All methods and procedures were performed in accordance and compliance with all relevant regulations ('European Convention for the Protection of Vertebrate Animals used for Experimental and other Scientific Purposes' (Directive 2010/63/EU) and animal care was in accordance with institutional guidelines. Data from human subjects were analyzed in compliance with all institutional, national, and international guidelines for human welfare and was approved by the Local Ethics Committee (20-9218-BO). All experiments have been performed with male C57BL/6JRj at the age of 12 weeks.

1. Preparation of materials and equipment

NOTE: Figure 1 shows an example of a small-animal ultrasound workplace.

1. Make sure to operate in a silent controlled environment with dimmable light.
2. Pre-heat the ultrasound gel, e.g., using a gel warmer. Allow the gel to warm to 37 °C. This may take a while.
3. Clean all instruments including the platform with a disinfectant wipe.
4. Turn on and pre-heat the platform to 37 °C.
5. Turn on the ultrasound machine. Enter the animal ID and protocol ID as well as other relevant information. Use a high-frequency ultrasound transducer with a center transmit of 30 MHz for mice with approximately 30 g body weight.
6. Make sure to work with an active gas exhaustion system.
   NOTE: If using an activated charcoal filter for adsorbing the isoflurane in the exhaled flow, make sure to check the weight and replace the filter once the indicated maximal weight increase is reached.
7. If necessary, fill the vaporizer with the adequate amount of isoflurane.
   CAUTION: Do not inhale volatile anesthetics.
8. Prepare a dobutamine working solution of 2.5 µg/µL either by dilution of a ready-to-use injection solution or by dissolving dobutamine hydrochloride powder in 0.9% saline according to the manufacturer's instruction. The solution is stable at least for 24 h when stored at room temperature.

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Figure 1: Small-animal cardiac ultrasound workplace. An ergonomic setting is indispensable for small-animal stress echocardiography as examination times must remain short. The workplace consists of an ultrasound machine, a small-animal anesthesia system with oxygen supply and active gas exhaust, a heated echocardiography platform with embedded ECG and movement capabilities via micromanipulators as part of an integrated rail system as well as a physiological monitoring unit. A gel warmer to warm ultrasound gel and a heat lamp are useful aids. Please click here to view a larger version of this figure.

2. Preparation of the mouse for imaging and induction of anesthesia

1. Fill the induction chamber with 3%-4% v/v isoflurane in an oxygen-enriched gas mixture (with 1 L/min 100% O₂).
2. Weigh the mouse. Gently pick up the mouse by the tail and transfer it to the induction chamber. Ensure that the animal is sedated within a few seconds by closely observing the animal's movements.
3. If necessary, change the gas flow to the nose cone connected to the anesthesia system (1.0-1.5 vol% isoflurane with 1 L/min 100 % O₂ to maintain a stable sedation). Remove the mouse from the induction chamber and place it carefully on the pre-heated platform. Ensure that the paws lie on the ECG sensors embedded in the platform.
4. To prevent drying of the sclera, apply ointment gel to both eyes.
   NOTE: Stress measurements will take their time.
5. Apply a very small amount of electrode cream to the ECG sensors. Gently secure the animal with adhesive tape on all four limbs. Use a small part of adhesive tape to secure the animal's head position in the nose cone. The ECG is used to record the heart rate during image acquisition. Adjust the physiological imaging system for a stable and clear ECG signal.
   NOTE: Too much electrode cream may result in bad ECG signal quality.
6. To protect the animal from stress during the procedure, check the adequate depth of sedation by maintaining the heart rate range at 400-450 bpm. The heart rate is obtained by the ECG. A variation of 50 bpm within the range is acceptable.
   NOTE: Movements of the animal may indicate a too narrow level of sedation. The anesthesia must not lead to cardio-depression of the mouse. The sedation can be adjusted to obtain the above-mentioned target heart rate.
7. Using lubrication, gently insert a rectal thermometer for continuous monitoring of the body temperature. Keep the temperature in the physiological range (normally between 36.5 °C and 37.5 °C depending on the mouse strain and experimental setup). In a non-environmentally controlled animal cardiac ultrasound laboratory, the use of infrared lighting may be considered.
8. Use chemical depilatory cream to remove the body hair from the chest. Use a clean damp paper towel to wipe clean the chest. Make sure to remove all remaining cream components (Figure 2A).
   NOTE: An electric clipper can be used before depilatory application as well. The animal is now prepared for imaging. As it is critical to keep imaging time short, the whole preparation before imaging should take less than 3 min.

Figure 2: Animal and transducer positioning. (A) The mouse is attached to the heated platform with all four limbs fixed on the silver ECG electrodes. A rectal thermometer is inserted for body temperature measurement. The snout is gently inserted to the nose cone of the anesthesia system. (B) Probe orientation for parasternal long axis view (PSLAX); see step 3.2. (C) Probe orientation for parasternal short axis view (PSSAX); see step 3.3. (D) Probe orientation for apical four chamber view (4CH); see step 3.4. Please click here to view a larger version of this figure.

3. Basic cardiovascular imaging

NOTE: Images can be acquired using two basic transducer positions (parasternal and apical ultrasound window) (Figure 2) and at least three ultrasound modalities (B(rightness)-mode, M(otion)-mode and Doppler-mode (color doppler and pulsed-wave (PW) doppler) (Figure 3,Figure 4,Figure 5). For basics of imaging please refer to previously published articles^16,18. It is critical to obtain clear images for the comparison with later acquired stress images.

1. Apply pre-heated ultrasound gel bubble-free to the chest.
   NOTE: Non-heated ultrasound gel will result in rapid body temperature loss which will affect the heart rate.
2. Perform parasternal long axis (PSLAX) view.
   NOTE: PSLAX is performed to visualize the LV in its long axis. With this, e.g., the aortic root dimensions and proximal aorta dimension as well as the LV length can be obtained.
   1. With the head facing away from the investigator, tilt the table approximately 10-20° to the left and 5-10° to the front to bring the heart as anterior as possible. Place the transducer parasternally in line with the long axis of the heart with the marker (notch) pointing towards the animal's right shoulder (Figure 2B).
   2. Use micromanipulators to adjust the optimal view. Use image control panel controls to optimize the image. Acquire at least one 2D B-mode picture and one M-mode picture in midventricular level.
   3. Acquire any additional images if need for the specific question. Acquire at least 100 frames and at least 3(-6) full cardiac cycles.
3. Perform parasternal short axis (PSSAX) view.
   NOTE: PSSAX is performed to visualize the LV in its short axis. From this view, e.g., left-ventricular end-systolic volume (LVESV), left-ventricular end-diastolic volume (LVEDV), stroke volume (SV) and cardiac output (CO) can be calculated.
   1. Rotate the transducer 90° clockwise without changing of the angulation (marker now pointing towards the animal's left shoulder) (Figure 2C). Acquire at least one B-mode image in the basal, midventricular (level of the papillary muscles), and apical view.
   2. To define the most basal and the most apical view, scroll along the long axis to the most distant points, where the full cardiac cycle of the LV chamber is still visible. Take images on the midventricular level approximately in an intermediate position at the level of the papillary muscles.
   3. Acquire at least one M-mode image in the midventricular view.
      NOTE: Some ultrasound machines provide presets for the different views; it is recommended to check for the adequate preset before acquiring images.
4. Perform apical four chamber (4CH) view.
   NOTE: 4CH is important because it can be primarily used to evaluate the mitral valve using PW doppler.
   1. Tilt the platform that the animal is in a modified Trendelenburg position with the head down. Angle the transducer towards the head of the mouse, the marker facing towards the animal's left side (Figure 2D).
   2. Acquire at least one B-mode image as well as color doppler and PW doppler image of the mitral- and tricuspid valve. Depending on the experimental question, apply tissue doppler in the 4CH view.
      NOTE: The easiest way to reach apical 4CH position is to tilt the table from the PSAX view and to angulate the transducer. Be careful not to apply too much pressure to the thorax as this may interfere with measurements e.g. of diastolic function.

4. Dobutamine stress imaging

NOTE: Once the target heart rate is reached, standardized views should be acquired as long as the target heart rate is stable. This typically requires more than one switch between PSLAX and PSSAX. Because the switch between PSLAX and PSSAX only requires a 90° rotation, the views can be imaged easily.

1. Perform dobutamine stress testing in one and the same animal maintaining the same anesthesia to ensure comparability. Make sure the starting heart rate remains stable in the range of 400-450 bpm. Record the ECG readings and save it together with and on the acquired images. Make sure the ECG signal is clear. Otherwise, try to re-tape all four limbs until a clear ECG signal is displayed.
2. Again, perform PSLAX view (B-mode and M-mode images). Save the images as "baseline" image. Be sure, to save and keep in mind the initial heart rate as well.
3. Prefill the syringe and inject 5 µg/g body weight dobutamine intraperitoneally using a 27 G needle and 1 mL syringe. Closely watch for the heart rate. Record echocardiographic images until the target heart rate is reached and use the increase of the heart rate for later analysis. A sustainable significant dobutamine-induced heart rate increase is reached after an increase of 15-30% after about 1 min, depending on the dobutamine dose.
   NOTE: Always use single-use sterile injection needles for every animal to prevent from infections. The dobutamine susceptibility and the (sub)maximal load may vary with the mouse strain and may be dependent on the experimental setup and should be defined in pre-experiments. It is recommended adjusting the dobutamine dose to the experimental setup.
   CAUTION: Follow institutional guidelines for the use of sharp and potential infectious items. Always dispose the needle into an approved medical waste container!
4. Once the target heart rate is reached and remains stable for approximately 30 s, acquire PSLAX B-mode and M-mode images as described in step 3.
5. Again, rotate the transducer clockwise to obtain PSSAX view as described in step 3. Here, acquire B-mode images of the basal, midventricular (level of papillary muscles) and apical level and M-mode images of the midventricular (level of the papillary muscles) level. Reassure, that the target heart rate remains stable. Otherwise, switch back to PSLAX position and start imaging again.
   NOTE: As the heart rate will drop without continuous infusion of dobutamine¹² (not covered in this article), the images should be acquired within two minutes. PSLAX and PSSAX images are essential for most of the relevant stress-induced measurements (see "Representative Results" section).
6. Now, perform apical 4CH view again (as explained in step 3.4.). Using PW doppler, measure the flow patterns of interest (as explained in step 3.4.2.). Under unstressed conditions, two characteristic waves are measured using PW doppler, one representing the passive filling of the ventricle (E(arly) wave) and one representing the active filling after atrial contraction (A(trial) wave). With increasing heart rates, these waves tend to fuse and may not be measurable distinctively under dobutamine induced stress.

5. Final steps

1. After approximately 5 min when the heart rate will start to decrease again, make sure all views are captured.
2. Gently remove the ultrasound gel from the chest using a clean damp paper towel. Carefully remove the tape fixation. Pay special attention to the tape fixing the animal's head to avoid pulling out the whiskers.
3. Turn off the anesthesia. If using an active gas exhaust, make sure to continue gas exhaustion. Place the animal on a paper towel in a separated heated cage during the wake-up period. Observe the animal closely. It must not be left unattended until it has regained sufficient consciousness to maintain sternal recumbency. Once awake and fully recovered, transfer the animal to its cage.
   ​NOTE: Due to the non-final character of this technique, the animal may stay within the experiment in accordance with all relevant regulations.

6. Offline evaluation

1. Transfer the image data to the offline analysis software on a working-station to perform detailed evaluation of the cardiac function. Pay special attention to the difference between unstressed and stressed heart function. The heart rate should always be recorded and presented.
   NOTE: Because software analysis varies between different software, it is not covered in this protocol. Please refer to the manufacturer's instructions.