 3L).
      NOTE: The bluish dye allows for easy visualization of the injection. Do not include the dye when injecting test materials. An inaccurate injection will result in the accumulation of bluish dye around the IV injection site.
   3. Remove the needle and use a cotton swab to apply pressure to the injection site to prevent bleeding (Figure 3M).
   4. Suture the skin with 5-0 suture (Figure 3N). After surgery, move the animal to a warm, clean, dry area and provide Meloxicam (1 mg/kg, SQ, q24h). Place the animal in a clean recovery cage with no bedding but the bottom covered in a paper towel. 
      NOTE: The mouse should be awake from anesthesia and regain consciousness within 5 min once back to the recovery cage. Monitor the mouse for signs of distress.
   5. Return the animals to their home cage and put the mouse cage back in the hypoxia chamber.
      ​NOTE: The entire procedure, from anesthetizing a mouse to finishing jugular vein injection, takes about 10-15 min by a single experimenter. To shorten the normoxia exposure in mice, it is recommended that at least two investigators collaborate to accomplish a jugular vein injection procedure.

3. Blood pressure measurement

1. Prepare instruments for blood pressure measuring.
   1. Soak the tip of 1.0 F catheter in 37 °C pre-heated PBS at least 30 min before hemodynamic measurement (Figure 4A).
   2. Measure the distance from the catheter insertion site to the desired catheter tip location. For example, the distance between the mouse ascending aorta to the mid of the neck is approximately 1-1.2 cm. The distance between the right ventricle of the heart to the mid of the neck is approximately 2.3-2.8 cm.
   3. Mark two catheter distance markings to provide a visual indication of insertion depth (Figure 4B).
   4. Attach the catheter to the pressure transducer, connect the pressure transducer to input channel 1 on the data acquisition device, turn on the pressure-volume control unit, and initiate data acquisition blood pressure analysis software. Create a new blood pressure analysis document and set channel 1 for pressure.
   5. Perform a pressure calibration according to the manufacturer's protocol. Allow the whole setup to stabilize for at least 5 min (Figure 4C).
   6. In blood pressure analysis software, select Units Conversion from the Channel 1 dropdown menu (Figure 4D, red arrow).
   7. Set the default Units Conversion Values (Figure 4E).
      NOTE: Blood pressure is represented as millimeters of mercury (mmHg). The standardized pressure output from Pressure Control Unit is 1 V per 100 mmHg. 25 mmHg corresponds to 0.25 V output, and 100 mm Hg corresponds to 1 V output.
2. Prepare mouse for blood pressure measurement procedure.
   1. Anesthetize the mouse with 3% isoflurane inhalation through a nose cone.
   2. Apply the veterinary ointment directly onto the ocular surface of the mouse eyes to prevent dryness, as the mouse cannot close its eyes under anesthesia. Shave the fur from the mouse's neck while under anesthesia.
   3. Scrub the shaved region with three alternating rounds of povidone-iodine solution and 70% ethanol swab. Place the anesthetized mouse in a supine position on an injection platform underneath a dissection microscope. Place the mouse's nose into the nose cone to maintain anesthesia (1.5% isoflurane) throughout the surgical procedure.
   4. Test the anesthetized mouse's motor response to the noxious stimulus. The anesthetized mouse should not respond to a noxious stimulus before and during the surgery.
      NOTE: Inhalable (isoflurane) and injectable (ketamine/xylazine) anesthetics can decrease blood pressure. In general, isoflurane inhalation anesthesia has a slight effect on lowering the blood pressure than ketamine/xylazine. Therefore, isoflurane is the preferred inhalant anesthetic over ketamine/xylazine. Achieving the appropriate depth of anesthesia is critical for accurate and reproducible hemodynamic measurements. The investigator needs to keep the depth of anesthesia constant for each mouse.
3. Ascending aorta catheterization
   1. Apply a noxious stimulus (i.e., toe pinch) with straight forceps to ensure an adequate level of anesthesia. Make a midline incision of the skin from the mandible to the sternum (Figure 5A).
   2. Separate the salivary glands and expose the trachea (Figure 5B).
   3. Use forceps to clear the soft tissue along the vessels to expose the right carotid artery and right external jugular vein (Figure 5C).
   4. Put 0.5 mL of PBS in the cavity to slow the development of vasospasm while manipulating the carotid artery.
   5. Carefully isolate a 5 mm section of the right carotid artery. Place a piece of sterile white paper underneath the vessel as a background to make the artery more visible (Figure 5D).
      NOTE: Carefully separate the vagus nerve (white) from the artery and ensure not to cut or damage the nerve or the artery.
   6. Using an 8-0 suture tie a permanent knot (#1) to close off the cranial end of the vessel (Figure 5E).
   7. Tie a first loose knot (#2) to temporarily occlude blood flow from the aorta. Then, tie a second loose knot (#3) between the first two sutures (Figure 5F). The second loose knot (#3) will be used to quickly secure the catheter after placement.
   8. Using a 25 G needle, make a small hole, large enough to pass the catheter, in line with the vessel between #3 and #1 ligatures (Figure 5G).
      NOTE: Carotid arteries carry oxygenated blood from the heart and have very high pressure. If the carotid artery is cut, that pressure will cause the blood to spurt out (Figure 5H).
   9. Hold the catheter 1.5 inches from the tip and gently insert the tip of the catheter through the hole of the artery (X-mark). Tighten the middle suture node (#3) around the catheter and vessel that still allows passage of the catheter (Figure 5I-J).
      NOTE: This step requires practice. The potential complications with this step include bleeding at the catheter insertion site and vasospasm. When bleeding occurs, blood loss from the bleeding artery reduces blood volume, leading to a severe drop in systemic blood pressure. Due to the severity, the animal has reached a humane endpoint and must be euthanized. For mechanically induced vasospasm, it usually occurs during catheter insertion caused by a persistent contraction of the blood vessels. This makes the blood vessel opening smaller and prevents catheter advancement to the carotid artery. Do not use excessive force against resistance to advance the catheter. When moderate or severe vasospasm resistance is encountered, try again in a little while or use a smaller catheter (e.g., 1.0 F). Experienced microsurgeons can achieve 100% success rates for ascending aorta catheterization.
   10. After the catheter passes the first loose knot (#2) with the sensor tip, fasten the second loose knot (#3) more tightly to secure the catheter and gently release the first loose knot (#2) (Figure 5K, L).
   11. Continue to insert the catheter toward the ascending aorta according to the mark on the catheter (Figure 4B) until pressure analysis shows an arterial blood pressure profile (Figure 5M). Record systemic blood pressure (SBP) data using the data acquisition system and software.
   12. Loosen the middle suture node (#3) to allow the catheter to be pulled out (Figure 5N).
   13. Tie the middle suture node (#3) around the vessel before pulling the catheter out of the carotid artery (Figure 5O-P).
   14. Place the catheter in PBS.
4. Right heart catheterization.
   1. Carefully isolate the right external jugular vein from the surrounding connective tissue and ligate all the small branches with 8-0 suture (blue arrowheads) (Figure 6A).
      NOTE: For right heart catheterization, the heart is commonly accessed via the right jugular vein.
   2. Using an 8-0 suture, tie a permanent knot (#1) to close off the cranial end of the vessel (Figure 6B). Then, tie a loose knot (#2) on the caudal end of the vessel (Figure 6C).
   3. Use a 25 G needle to make a small hole proximal to the permanent knot (#1) (Figure 6D).
      NOTE: Jugular veins carry deoxygenated blood to the heart and have low pressure. If the jugular vein is cut, the blood will not spurt out (Figure 6D, E).
   4. Hold the catheter and insert the catheter into the cut of the vein (X-mark) (Figure 6E) and tighten the caudal knot (#2) around the catheter and the vessel (Figure 6F).
   5. Slowly and gently push the catheter into the right heart. Monitor the catheter tip depth based on the catheter mark (Figure 4B).
      NOTE: Assessment of the right ventricular systolic pressure (RVSP) in closed-chest mice is a challenge because of the complex RV anatomy and structure. This step requires a high level of expertise and lots of practice. In the hands of an experienced microsurgeon, the successful rate for right ventricle catheterization can approach 90%.
   6. Assess the catheter tip position according to the pressure wave tracing in the software. When the tip of the catheter is in the right ventricle, the monitor will show a typical RVSP tracing (Figure 6G, H).
      NOTE: When the shape of pulmonary pressure curves looks atypical (e.g., spiky curves), this implies incorrect positioning of the catheter. Adjust the catheter position by gently pulling the catheter a little bit back, and then slowly advancing the catheter to a more central position within the right ventricle. To avoid the generation of artifacts in research data, the investigator should avoid prolonged (not more than 1 min) or repeated attempts (not more than two attempts) at right ventricle catheterization.
   7. Keep the catheter immobile and collect the data for 5 min.
   8. After the recording is complete, carefully pull the catheter out and tie the caudal knot (#2) around the vessel (Figure 6I). Place the catheter back in PBS solution.
      ​NOTE: After completion of the experiment, clean the catheter with 1% digestive enzyme solution according to the manufacturer's instructions. In addition to assessing hemodynamic status, investigators may harvest the hearts and the lungs for PAH histopathological examination. To ensure the effectiveness of multiple IV bolus dosing, investigators can isolate lung endothelial cells and measure let-7 miRNA levels.

4. Blood pressure data analysis

1. Examine the blood pressure recording.
   1. Open the blood pressure analysis software data file (PAH JOVE.adicht).
   2. In channel 1, select an area representing the pressure signal and place the Waveform Cursor on the Peak (X-mark) to measure pressure amplitude (Figure 7A).
   3. Determine the maximum amplitude of the pressure wave. This represents the systolic pressure (Figure 7A, red arrow).
   4. Extract the region of interest (Figure 7B gray area) from the image by pressing the Shift + Command + 3 (for Mac) or Windows + Shift + S (for Windows PC) and paste it into a graphics file.
2. Statistical analysis of blood pressure data.
   1. Enter the individual mouse blood pressure data in statistical analysis software.
   2. Perform an unpaired Student's t-test for statistical analysis of two study groups (normoxia vs. hypoxia; hypoxia vs. hypoxia + 7C1/let-7 miRNA). Consider the differences in mean values as significant as p < 0.05.

Results

Anesthesia often reduces blood pressure. Therefore, a minimum dose of anesthesia was used to abolish the movements in response to a noxious stimulus. Successful right ventricular chamber access can be visualized as the hemodynamic waveform changes in different regions of venous systems (Figure 8).

In this study, mice were randomly assigned to the normoxic (21% O₂) group (n = 10), hypoxia (10% O₂) group (n = 10), or hypoxia + 7C1/let-7 treatme...

Discussion

Several pulmonary hypertension animal models have been established to mimic the elevated pulmonary vascular resistance events in human subjects. Among them, the mouse hypoxia-induced PAH model has been widely used for evaluating the effectiveness of new experimental therapies for PAH. Research using this model often requires the administration of compounds to the mice. In comparison with other published intravenous (IV) injection and invasive hemodynamic assessment protocols, this method provides both visual illustration...

Materials

Name	Company	Catalog Number	Comments
5-0 prolene suture pack	Ethicon	8698G	for incision closure
8-0 nylon suture pack	AROSurgical Instruments	T06A08N14-13	for ligation
Anesthesia induction chamber	VETEQUIP	#941444	Holds the animal during anesthesia exposure
Catheter Interface Cable PEC-4D	Millar		for connecting Millar Mikro-Tip catheter to PCU-2000
Charcoal canister filters	VETEQUIP	#931401	to help remove waste anesthetic gases
Cotton swabs	McKesson	24-106	for applying pressure to the injection site to prevent bleeding
Fine scissors	Fine Science Tools	14059-11	Surgical tools
Insulin syringe 28 G	EXEL	26027	for jugular vein IV injection
Isoflurane	COVETRUS	#029405	for mouse anesthesia
LabChart 8 Software	ADInstruments		for data analysis
Mikro-Tip Pressure Catheter SPR-1000 (1.0 F)	Millar		for invasive blood pressure measurement
Needle-25 G	BD	305124	for making a samll hole in a vessel
Oxygen controller ProOx Oxygen Sensor	BioSpherix	E702	for oxygen concentration monitoring
PCU-2000 Pressure Control Unit	Millar		for connecting Millar Mikro-Tip catheter to PowerLab 4/35
PowerLab 4/35	ADInstruments		for Data Acquisition. Investigator needs to connect the PowerLab 4/35 to a personal laptop containing LabChart 8 software for operation.
Prism 8	GraphPad		for statistics and scientific graphing
Semisealable hypoxia chamber	BioSpherix		an artificial environment that simulates high-altitude conditions for animals
Spring Scissors	Fine Science Tools	15021-15	Surgical tools
Tweezer Style 4	Electron Microscopy Sciences	0302-4-PO	Surgical tools
VasoRx compound 7C1/let-7 miRNA	VasoRx, Inc.	Lot# B2-L-16Apr	IV injection compound
VIP 3000 Veterinary Vaporizer	COLONIAL MEDICAL SUPPLY CO., INC.		for accurate anesthesia delivery