Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Summary

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Abstract

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

Introduction

Animal models have been a valuable tool used for understanding the underlying mechanisms of human physiology and pathophysiology, as well as a platform for preliminary testing of therapies to treat various diseases¹. Great strides have been taken in the field of biomedical research based on these animal studies². However, significant interspecies differences exist between human and animal physiologies, including mice, rats, guinea pigs, rabbits, sheep, pigs, and dogs^3,4. As a result, there have been numerous drug, gene, and cell therapies that showed promise during the animal testing stage but failed to live up to the results in clinical trials⁵. To bridge this gap, isolated cardiac myocytes and human induced pluripotent stem cells (iPSCs) were developed as models to test the response of human physiology to various drugs and diseases⁶. Stem cell-derived cardiomyocytes have been widely used in organ-on-a-chip systems as a surrogate of the heart^6,7,8. However, the usefulness of iPSC-derived cardiomyocytes (iPSC-CMs) is impeded by their relatively immature phenotype and the lack of representation of the cardiomyocyte subpopulation; the mature myocardium is a complex structure comprised of several coexisting cell types such as fibroblasts, neurons, macrophages, and endothelial cells. On the other hand, isolated human cardiomyocytes are electrically mature, and different cardiomyocyte subpopulations can be obtained by altering culturing parameters⁹. Still, these myocytes generally exhibit altered action potential morphologies due to the lack of cell-cell coupling, rapid de-differentiation, and occurrence of proarrhythmic behavior in vitro^10,11. Some of the limitations were addressed by 3D cell culture models of iPSC-CMs and cardiac myocytes. These models, which include spheroids, hydrogel scaffold encapsulated 3D cultures, engineered heart tissues (EHTs), and heart-on-a-chip systems, use multiple cardiac cell populations such as cardiomyocytes, fibroblasts, and endothelial cells. They either self-assemble or assemble along a scaffold to form 3D structures, and some even reproduce the complex anisotropic nature of the myocardium. These models have been reported to have cells of mature phenotypes, contractile properties, and molecular profiles similar to cardiac tissue. The heart-on-a-chip system also allows the study of systemic effects in drug testing and disease models. However, in vitro cell-based models lack the native extracellular matrix and therefore cannot accurately mimic organ level electrophysiology. Human cardiac slices, by contrast, have an intact extracellular matrix and native cell-to-cell contacts, making them useful for more accurately examining arrhythmogenic properties of the human myocardium.

Researchers have developed human cardiac organotypic slices as a physiological preclinical platform for acute and chronic drug testing and to study cardiac electrophysiology and cardiac disease progression^{12,13,14,15,16,17,18,19}. When compared with iPSC-derived cardiomyocytes, human cardiac slices more faithfully replicate adult human cardiac electrophysiology with a mature cardiomyocyte phenotype. When compared with isolated human cardiomyocytes, cardiac slices exhibit physiological action potential durations because of the well-preserved cell-cell coupling and the intrinsic existence of their native intra- and extracellular environments.

This protocol describes the process of generating human cardiac slices from whole donor hearts, performing acute (i.e., hours-long) and chronic (i.e., days-long) studies to test cardiac electrophysiology parameters via optical mapping. While this protocol describes only the use of the left ventricular (LV) tissue, it has been successfully applied to other regions of the heart as well as other species such as mice, rats, guinea pigs, and pigs^14,20,21,22. Our laboratory uses whole human donor hearts that have been rejected for transplantation for the last 5 years, but it is feasible for these same procedures to be carried out on any donor heart sample tissues obtained by alternative means (e.g., left ventricular assist device [LVAD] implantations, biopsies, myectomies) as long as the tissues have the ability to be sectioned into cubes. Optical mapping is employed for analysis in this study due to its capacity to simultaneously map optical action potentials and calcium transients with high spatial (100 x 100 pixels) and temporal (>1,000 frames/s) resolution. Alternative methods can also be used, such as multielectrode arrays (MEAs) or microelectrodes, but these techniques are limited by their relatively low spatial resolutions. Additionally, MEAs were designed for use with cell cultures, and sharp microelectrodes are more easily managed for use with whole hearts or large tissue wedges.

The goal of the article is to enable more researchers to use human cardiac tissues for cardiac electrophysiology studies. It should be noted that the technology described in this article is relatively simple and beneficial for short-term studies (on the order of several hours to days). More physiological biomimetic culture for longer-term studies (on the order of weeks) has been discussed and described by a number of other studies^12,18,23. Electrical stimulation, mechanical loading, and tissue stretching are advantageous conditioning mechanisms that can help limit the onset of in vitro tissue remodelling^12,18,23.

Protocol

All methods described have been performed in compliance with all institutional, national, and international guidelines for human welfare. Research was approved by the Institution Review Board (IRB) at The George Washington University.

NOTE: Donor human hearts were acquired from Washington Regional Transplant Community as deidentified discarded tissue with approval from the George Washington University IRB. Explanted hearts are cardioplegically arrested by flushing the heart with a solution of ice-cold cardioplegia (the blood was cleared from the heart in this process) and transferred to the lab under standard organ transplant conditions.

1. Preparation of solutions

1. For each heart, make 4 L of cardioplegic solution (110 mM NaCl, 16 mM KCl, 16 mM MgCl₂, 10 mM NaHCO₃, 1.2 mM CaCl₂; pH = 7.4). Store 3 L at 4 °C and the remaining 1 L at -20 °C.
   NOTE: This solution can be made up to several days in advance.
2. Freshly prepare 1 L each of Tyrode’s slicing solution (140 mM NaCl, 6 mM KCl, 1 mM MgCl₂, 1.8 mM CaCl₂, 10 mM glucose, 10 mM HEPES, 10 mM 2,3-butanedione monoxime [BDM]; pH = 7.4) and Tyrode’s recovery solution (140 mM NaCl, 4.5 mM KCl, 1 mM MgCl₂, 1.8 mM CaCl₂, 10 mM glucose, 10 mM HEPES, 10 mM BDM; pH = 7.4).
   NOTE: Both the slicing and recovery solutions should be made the day of the experiment and can be stored at 4 °C.
3. Prepare stock solutions of the fluorescent dyes. Reconstitute the voltage-sensitive dye RH237 at 1.25 mg/mL in dimethyl sulfoxide (DMSO) and store it in 30 µL aliquots at 4 °C. Reconstitute the calcium indicator Rhod-2AM at 1 mg/mL in DMSO. Store in 30 µL aliquots at -20 °C.
   NOTE: Di-4-ANEPPS (stock solution at 1.25 mg/mL in DMSO) can be used in experiments for single camera imaging of the transmembrane potential alone.
   1. Before the start of the experiment, sonicate the dyes using an ultrasound sonicator for at least 10 min and dilute each aliquot of fluorescent dyes in 1 mL of the recovery solution. Add Pluronic F-127 (Table of Materials) to Rhod-2AM at a 1:1 ratio before dilution in recovery solution.
4. Prepare a stock solution of the excitation-contraction uncoupler blebbistatin at 2 mg/mL solution in DMSO. Store in aliquots at -20 °C. During optical mapping experiments, dilute the stock solution of blebbistatin to a working concentration of 5−10 μM in the Tyrode’s recovery solution.
5. Make fresh culture medium by supplementing medium 199 with 2% penicillin-streptomycin, 1x insulin-transferrin-selenium (ITS) liquid media supplement, and 10 mM BDM. Filter the medium using a 0.2 µm sterile filter.
   NOTE: For pharmacological perturbation studies, drugs can be added directly to the culture medium. The culture medium can be stored at 37 °C.
6. Make a 4% agarose gel for tissue mounting by dissolving low-melting point agarose in distilled water and heating the mixture in a microwave until fully dissolved. Cure the agarose in a Petri dish at a thickness of 5 mm and store at 4 °C.

2. Equipment setup

1. Vibrating microtome setup
   1. Calibrate the vibrating microtome prior to each experiment.
      1. Using a high precision vibrating microtome (Table of Materials), load a ceramic cutting blade into the holder and attach the calibrating device provided with the vibrating microtome. Choose the blade adjustment option from the menu and select ceramic for type of blade.
      2. Select the vibrate option, check for Z axis value. If this value is <1 μm, exit the calibration menu. If not, finely adjust the calibration screw attached to the top of the vibrating head and select the vibrate option. Repeat as many times as needed to set the Z axis to <1 μm.
   2. Set the vibratome settings to 400 μm cutting thickness, 0.02 mm/s advance speed for atrial tissue and 0.04 mm/s advance speed for ventricular tissue, 2 mm horizontal vibration amplitude, and 80 Hz vibration frequency.
      NOTE: While 400 μm is the recommended thickness to compensate for cell damage on the cut surfaces of the slice, thinner slices can also be prepared. Given that the oxygen diffusion limit is around 150 μm, slices around 300 μm are often used^14,17,18,24.
   3. Fill the bath of the vibrating microtome with slicing solution at 4 °C and maintain the temperature by surrounding the outside of the bath with ice, replenishing as needed throughout the slicing protocol. Continuously oxygenate the slicing solution in the bath by bubbling with 100% oxygen during slicing.
   4. Set up a second dish with as many 100 μm nylon mesh cell strainers and meshed washers as needed (one cell strainer per slice). Fill this dish with recovery solution and oxygenate it by bubbling with 100% oxygen at room temperature (RT).
      NOTE: This solution is maintained at RT during the experiment.
2. Optical mapping setup
   NOTE: A more detailed description of the optical mapping system is provided in previous publications^16,25,26.
   1. Attach a tissue bath with polydimethylsiloxane (PDMS) gel layer at the bottom (to pin the slices) to a perfusion system. Circulate 1 L of the recovery solution at 37 °C and oxygenate with 100% oxygen, through the perfusion system at a flow rate fast enough to maintain the temperature and clear accumulation of bath perfusate.
   2. Adjust the focus and alignment of the two CMOS cameras (Table of Materials) using a target.
      NOTE: More details on alignment can be found in previous studies²⁶.
   3. Use a green LED light source with a wavelength of 520 ± 5 nm to excite the voltage-sensitive and calcium indicator dyes simultaneously.
      NOTE: The excitation light source is attached to the excitation filter cube of the optical mapping system and is reflected off a 550 nm dichroic mirror for epicentric illumination. The emitted light is collected by a 1x lens and split by a second dichroic mirror at 630 nm into voltage and calcium components before it is filtered by 690 ± 50 nm and 590 ± 33 nm filters, respectively, and recorded by the two cameras.

3. Slicing protocol

1. When ready for tissue dissection and experimentation, prepare a bath of ice-cold cardioplegia by mixing frozen and liquid cardioplegia. Keep the heart submerged in the cardioplegia bath until tissue collection for slicing (Figure 1A).
2. Glue the premade agarose gel to the back of the metal tissue holders of the vibratome.
3. Identify the left ventricular free wall and cut 1 cm³ cubes of tissue in cold cardioplegic solution. Then, quickly mount the tissue blocks onto the metal tissue holders with the endocardial surfaces facing up and attach them to the agarose gel using topical skin adhesive (Figure 1B).
   NOTE: The selected tissue region should be away from large blood vessels to avoid holes in the slices. The plane of sectioning is approximately parallel to the fiber orientation in the endocardial layer (Figure 1E) and the plane of sectioning could be at a slight angle due to rotational anisotropy in a deeper layer of the LV wall. To increase throughput, up to two tissue blocks can be mounted side by side on the same holder.
4. Transfer the metal holders into the vibratome bath filled with the ice-cold oxygenated slicing solution. Ensure that the tissue cubes are completely submerged.
5. Move the blade to the front edge of the tissue and turn on the vibratome to begin slicing with the preset parameters. Discard the first several slices until the blade reaches beyond the trabeculae into the smooth endocardial tissue.
6. Once each slice is cut, carefully transfer the slice to an oxygenated (100% O₂) bath of Tyrode’s recovery solution at RT. Gently place each slice in individual 100 µm nylon mesh cell strainers and cover with meshed washers to keep the tissue slices from curling (Figure 1C). Keep slices in the recovery solution for at least 20 min.
   NOTE: Slices can be kept in the bath under these conditions for up to 3−4 h without detrimental electrophysiological effects.

4. Slice culturing under static conditions

NOTE: To minimize the chance of contamination, sterilize the forceps using a bead sterilizer before each transfer step.

1. When ready to culture, carefully transfer each slice to the individual wells of a 6 well plate filled with sterile phosphate-buffered saline (PBS). Gently rock the well plate to rinse the slices of recovery solution.
   NOTE: From this point forward, the slices should only be exposed in a BSL2 laminar flow culture hood, and sterile forceps should be used to handle the tissue.
2. Transfer the slices to wells of fresh PBS to thoroughly rinse and sterilize the slices for culture. Perform this wash step 3x to ensure complete removal of recovery solution from the slices.
3. Transfer the slices to individual wells of a 6 well plate filled with 3 mL of prewarmed (37 °C) culture medium (with or without drugs, Figure 1D). Place plates onto an orbital shaker at 20 rpm in a humidified incubator at 37 °C with 30% O₂ and 5% CO₂. Aspirate and replace culture medium every 48 h.

5. Functional characterization―optical mapping

1. For optical mapping studies either immediately following slicing or after culturing, carefully transfer the slice of interest to the tissue bath of the perfusion system at 37 °C with 100% O₂ in Tyrode’s recovery solution, and pin down the four corners to the PDMS gel layer while applying minimal stretch (i.e., just enough to keep the slice from easily moving under flow conditions). Allow the slices to rest in this bath with circulating recovery solution for approximately 10 min.
2. Add 0.3−0.5 mL of the diluted blebbistatin to the reservoir containing the recovery solution. Let the slice incubate with blebbistatin for approximately 10 min.
3. Reconstitute 30 µL of the stock voltage-sensitive dye, RH237, in 1 mL of recovery solution at 37 °C. Turn off the pumps and slowly load 0.2−0.3 mL of working dye solution onto the surface of the slice over a period of 30 s. Allow the slice to incubate in the dye with the pumps off for a period up to 90 s, then turn the pumps on again to allow any excess dye to wash out.
   NOTE: Care should be taken to apply the dye evenly all over the slice. Additionally, very slow dye application is crucial to prevent the dye from floating away from the slice.
4. Repeat step 5.3 to reconstitute the stock calcium indicator Rhod-2AM and load it onto slice.
5. Focus the cameras onto the slice by adjusting the distance of the slice from the lens.
6. Position a bipolar platinum-iridium pacing electrode such that its tip is in contact with the middle of the slice and apply pacing stimuli of increasing amplitude to determine the minimum pacing threshold required to elicit an electrical response. Pace the slice at 1 Hz with a 2 ms pulse width duration at 1.5x the amplitude of the predetermined pacing threshold. Place a coverslip over the tissue slice.
7. Illuminate the slice using a 520 ± 5 nm LED excitation light source. Record the emitted voltage and calcium signals with the two CMOS cameras at 1,000 frames/s.
8. Check the recording for good signal quality and suppressed motion artifacts (Figure 1F). Add more blebbistatin to the reservoir in steps of 0.1 mL until the motion no longer produces artifacts in the optical signals. Add more dye, if needed, to improve the signal quality.
   NOTE: Good signal quality refers to qualitatively assessed large signal amplitude and low background noise in the recorded optical signals.

6. Data processing with RHYTHM1.2

NOTE: RHYTHM1.2 is a MATLAB based user interface that is used to display, condition, and analyze optical mapping data acquired by single or dual camera optical mapping systems (Figure 3). It is used in conjunction with the imaging system.

1. Loading data files
   1. Load the data files into RHYTHM1.2 by selecting any one of the four display windows on the screen and using the Select Directory and Load buttons.
   2. The program will prompt the user to select Camera 1 or Camera 2 data for the dual voltage and calcium recordings. Select Camera 1 for the voltage signal.
   3. Repeat the process to load the calcium signal (Camera 2) to one of the adjacent display windows.
   4. Select the Dual Data Linking checkbox between the two selected display windows to link the two display windows, since the two data sets are recorded from the same region on the slice.
      NOTE: Alternatively, if the same field of view is maintained, this function can be used to link two images that were obtained at separate time points to see the time-response in electrophysiology of the drug being investigated. Double clicking any one display window allows for that one window to be maximized to allow for ease of analysis.
2. Signal conditioning
   1. Select Data Analysis | Condition Parameters to condition the voltage and calcium optical mapping data prior to further analysis.
   2. Use the “Remove Background” function to remove pixels that do not contain any signal based on a fluorescent intensity threshold (background [BG] threshold) and degree of pixel clustering (excitation [EX] threshold).
   3. Perform spatial averaging of the optical mapping data using the “Bin” function to improve signal quality.
   4. Filter the optical mapping data using a band pass filter with the “Filter” function.
   5. Use the “Drift” function to remove the baseline drift in the optical mapping signal.
   6. Using the “Normalize” function, normalize the optical mapping data from each pixel to have a maximal amplitude of 1.
   7. Use the “Inverse Signal” function to invert the calcium traces for further analysis.
   8. Click Display Wave to select the conditioned trace from any given spot in the display window and plot it in the Waveform window. Adjust the signal conditioning parameters to obtain optimal action potential and calcium transient traces.
3. Conduction velocity (CV) calculation
   NOTE: The activation time is defined as the time of maximum derivative of the optical signal (dF/dt_max).
   1. Select Data Analysis | Activation Map and enter a Start Time and End Time to encompass a single action potential in the trace. Press Calculate and select the region of interest (ROI) to display the activation map of the selected region (Figure 1G).
   2. Select Data Analysis | CV Map and similarly, choose the Start Time and End Time. Enter the interpixel resolution values based on the setup.
      NOTE: For 1x magnification systems, interpixel resolution is 0.1 mm in the X and Y direction.
   3. Press the Generate Vec. Map button to select an ROI and display CV vectors within that region. The mean, median, standard deviation, and number of vectors included in the analysis as well as the average angle of propagation of the CV vectors in the selected region is calculated and displayed in the Statistics section.
   4. Click the Draw Line button to draw a line along a given direction of propagation. All CV vectors in that direction will be selected. Click Calculate CV to use only those selected CV vectors to calculate the statistics that will be displayed in the Statistics section.
4. Action potential duration (APD) and calcium transient duration (CaTD) calculation
   NOTE: The APD is another fundamental parameter of the cardiac electrophysiology. APD maps can be generated by determining the time difference between activation and a specified percentage of repolarization of each optical action potential. APD heterogeneity or APD prolongation and shortening can be used to predict arrhythmia susceptibility.
   1. To calculate APD in RHYTHM1.2, select Data Analysis | APD/CaT Map.
   2. Select the Start Time and End Time to encompass one full action potential. Set a minimum and maximum value of APD/CaTD to remove any outliers (for example, 0 and 1,000 ms, respectively). Enter the percent repolarization that will determine the APD, for example, 0.8 for APD₈₀/CaTD₈₀ or 0.5 for APD₅₀/CaTD₅₀.
   3. Click Regional APD Calc to select an ROI and generate the APD map. The mean, median, standard deviation, and number of pixels included in the analysis will be displayed in the Statistics section.
5. Rise time
   NOTE: The rise time is another electrophysiological parameter that can be measured from optical signals of voltage and calcium transient traces. It provides an estimate for how long it takes for the depolarizing ion channels to trigger an action potential, or how long it takes for calcium to be released into the cytoplasm from the sarcoplasmic reticulum. Optical rise time is not a perfect substitute for rise times measured by microelectrodes and may not be as sensitive to changes in depolarization, because optical action potentials are an average of the transmembrane potential of many cells. Spatial and temporal resolution of the system can also affect optical rise time values. However, it can still be used to predict large changes in depolarization²⁷.
   1. To determine the rise time, select Data Analysis | Rise Time.
   2. Select Start Time and End Time to select the upstroke of one single action potential or calcium transient. Enter the values of Start% and End% (typically 10–90% is recommended for non-noisy signals), which allows the user to select just the portions of the signal without including noise in the case of noisy signals.
   3. Click Calculate to select the ROI and determine the mean, median, standard deviation, and number of pixels included in the analysis of rise time.
6. Calcium decay
   NOTE: For calcium traces, the time constant of the decay of the calcium transient can be determined. This allows for the analysis of changes in the removal of calcium ions from the cytoplasm back into the SR.
   1. Select Data Analysis | Calcium Decay and enter the start time and end time to encompass the entire decay portion of a single calcium transient signal.
   2. Click Calculate Tau to select the ROI and determine the mean, median, standard deviation, and number of pixels included in the analysis of calcium decay time constant.

Results

Human organotypic slices were collected from the left ventricle of a donor human heart according to the protocol detailed above and illustrated in Figure 1. A dual camera optical mapping system like that in Figure 2 was used in the upright imaging configuration to perform simultaneous optical mapping of voltage and calcium about 1 h after the completion of the slicing protocol. Data were analyzed using RHYTHM1.2 (Figure 3), an open ...

Discussion

Here, we present step-by-step methods to obtain viable cardiac slices from cardioplegically arrested human hearts and to functionally characterize the slices using dual optical mapping of transmembrane potential and intracellular calcium. With preserved extracellular environment and native cell-cell coupling, human cardiac slices can be used as an accurate model of the human heart for fundamental scientific discovery and for efficacy and cardiotoxicity testing of pharmacological agents and gene therapies. The technology ...

Materials

Name	Company	Catalog Number	Comments
1mL BD Syringe	Thomas Scientific	309597
2,3-butanedione monoxime	Sigma-Aldrich	B0753
6 well culture plates	Corning	3516
Biosafety cabinet	ThermoFisher Scientific	1377
Blebbistatin	Cayman	13186
Bubble Trap	Radnoti	130149
Calcium chloride	Sigma-Aldrich	C1016
Corning Cell Strainers	Fisher Scientific	07-201-432
Di-4-ANEPPS	Biotium		stock solution at 1.25 mg/mL in DMSO
DMSO	Sigma-Aldrich	D2650
Dumont #3c Forceps	Fine Science Tools	11231-20
Emission dichroic mirror	Chroma	T630LPXR-UF1
Emission filter (RH237)	Chroma	ET690/50m
Emission Filter (Rhod2AM)	Chroma	ET590/33m
Excitation dichroic mirror	Chroma	T550LPXR-UF1
Excitation Filter	Chroma	ET500/40x
Falcon 50mL Conical Centrifuge Tubes	Fisher Scientific	14-959-49A
Glucose	Sigma-Aldrich	G8270
Heat exchanger	Radnoti	158821
HEPES	Sigma-Aldrich	H3375
Incubator	ThermoFisher Scientific	50145502
Insulin Transferrin Selenium (ITS)	Sigma-Aldrich	I3146
LED excitation light source	Prizmatix	UHP-Mic-LED-520
Magnessium chloride hexahydrate	Sigma-Aldrich	M9272
Medium 199	ThermoFisher Scientific	11150059
Micam Ultima L type CMOS camera	Scimedia	N/A
Minutien Pins	Fine Science Tools	26002-10
Pennicillin-Streptomycin	Sigma-Aldrich	P4333
Peristaltic Pump	Cole Parmer	EW-07522-20
Platinum pacing wire	Alfa Aesar	43275
Pluronic F127	ThermoFisher Scientific	P6867	nonionic, surfactant polyol
Potassium chloride	Sigma-Aldrich	P3911
Powerlab data acquisition and stimulator	AD Instruments	Powerlab 4/26
RH237	Biotium	61018
Rhod2AM	ThermoFisher Scientific	R1245MP
Rhod-2AM	Invitrogen, Carlsbad, CA
Sodium bicarbonate	Sigma-Aldrich	S6014
Sodium chloride	Sigma-Aldrich	S9625
Sterilizer, dry bead	Sigma-Aldrich	Z378550
Stone Oxygen Diffuser	Waterwood	B00O0NUVM0
TissueSeal - Histoacryl Topical Skin Adhesive	gobiomed	AESCULAP
UltraPure Low Melting Point Agarose	Thermo Fisher Scientific	16520100
Ultrasound sonicator		Branson 1800
Vibratome	Campden Instruments	7000 smz