Optical System for Assessment of Atrial and Ventricular Mechanics in Langendorff-perfused Mouse Hearts

Summary

This protocol describes the procedure for mouse heart extraction and cannulation, as well as the preparation and configuration of the Langendorff retrograde perfusion system. The experimental setup is coupled with a high-resolution optical system designed to evaluate the mechanical responses of the atria and ventricles while preserving their natural dynamics.

Abstract

The mouse heart serves as a widely used model in cardiovascular research, facilitated by advances in engineering technologies. However, assessment of atrial and ventricular contractility remains challenging due to the small size of the mouse heart. This study introduces a novel methodology for evaluating contractility and heart rate in an intact, isolated heart. Using a Langendorff-perfused mouse heart in combination with a high-resolution optical system enables evaluation of atrial and ventricular contractility. The experimental setup addresses challenges associated with small heart size and rapid contraction rates while acquiring high-resolution signals. Synchronization between stimulus frequency and heart contraction frequency in the range of 420 to 660 beats per minute (bpm) was consistent with results from the auxotonic method. Additionally, measurements of ventricular shortening and atrial dynamic signals allow for the determination of atrioventricular contraction. This method is suitable for assessing contractility and heart rate in an intact, isolated heart without pharmacological or systemic influences. These results demonstrate that the optical system enables measurement of ventricular and atrial shortening responses.

Introduction

There is much to discover in cardiac physiology; key processes, such as cardiac pace-making and excitation-contraction coupling, are central tenets that are continually researched¹. Although the effects of biological noise on cardiac contractility² and pace-making activity remain poorly understood¹, new optical imaging methods have allowed the presentation of new paradigms^3,4.

Two experimental methodologies using the Langendorff system have made invaluable contributions to measuring cardiac function. First, the isovolumic contraction method, where a signal is obtained from a small balloon inserted into the left ventricle^5,6,7. Second, the auxotonic recording method, where a signal is obtained by suturing one end of a thread to the apex of the heart and the other end to a mechanical recording device to measure auxotonic contractions along the axis of the heart^2,8. Nevertheless, evaluating atrial and ventricular contractility in the mouse heart is challenging due to its diminutive size. Furthermore, preparation requires a certain degree of gentleness, since the heart is susceptible to injury^{9,10,11,12,13}.

The optical system for assessing the mechanical dynamics of mouse hearts has several novel features and advantages. First, this method is suitable for assessing contractility and heart rate in an intact, isolated heart in the absence of pharmacological or systemic influences. The protocol could also be effectively employed in experiments assessing ischemia-reperfusion injury when looking into pharmacological, cell-regeneration, or oxygen-therapy-based treatments.

In this paper, we present a protocol for recording biophysical parameters of mouse heart contractile function using a high-resolution optical system. This method helps assess contractility and heart rate in an intact, isolated heart. Furthermore, it permits evaluation of atrial and ventricular contractility. The Langendorff mouse heart preparation presents a refinement that includes a slight modification to minimize loss of perfusate temperature and motion artifacts without inhibiting muscle contractions.

Protocol

All animal procedures were conducted in accordance with Federal Regulations for Animal Experimentation and Care and were approved by the Animal Care Committee of CINVESTAV (CINVESTAV Zacatenco, D.F., Mexico). In this study, C57BL/6J male mice are used, weighing 28–34 g, and aged 8–14 weeks. All materials and equipment used in the protocol are listed in the Table of Materials. Supplementary Files 1–11 provide supporting diagrams, software guides, analysis programs, pacing data, and application files for the protocol.

1. Stock preparation

1. Prepare two stock solutions in advance and store them in a refrigerator at 4 °C: (1) Stock I, 1 L (in deionized water; in g/L: 76.13 NaCl, 4.02 KCl, 1 MgCl₂(6H₂O), 0.4 NaH₂PO₄) and (2) Stock II, 0.1 L (in deionized water; in g/L: 73.5 CaCl₂(2H₂O)).
   NOTE: The solutions may be used for up to 30 days after preparation. Store them at 4 °C.

2. Langendorff system connection and cleanup

1. Connect the reservoir (1), reservoir thermal bath (2), perfusion pump (3), serpentine (4), serpentine recirculator (5), monitoring system (6), and drainage system (8), as shown in Figure 1A.
   NOTE: The connection between the monitoring system and the drainage system is shown in greater detail in Supplementary File 1.
2. Fill the Langendorff system with 1 L of deionized water at 50 °C, then start the perfusion and drainage pumps (Supplementary File 1, drainage system). After approximately 20–30 min, remove the liquid from the system, then proceed to the next step.
   NOTE: Verify that there are no leaks or pressure drops in the system. If this occurs, do not proceed with the experiment.
3. Fill the system with 1 L of 1% acetic acid (dissolved in deionized water), then turn off the perfusion and drainage pumps and wait for 60 min, then turn on the pumps to remove the liquid.
4. Repeat step 2.2 twice.
5. Fill the system with 70% ethanol (dissolved in deionized water) to prevent adhesion of solution components.
   NOTE: The Langendorff perfusion system must be cleaned every ten experiments or monthly. The pre-experiment cleaning protocol is explained in section 4. While the post-experiment cleaning protocol is detailed in section 8.

3. Preparation of Tyrode solution during the experiment

1. Prepare 1 L of solution in mM: 130.25 NaCl, 5.4 KCl, 0.53 MgCl₂(6H₂O), 0.33 NaH₂PO₄, 1.8 CaCl₂(2H₂O), 10 D-glucose, 10 HEPES, adjusted for pH 7.35 ± 0.05 by NaOH.
   1. Mix 100 mL Stock I with 700 mL of deionized water under constant stirring, then raise the temperature of the solution to 37 °C; maintain solution oxygenation by connecting the O₂ tank to the air stone as shown in Figure 1A (1).
   2. Add 3.6 mL of Stock II and 96.4 mL of deionized water to obtain 1.8 mM CaCl₂(2H₂O), then add NaOH to adjust the pH to 7.35 ± 0.05.
   3. Complete with deionized water to obtain 1 L of Tyrode solution.
      NOTE: It is recommended to store 100 mL of the solution in the refrigerator at 4 °C; this volume will be required later in step 6.

4. Langendorff system preparation

NOTE: If the Langendorff system is not filled with ethanol on the day of the experiment, it is not recommended to proceed with the procedure; instead, repeat section 2.

1. Turn on the perfusion and drainage pumps. Set the perfusion pump flow rate to 3 mL/min and the drainage pump to 10 mL/min. Drain the system.
2. Fill the system with 1 L of deionized water at 37 °C, then drain the water.
3. Fill the system with the Tyrode’s solution prepared in step 3 and maintain the solution at physiological temperature and constant oxygenation.
4. Turn on the reservoir thermal baths and the serpentine recirculator. Set the reservoir temperature (Figure 1A (2)) to 38 °C to maintain the solution under physiological conditions. Set the temperature of the serpentine (Figure 1A (5)) at 40 °C. This compensates for the loss of temperature in the Langendorff system solution.
   NOTE: If the temperature of the Tyrode solution at the system outlet is not physiological (37 ± 0.5 °C), adjust the serpentine recirculator temperature. It is not recommended to proceed to step 6 until the temperature has stabilized at the appropriate value.
5. Turn on the pressure monitor (Supplementary File 1, monitoring system) and calibrate in the range of 0–200 mmHg. The monitoring and pacing application is available to help calibrate the system (Supplementary File 3 and Supplementary File 11).
   NOTE: It is essential to verify that there is no pressure loss and no bubbles in the system. If the system contains bubbles, temporarily block the lower outlet of the serpentine and open the upper valve; this will cause the column to fill, allowing the bubbles to be released into the outside. It is not recommended to proceed with the experiment until this condition has been verified.

5. Optical system¹⁴

1. Assemble the CMOS camera (Figure 1A, 10), adjustable camera lens (LC), and illumination system.
   NOTE: The illumination system consists of two LEDs (Figure 1A, 9), two plano-convex illumination lenses (LI), an illumination control (Figure 1A, 11), an acquisition card (Supplementary File 1, monitoring system), and 3D-designed support pieces (Supplementary File 1).
2. Place a reference object with a volume of approximately 1,000 mm³ at 100–200 mm from the camera’s lens (center of Figure 1A).
   NOTE: The object must be positioned at the heart position in the Langendorff system. This position can be observed in Figure 1B, specifically in the center of the image where the light beams impinge.
3. Start the camera acquisition software, then set the camera with a subsampling of 4x (4 pixels = 1 subpixel). Set a region of interest (ROI) of 200 x 200 subpixels (Supplementary File 2).
   NOTE: With this configuration, a temporal resolution of 750 frames per second (fps) is possible.
4. Turn on the illumination system (Supplementary File 1), adjust the light beams to focus on the reference object, and achieve homogeneous illumination (Figure 1B).
5. Adjust the light intensity on the reference object to avoid overexposing of the image.
   NOTE: The image must look like Figure 2A (or Figure 1C, Controller PC). Reflections on the surface of the object (heart) must be reduced as much as possible without losing the definition of its contour. This can be done by adjusting the Illumination control power or by reducing the exposure time in the camera settings.
6. Focus the camera lens to obtain a clear image of the reference object (Figure 1C, controller PC). Ensure that the reference object occupies 80% of the field of view (Figure 2A) to obtain a conversion scale of approximately 61 µm per subpixel.
   NOTE: The optical system automatically activates the Illumination system when the camera is set to acquisition mode (Figure 1A and Supplementary File 2). It is recommended to keep the camera in standby mode so that the Illumination system remains off, activating only during experimental data collection periods to avoid overheating.

6. Surgical and cannulation procedures

1. Heparinization and anesthesia
   1. Prepare an anesthetic mixture containing tiletamine-zolazepam (40 mg/kg mouse body weight) and xylazine (10 mg/kg mouse body weight), then dilute to 200 µL with saline solution.
   2. Inject the anesthetic mixture intraperitoneally using a 27 G needle. Allow the mouse to enter deep anesthesia.
      NOTE: Verify the absence of a reflex in response to a stimulus in the limbs or tail (usually, 10 min is enough).
   3. Inject 200 UI of unfractionated heparin intraperitoneally using a 27 G needle gauge and wait approximately 5 min to continue with the experiment.
2. Extraction and cannulation
   1. Place the mouse in a supine position, remove the fur and skin from the thoracic area with surgical scissors.
   2. Make an abdominal incision below the diaphragm, making sure to maintain its integrity.
   3. Continue with a transverse incision in the diaphragm and quickly perform a bilateral ascending thoracotomy through the ribs, then remove the rib wall.
      NOTE: Taking care not to damage the heart. If the heart is compromised or damaged during the procedure, it is recommended that the experiment not proceed.
   4. Smoothly hold the superior structures of the heart using fine tweezers, the aortic arch, and adjacent vessels. Smoothly lift to make a cut underneath and upwards, and remove the heart and lungs together.
   5. Quickly transfer the heart into a petri dish containing 50 mL of Tyrode solution at 4 °C for an initial wash.
      NOTE: From the moment the diaphragm is incised, the animal will begin to experience respiratory distress; from this point forward, there is a 60 s window to perform steps 6.2.3–6.2.6. Exceeding this time limit may induce ischemic damage.
   6. Remove lungs and excess adipose tissue to expose the aortic arch using a stereoscopic microscope and fine scissors.
      NOTE: When cleaning the tissue, ensure the aortic arch is not damaged. If the aorta is cut below the brachiocephalic artery, cannulation will be difficult, potentially leading to pressure leaks.
   7. Transfer the heart into the custom-made cannulation base containing 50 mL of Tyrode buffer at 4 °C and insert the aortic extremity into the cannula custom-made from a 20 G needle (Supplementary File 1).
      NOTE: Be careful not to damage the aortic valve. If the valve is compromised, the heart swells, and the coronary circulation will not properly flush.
   8. Tie two cannulation knots using 4/0 silk suture: the first in the indentation distal to the tip of the cannula and the second before the brachiocephalic trunk (Supplementary File 1).
   9. Slowly apply Tyrode solution using a 1 mL syringe filled with buffer at room temperature. Verify that the coronary tissue clears to confirm good cannulation.
   10. Remove adipose tissue to allow access to the ventricles and atria.
       NOTE: Avoid exceeding 5 min of cannulation to preserve heart viability.
   11. Transfer the heart into a Langendorff system and allow it to stabilize for 30 min.
       NOTE: The heart is placed in the center of the system (Figure 1A,B), suspended vertically. When hanging the heart, take extreme care to prevent air bubbles from forming. If this occurs, the pressure will increase above 140 mmHg, the heart will begin to swell, and it will no longer be viable.
       NOTE: The Langendorff montage is complete when the heart exhibits a mechanically stable response, a natural oscillation frequency of approximately 360 beats per minute (BPM), and a perfusion pressure between 60 and 110 mmHg.

7. Pacing protocol and video recording

1. Heart viability
   1. Verify that the coronary pressure is in the range 60–110 mmHg with a 3 mL/min perfusion pump. If the coronary perfusion pressure drops below 60 mmHg, slightly increase the perfusion pump flow rate (low pressure will cause the heart to become pale and flaccid). Conversely, if the pressure exceeds 110 mmHg, slightly decrease the pump flow rate (excessive pressure will induce tissue edema and a progressive increase in pressure until cardiac arrest occurs).}
      NOTE: Maintaining the perfusion pressure outside the aforementioned optimal range for an extended period will irreversibly compromise heart viability.
   2. Place the heart in a frontal position so that a full-frontal view is obtained in the camera. Adjust the position so that both atria are visible in the camera (Figure 2A).
   3. Gently place the platinum electrodes superficially on the top of the right atrium at the pacemaker node. Use the camera feed as a guide to bring the electrodes into the field of view. Position the electrode so that it does not obstruct the right atrium. While monitoring the camera live view, slowly advance the electrode until it contacts the atrial tissue, taking care not to exert excessive compression (which would flatten the right atrium in the image).
      NOTE: If performed correctly, the camera view will resemble Figure 2A.
   4. Launch the monitoring and pacing application (Supplementary File 11) and turn on the acquisition card (Figure 1A (6)). Set stimulus parameter values: amplitude at 2 V, frequency at 420, 540, and 720 pulses per minute (PPM) with a 7 ms duty-cycle square pulse (inset of Figure 2A).
   5. Apply the viability stimulations starting with the lowest frequency, using a stimulation duration of 20 s followed by a 20 s recovery period (resting time) between events.
      NOTE: Assess heart viability via camera monitoring. If the organ is physiologically functional, a synchronization between the applied stimulus and the cardiac response will be observed, accompanied by proper coordination between the heart chambers. Conversely, a lack of viability will be manifested by a loss of the chamber’s coordination or by the exclusive presence of isolated atrial contractions.
   6. Close the chamber door because the experiment is sensitive to the room light and thermal fluctuations (Figure 1C).
      NOTE: The chamber door must remain closed during experimental time. Monitor the heart viability with the camera and perfusion pressure.
2. Data recording
   1. Apply the pacing protocol, which consists of square pulse trains of 30 s duration, followed by 30 s without a signal. Set stimulus parameter values in each pacing train: amplitude at 2 V, 7 ms duty cycle, and frequency in the range of 360 to 720 PPM.
      NOTE: Pacing protocol begins at 360 PPM and increases by 60 PPM at each time point (inset of Figure 2A).
   2. Continuously record videos of the dynamics of the heart’s frontal view in .avi format using camera acquisition software (Supplementary File 2). Make a 20 s video recording of each pacing train. Record the 20 s segment beginning 5 s after pacing. A representative raw recording is provided in Supplementary Movie 1.
      NOTE: A computer simultaneously records the pacing train applied (.xls, Supplementary File 9) to each recorded video (.avi), generated by the monitoring and pacing application and the camera acquisition software, respectively (Supplementary File 2 and Supplementary File 3).
   3. Verify heart viability by comparing the first video, recorded before starting the stimulation protocol, with the second video, recorded after applying the protocol (Figure 2B).

8. Langendorff perfusion system post-experimental clean-up

1. Remove stimulus and ECG platinum electrodes. Remove the heart from the system and drain all residual Tyrode solutions.
2. Turn off the optical system and close the O₂ valve.
3. Rinse the system with 1 L of deionized water at 37 °C, and drain the system.
4. Repeat steps 2.4 and 2.5.
5. Turn off all systems and equipment.

9. Video processing

1. Begin by importing each video into computational software and applying a video normalization function¹⁵.
2. Select a region of interest (ROI) that includes the heart area. Perform image segmentation to remove background noise, electrodes, and the cannula (Supplementary File 4).
   NOTE: To correctly select the image area to be removed, please refer to Supplementary File 10.
3. Conduct a global mechanical dynamics analysis (Supplementary File 5). Supporting variability and signal examples are shown in Supplementary Figure S1, Supplementary Figure S2, and Supplementary Figure S3.
   1. Apply morphological operators to each video frame to detect the heart perimeter, then apply a fill frame algorithm to generate a mask¹⁵.
   2. Count the pixels within the segmented area in each frame, then normalize this count by the total number of pixels in the image to produce a dynamic signal (inset in Figure 2B and Supplementary Figure S3).
   3. Apply a 5th-order bidirectional Butterworth filter between 3 and 100 Hz to the obtained signal to suppress high-frequency noise and perfusion dripping artifacts¹⁵.
   4. Normalize the signals across all videos using the recorded maximum and minimum values.
   5. Determine the heart rate by calculating the interval between two maximum-amplitude values (IAA (s)) for each dynamical and stimulus signal.
   6. Obtain the frequency in BPM with the next formula:
      
   7. Generate Poincaré maps from IAA data to illustrate variability over short and long-term periods^16,17,18 (Supplementary Figure S1 and Supplementary Figure S2).
      NOTE: For more detailed information, please refer to Supplementary File 10.
4. Conduct a segmental analysis of the ventricular mechanical dynamics (Supplementary File 6).
   1. Apply image binarization. Use morphological operators to delimit the ventricular area, adjusting it to an elliptical geometry without compromising the ROI¹⁵.
   2. Fit a minimum ellipse to the ventricular area in each frame. Extract the coordinates of a set of distal points in a frame. Use a set of 10–20 points in each extreme along the Y axis (Figure 3A). A representative ellipse-estimation output is provided in Supplementary Movie 2.
   3. Calculate the ventricular shortening as the mean distance between extreme points along the Y axis in each video¹⁹.
   4. Apply a 5th-order bidirectional Butterworth filter between 3 and 100 Hz to the obtained signal to suppress high-frequency noise and perfusion dripping artifacts¹⁵.
   5. Normalize the ventricular shortening signal relative to all videos obtained for each pacing, and calculate heart rate and delay as in steps 9.3.5–9.3.6.
      NOTE: Parameters, such as ventricular cycle duration (DVE), relaxation duration (DVW), interventricular interval (IVV), and ventricular amplitude (VA), are obtained (Figure 3A).
      NOTE: For more detailed information, please refer to Supplementary File 10.
5. Conduct a segmental analysis of the atria’s mechanical dynamics (Supplementary File 7).
   1. Use two ROIs to delimit the atria (Figure 3C).
   2. Analyze each atrium individually.
      1. Use morphological operators to delineate atria while adjusting them without compromising the ROI¹⁵. Use a contrast-increase algorithm to delineate each atrium^20,21.
      2. Apply image binarization and quantify the number of pixels that differ from zero per frame to obtain an atrial dynamic signal.
      3. Apply a 5th-order bidirectional Butterworth filter between 3 and 100 Hz to the obtained signals to suppress high-frequency noise and perfusion dripping artifacts¹⁵.
      4. Calculate the average of two atrial signals (Figure 3D).
   3. Normalize the atrial shortening signal relative to all videos obtained for each pacing and calculate heart rate and delay as in steps 9.3.5–9.3.6.
      NOTE: Parameters, such as atrial cycle duration (DAE), atrial-atrial interval (IAT), and atrial amplitude (AA), are obtained (Figure 3D). For more detailed information, refer to Supplementary File 10.
6. Conduct an electro-mechanical analysis^13,22 (Supplementary File 8).
   1. Compare the pacing data with the atrial and ventricular signals (Figure 3D).
   2. Calculate the propagation time in the heart conduction system, obtaining the delay between the atrium and ventricular events (AVD), taking the applied stimulus as a reference.

Results

The cardiac mechanical function experiments are performed using the Langendorff apparatus, as shown in Figure 1A (Top). The monitoring and optical systems enable video recording when illumination control is triggered, as shown in Figure 1A (Bottom). Figure 1C illustrates the external view of the thermostatically controlled cabinet, designed to maintain the heart isolated from ambient thermal fluctua...

Discussion

The mouse heart is very sensitive to temperature variations^2,9,10,11,22. In this experiment, the perfused heart was vertically positioned. We included a slight modification to minimize loss of perfusate temperature and motion artifacts in the Langendorff perfusion method. A recirculation pump was added to the serpentine to minimize loss of perfusate temperatur...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1 mL syringe	BD Plastipak	302485	Surgical procedure
20 G needle	NIPRO	AH-2032	Aortic cannula
25 mm adjustable lens with 8x-100x magnification	Generic	N/A	Optical system camera lens; cited generically as adjustable camera lens
27 G needle	BD PrecisionGlide	302358	Surgical procedure
3D-designed support pieces	CINVESTAV	N/A	Custom 3D-designed support pieces
4-0 silk suture	AMERICAN SUTURE	TGSN0901	Aortic cannula
40-A power supply	STEREN	PRL-25	LED power supply
Air stone	N/A	N/A	Tyrode's solution oxygenation
CaCl2 (2H2O)	Sigma-Aldrich	C3306	Tyrode's solution
D-Glucose	J.T. Baker	1916-01	Tyrode's solution
Drainage pump	CINVESTAV	N/A	Custom drainage pump; cited generically as drainage pump
F = 25.4 mm, plano-convex lens	THORLABS	LA1951-A	Optical system illumination lens; cited generically as plano-convex illumination lens
Glacial acetic acid	Sigma-Aldrich	695092	Langendorff system cleaning
Heparin	PiSA	060M91	Anticoagulant
HEPES	Sigma-Aldrich	H3375	Tyrode's solution
High-power LEDs	Luminius	CBT-90-G-L11-C12	LED light source
Illumination system	CINVESTAV	N/A	Custom illumination system; cited generically as illumination system
KCl	CTR SCIENTIFIC	CTR02196	Tyrode's solution
LabVIEW software version 2018	NATIONAL INSTRUMENTS	N/A	Software used to run the monitoring and pacing application; cited generically as monitoring and pacing application
MATLAB software	The MathWorks	N/A	Computational software for video normalization, segmentation, and analysis programs; cited generically as computational software
Medical-grade oxygen	GRUPO INFRA	N/A	Tyrode's solution oxygenation
MgCl2 (6H2O)	CTR SCIENTIFIC	CTR02104	Tyrode's solution
Mini-Pump variable flow	CONTROL COMPANY	3385	Perfusion pump
Mouse Surgical Kit	Kent Scientific	INSMOUSEKIT	Surgical procedure
NaCl	Sigma-Aldrich	S5886	Tyrode's solution
NaH2PO4	DEQ	DEQF290400250	Tyrode's solution
NaOH	CTR SCIENTIFIC	CTR03108	pH adjustment
NI USB-6009	NATIONAL INSTRUMENTS	779026-01	Acquisition card
Platinum electrodes, diam. 0.25 mm	Sigma-Aldrich	349402	Electrical stimulation electrodes
Power supply (12 V, 30 A)	Karnick	PTA-12V30A	Recirculator power supply
Pressure monitor	CINVESTAV	N/A	Custom pressure monitor; cited generically as pressure monitor
Pressure transducer	LA BOUVET	B17295	Langendorff system pressure monitor
Pure ethyl alcohol	Sigma-Aldrich	493511	Ethanol for system cleaning
Recirculating thermal bath (LXC)	PolyScience	L113A0412	Reservoir recirculator
Saline solution	PiSA	111848	0.9% sodium chloride (NaCl) solution
Serpentine thermal bath	CINVESTAV	N/A	Custom serpentine recirculator/thermal bath
Stereomicroscope	N/A	N/A	Surgical procedure
Stereomicroscope illumination	N/A	N/A	Surgical procedure
uEye Cockpit software	IDS Imaging Development System GmBH	N/A	Camera acquisition software; cited generically as camera acquisition software
UI-3360CP-NIR-GL Rev.2	IDS	AB00624	High-speed CMOS camera; cited generically as CMOS camera
Xylazine	PiSA	4001211	Anesthetic
Zoletil 100	Virbac	83048907	Tiletamine-zolazepam anesthetic; cited generically as anesthetic mixture