Measurement of Endothelium-Dependent Vasorelaxation in the Mouse Thoracic Aorta Using Tensometric Small Volume Chamber Myography

Summary

The present protocol describes the concepts and technical application of the tensometric myograph technique using a multi-chamber myograph system in the experimental ex vivo assessment of mouse aortic endothelial function.

Abstract

Small volume chamber tensometric myography is a commonly used technique to evaluate the vascular contractility of small and large blood vessels in laboratory animals and small arteries isolated from human tissue. The technique allows researchers to maintain isolated blood vessels in a tightly controlled and standardized (near-physiological) setting, with the option of adjusting to various environmental factors, while challenging the isolated vessels with different pharmacological agents that can induce vasoconstriction or vasodilation. The myograph chamber also provides a platform to measure vascular reactivity in response to various hormones, inhibitors, and agonists that may impact the function of smooth muscle and endothelial layers separately or simultaneously. The blood vessel wall is a complex structure consisting of three different layers: the intima (endothelial layer), media (smooth muscle and elastin fibers), and adventitia (collagen and other connective tissue). To gain a clear understanding of the functional properties of each layer, it is critical to have access to an experimental platform and system that would allow for a combinational approach to study all three layers simultaneously. Such an approach demands access to a semi-physiological condition that would mimic the in vivo environment in an ex vivo setting. Small volume chamber tensometric myography has provided an ideal environment to evaluate the impact of environmental cues, experimental variables, or pharmacological agonists and antagonists on vascular properties. For many years, scientists have used the tensometric myograph technique to measure endothelial function and smooth muscle contractility in response to different agents. In this report, a small volume chamber tensometric myograph system is used to measure endothelial function in the isolated mouse aorta. This report focuses on how small volume chamber tensometric myography can be used to evaluate the functional integrity of the endothelium in small segments of a large artery such as the thoracic aorta.

Introduction

For the last few decades, the small chamber myography system has been used to measure the reactivity of different layers of blood vessel walls in response to various pharmacological agents and neurotransmitters in an ex vivo, real-time setting. Vascular reactivity is a major component of a healthy functional blood vessel and is critical for the regulation of blood flow and perfusion in peripheral and cerebral vasculature¹. Within the blood vessel wall, the interaction between endothelial and smooth muscle layers is a major determinant of vascular tone, which is also constantly impacted by structural changes in the connective tissue layer surrounding the blood vessel wall (adventitia).

The endothelial layer controls vasomotion by releasing a few vasodilatory factors, including nitric oxide (NO), prostacyclin (PGI2), and endothelium-derived hyperpolarizing factor (EDHF), or by producing vasoconstrictive agents such as endothelin-1 (ET-1) and thromboxane (TXA2)^2,3,4. Among these factors, NO has been extensively studied, and its important regulatory roles in other critical cellular functions such as inflammation, migration, survival, and proliferation have been highly cited in scientific literature^2,5.

In the field of vascular biology, chamber myography has provided vascular physiologists and pharmacologists with a valuable and reliable tool to measure endothelial function in a tightly controlled semi-physiologic system¹. Currently, there are two different myograph systems available to scientists: wire (or pin) tensometric (isometric) myography and pressure myography. In a wire myography system, the blood vessel is stretched between two wires or pins, allowing for the isometric measurement of force or tension development in the wall of the blood vessel, while pressure myography is a preferable platform for measurements of vascular reactivity in small resistance arteries, where changes in blood pressure are considered the main stimulus for changes in vascular tone and vasomotion. There is a general agreement that, for small resistance arteries such as mesenteric and cerebral arteries, pressure myography creates a condition that is closer to the physiological conditions in the human body. The small chamber myograph can be utilized for vessels with very small diameters (200-500 µm) to much larger vessels such as the aorta.

While the wire myograph is a powerful system for recording blood vessel tension under isometric conditions, the pressure myograph is a more appropriate system for measuring changes in vessel diameter in response to changes in isobaric conditions. The diameter changes in the vessel in response to changes in pressure or flow are much larger in a small muscular artery (arteriole) compared to large elastic arteries such as the aorta. For these reasons, the pressure myograph is considered a better tool for small blood vessels with substantial vasoreactivity¹. One of the other practical strengths of multi-chamber small volume chamber tensometric myography is that one can discern the contribution of different mechanisms to vascular reactivity by studying multiple (up to four) segments of the same artery and from the same animal to reduce variability and produce robust and conclusive data. It is also relatively easy to set up and maintain technically. Vessels of almost any size can be studied with a wire myograph. It is a more cost-effective solution for assessing vascular function and is a good alternative to pressure myography in experiments where the length of the dissected vessel is too short for the pressure myograph protocol.

This report provides a detailed protocol for the assessment of endothelial function in the isolated mouse thoracic aortic ring using mounting pins in the small volume chamber tensometric myography technique using the DMT-620 multi-chamber myograph system (DMT-USA). This protocol utilizes a 6-month-old male C57BL6 mouse with an average weight between 25-35 g. Fortunately, this protocol can be applied to various animal types and weights, considering the broad range of vessel types and diameters that this protocol can be used for.

Protocol

All surgical procedures and animal care were approved by the Institutional Animal Care and Use and Care Committee (IACUC) of Midwestern University (IACUC# AZ-3006, AZ-2936).

1. Buffer preparation

NOTE: Although the HEPES physiological salt solution (HEPES-PSS) buffer is stable at 4 °C for 7 days, it is recommended that all buffers are freshly made on the day of each experiment. All other reagents and agonists must be prepared freshly for each experiment. The HEPES-PSS buffer used in this protocol is a well-established buffer for ex vivo vascular studies that has been shown to be cytoprotective for more than 12 h while preserving the vessel's vasodilatory responses-the main focus of this experimental protocol^6,7.

1. Prepare HEPES-PSS solution (pH 7.4) as follows: Mix 10 mmol/L HEPES, 6 mmol/L glucose, 1.5 mmol/L CaCl₂, 130 mmol/L NaCl, 4 mmol/L KCl, 4 mmol/L NaHCO₃, 1.2 mmol/L MgSO₄, 1.2 mmol/L KH₂PO₄, and 0.03 mmol/L EDTA.
2. Prepare HEPES-PSS high K⁺ buffer. This is identical to the HEPES-PSS solution, except that it contains 5 mmol/L HEPES, 65 mmol/L NaCl, 10 mmol/L glucose, 1 mmol/L MgCl₂, 80 mmol/L KCl and does not contain MgSO₄ and EDTA.

2. Myograph unit preparation

1. Turn on and set the water bath to 37 °C. Ensure an appropriate water level.
2. Place two beakers labeled appropriately into the water bath, one with 600 mL of HEPES-PSS solution and one with 150 mL of high K⁺ solution.
3. Aerate a beaker of 300 mL of HEPES-PSS solution with carbogen gas (5% CO₂ and 95% O₂) for at least 10 min.
4. Add 30 mL of the aerated HEPES-PSS solution to a 50 mL centrifuge tube, label appropriately (thoracic cage, mouse identification number), and place it on ice. Keep the remaining aerated HEPES-PSS solution on ice to be used during the aortic dissection process.
5. Turn on the four-chamber myograph unit; add 6 mL of HEPES-PSS solution to each myograph chamber, and set the heat to 37 °C. Allow the solution in each chamber to be aerated (with carbogen mix) for 30 min and check to ensure that the chambers have reached the desired 37 °C (use this waiting period to dissect the mouse aorta).
6. Turn on the myograph data acquisition hardware and computer.

3. Mouse aorta isolation

1. Anesthetize the experimental mouse with 5% isoflurane inhalation and euthanize by cervical dislocation (a 6-month-old male C57BL/6J mouse is used for this demonstration).
2. Lay the mouse on a surgical board in the supine position and secure the appendages to the board using surgical tape.
3. Spray the abdominal region with 70% alcohol so that the fur is completely wet and any loose/dry hairs do not enter the incision. Gently wipe away excess solution.
4. Use forceps to locate and isolate the abdominal skin just inferior to the xiphoid process of the sternum.
5. Create tension by lifting the skin straight up. Make a blunt cut using scissors, and remove the superficial skin, exposing the thoracic cavity.
6. Lift the xiphoid process and make lateral incisions just inferior to the process along the subcostal margins.
7. Extend the lateral incisions cranially to remove the anterior portion of the ribcage.
8. Remove the thoracic cage by making a blunt cut through the spine, below the diaphragm, and in the neck.
   NOTE: Researchers may also remove the aorta directly from the animal, but care should be taken to protect the vessel and flush the clotted blood from the aorta. This protocol uses the example of removing the thoracic cage and carefully cleaning and dissecting the aorta. With practice, this method can be performed quickly and efficiently and is beneficial for preserving the extra tissue for other studies such as immunohistochemistry, histology, and western blotting.
9. Transfer the thoracic cage to a clear silicone elastomer-coated Petri dish filled with ice-cold aerated HEPES-PSS buffer and pin it on both sides.
   NOTE: Use a silicone elastomer kit (Table of Materials). Mix base and curing agent (10:1 by weight), then pour into a glass Petri dish, and allow to cure for 24 h at 25 °C.
10. Place the Petri dish under a stereo zoom microscope, gently cut and remove the heart and attached aorta from the rib cage, and transfer to a clean, clear silicone elastomer-coated dish (Figure 1) .
11. Then, gently remove fat and connective tissue and any clotted blood from the aorta. Using sharp, small scissors, dissect and isolate the whole aorta starting from the arch area to the bottom part of the descending aorta. Remember that the aortic arch is not suitable for myography experiments but can be used for histological studies.
12. Use cold aerated HEPES-PSS solution throughout the dissection. Change the solution every 10 min or when visibility is compromised, whatever comes first. It is recommended that each aortic ring is mounted onto the myograph chamber within 20 min post dissection (max 60 min).

4. Mounting of the aortic segments onto the myograph chambers

1. Cut the dissected aorta into four segments of 2 mm each using sharp small surgical scissors.Use the mini-ruler within the dish as a reference.
2. Place the myograph chamber under the stereo zoom microscope to easily visualize the pins and set the micrometers such that the pins are nearly touching (Figure 2).
3. Make sure both tissue-holding pins are properly aligned.
4. Pay attention and avoid pinching the aorta while mounting the segments to prevent damage to the endothelium.
5. In chambers already containing 5 mL of warmed and aerated HEPES-PSS buffer, carefully slide each of the 2 mm aortic segments onto the two mounting pins using forceps. Be very gentle while sliding the aortic segments over the pins. The endothelial layer is very fragile and comes off very easily from the luminal side.
6. Slowly move the pins apart by rotating the micrometer counter-clockwise so the aortic segment does not slide off the pins when placing the chamber back in the myograph unit (Figure 3).
7. Use the previously calibrated eyepiece graticule of the dissecting microscope to measure the true and accurate length of the mounted aortic ring by positioning the beginning of the ruler at one end of the segment (marked as tissue end α1) and making a note of the measurement at the other end of the aortic segment in ocular divisions (marked as tissue end α2)⁸. These values will be used during the normalization steps in section 5.
8. Return each myograph chamber to the unit, and start aerating the chambers at 37 °C for 30 min.
   ​NOTE: The four segments of the aorta can be used as replicates for the same treatment, or each segment of the aorta can be used simultaneously for different experiments.

5. Normalization

NOTE: A normalization procedure is necessary to ensure that the experimental conditions are properly standardized and the collected data is reliable and reproducible. The "IC1/IC100", or "Normalization Factor", is defined as the ratio of the internal circumference of the artery at which it is possible to record the maximum response to a vasoconstrictor (e.g., 60 mM KCl) divided by the internal circumference at which a transmural wall pressure of 100 mm Hg (i.e., IC100) is recorded. Therefore, by multiplying the IC100 by this ratio, we can determine the internal circumference of the artery at which an optimal response (i.e., IC1) can be established.

1. Set the micrometer such that the pins are nearly touching.
2. Set forces to zero for all myograph chambers and let it equilibrate for another 1-2 min.
3. Make a note of the first diameter reading from the micrometer. This is the position where the gap between two pins is considered zero (needed for step 5.7.).
4. Open the data acquisition software Normalization Settings under the DMT drop-down menu; a new screen will open with the following settings and default values:
   Eyepiece calibration (mm/div): 0.36
   Target pressure (kPa): 13.3
   IC1/IC100: 0.9
   Online averaging time (s): 2
   Delay time (s): 60
   Play sound on delay completion (check box)
   Sounds (with drop-down selection menu or Browse function)
5. Depending on the number of segments, choose the correct number of channels available on-screen and start the chart recording.
6. Use the drop-down menu to select the channel of interest; the normalization screen will appear.
7. Enter the constant values into the windows as follows: Tissue end-points α1; Tissue end-points α2; Pin diameter: 40 µm; Micrometer reading value from the analog micrometer scale.
8. In order to record the first point (the initial value of X or X0), click on the Add Point button. Following a delay of 60 s, the values for the force and the effective pressure (ERTP) corresponding to this micrometer will be displayed, and the box for the micrometer reading becomes active and available.
9. Start stretching the vessel by turning the micrometer in a counter-clockwise direction. Use the micrometer reading box to enter the value and click the Add Point button (there will be a delay time of 60 s again).
10. Keep stretching the vessel and continue to add micrometer values until the value of Micrometer X1 is seen, which is the calculated micrometer setting used to stretch the vessel to its IC1.
11. Set the micrometer to the X1 value.
    NOTE: The normalized (optimal) tension for a 6-month-old C57BL6 mouse is 6 mN.
12. After normalization, let the tissue rest and equilibrate for 30 min. There is no need to change the HEPES-PSS solution in the chambers at this point.
    NOTE: Each chamber is capable of holding 8 mL of solution. This protocol is written with the use of 5 mL, which allows for full submergence of the vessel and mounting pins.
13. Use the rest and equilibration time to prepare the following.
    1. Prepare the working stock of acetylcholine serial dilutions in distilled water (RO water) from lowest to highest concentrations as follows: 50 nM, 100 nM, 500 nM, 1 µM, 5 µM, 10 µM, 50 µM, 100 µM, 500 µM, and 1 mM. Keep all the tubes on ice for the duration of the experiment.
    2. Prepare a 100 mM working solution of N-nitro-L-arginine methyl ester (L-NAME) in double-distilled water and keep the solution on ice for the duration of the experiment.
    3. Prepare the sub-maximum dose of phenylephrine (10 µM) determined in a separate set of experiments.
       ​NOTE: The equilibration period is necessary for the blood vessel segment to adjust to the new environment in the myograph chamber, reset ion gradients, and achieve a stable level of passive tension prior to being subjected to different pharmacological and mechanical challenges.

6. Measurement of endothelium-dependent vasorelaxation in aortic rings

1. At the end of the 30-min equilibration period, drain the chambers and add 5 mL of fresh, warm, aerated HEPES-PSS solution to each chamber. Drain one chamber at a time to minimize tissue exposure to air. Draining of the chambers is performed using a vacuum pump set to 60 cmHg and the myograph valves set to a 6 s delay to ensure the removal of the 5 mL of solution.
2. If needed, readjust the force measurements to read the optimal tension of 6 mN. It is very critical to readjust the force to optimal tension during every incubation period, as well as prior to each new set of experiments. Rest the tissue for another 15-20 min.
3. Open the data acquisition software, change the tracking numbers from 50:1 to 500:1, and press Start. At this stage, the registered force for each chamber can be seen in software.
4. Immediately prior to the start of a new experiment, be sure to add the appropriate label on the data acquisition software.
5. Drain the chambers one more time before adding 5 mL of high K⁺ solution to each chamber. This is suitable for testing the viability of aortic tissue and the integrity of the smooth muscle contractile response to membrane depolarization before using the tissue for any other experimental protocol. This helps determine whether the blood vessel segment is usable for further experimentation.
6. Drain the chambers again as soon as the contractile response to the high K⁺ solution reaches a plateau for force generation (Figure 4).
   NOTE: Do not leave the high K⁺ solution in the chambers for more than 3 min, as it would damage the tissue.
7. Wash the tissue with HEPES-PSS solution 3x. Add 5 mL of high K⁺ solution to each chamber. Drain the chambers as soon as the contractile response to high K⁺ solution reaches a plateau for force generation and wash the tissue 3x with HEPES-PSS before allowing the tissue to rest for another 15 min.
   NOTE: In some laboratories, the blood segments are subjected to a high K⁺ solution three consecutive times to ensure the variability of the blood vessel before initiating the myograph experiments. In this protocol, the newly isolated aortic segments are subjected to a high K⁺ solution twice before using the tissue for further experiments. It is important to keep this consistent across all experiments.
8. Use the average of the two registered peaks of force development (contraction force) in response to a high K⁺ solution to normalize the registered values in response to other vasoconstrictor and vasodilator agents used in the experiment.
9. Rest the tissue for 15-20 min.
10. Pre-contract the aortic segments with the vasoconstrictor agent phenylephrine (PE) at an already established sub-maximum dose (10 µM final concentration in the chamber).
    NOTE: The sub-maximum dose of phenylephrine (10 µM) was determined in a separate set of experiments, where aortic segments were subjected to increasing concentrations of phenylephrine solution at final concentrations of 1 nM, 5 nM, 10 nM, 50 nM, 100 nM, 500 nM, 1 µM, 5 µM, 10 µM, and 50 µM. As a result, it was determined that a final concentration of 10 µM of phenylephrine could generate the sub-maximum contraction force (90% of the maximum force) in 2 mm aortic segments, which is right before the contraction force reaches a plateau. The reason for using the sub-maximum concentration of phenylephrine is to avoid issues related to aortic tissue saturation or desensitization to the vasoconstricting agonist.
11. Allow the PE-induced contraction curve to reach a plateau for tension (force) development (Figure 5).
12. At a plateau of tension development to phenylephrine, perform the acetylcholine dose-response curve by adding 5 µL of increasing doses of acetylcholine working stock solutions in 3 min intervals in order to establish final concentrations of acetylcholine in the myograph chamber as follows: 50 pM, 100 pM, 500 pM, 1 nM, 5 nM, 10 nM, 50 nM, 100 nM, 500 nM, and 1 µM (Figure 6).
    NOTE: After adding each dose of acetylcholine, wait for at least 3 min (for the tension to reach a plateau) before adding the next dose.
13. At each step, pipette the acetylcholine stock solution into the chamber very slowly and far from the aortic rings to avoid any tissue disturbance.
14. After completing the dose-response experiment, drain the chambers and wash the aortic segments with warm and aerated HEPES-PSS solution 3x to remove any remaining residue of the drug.
15. Rest the tissue for 30 min before starting the next experiment steps. The viability of the aorta is checked before each experimental step using HEPES-PSS high K⁺ solution. If all the preparation and experimental steps are followed correctly, the aortic tissue viability will remain the same for the duration of the entire experiment (4-6 h).

7. Effects of general inhibitors of NO production on endothelium-mediated vasorelaxation

1. Use the same aortic tissue for this part of the experiment after carefully washing and resting the segments for at least 30 min.
   NOTE: Use the 30 min resting time to prepare a 100 mM working solution of N-nitro-L-arginine methyl ester (L-NAME) in double-distilled water and keep the solution on ice for the duration of the experiment.
2. After resting the aortic segments for 30 min, drain the chambers and add fresh, warm high K⁺(60 mM KCl) solution to each chamber.
   NOTE: It is important that, for each part of the experiment, the aortic tissue is challenged with a high K⁺ solution at least once. The recorded contraction peak will be used for normalizing the data collected during that part of the experiment.
3. Drain the chambers again as soon as the contraction response to high K⁺ solution reaches a plateau and wash the tissue with HEPES-PSS solution 3x.
4. In order to assess the contribution of NO production to the observed acetylcholine-induced vasorelaxation (see section 6.), at this stage, preincubate the aortic segments with a general inhibitor of NO production (L-NAME) by adding 10 µL of the prepared 100 mM working stock solution of L-NAME to each myograph chamber (containing 5mL buffer solution) in order to achieve a final concentration of 200 µM in each myograph chamber.
   NOTE: L-NAME is a non-selective and potent inhibitor of all isoforms of NOS (an enzyme responsible for NO production), and, therefore, it is considered an effective tool to block the production of NO in the blood vessel wall⁹.
5. Allow the aortic segments to rest during the pre-incubation time with L-NAME (30 min).
6. Do not wash at this point.
7. Without removing the L-NAME, add the sub-maximum dose of phenylephrine (10 µM final concentration) to the chamber to induce aortic contraction.
   NOTE: Due to the inhibitory action of L-NAME on endogenous NO production, the newly registered peak for phenylephrine-induced aortic contraction will be much higher than the initial peak that was recorded prior to incubating the tissue with L-NAME. This is expected due to L-NAME's effect on basal NO production in the aortic wall, which leads to higher force generation (due to higher contractile force generation by the smooth muscle).
8. Wait until the phenylephrine-induced contraction curve reaches a plateau for tension (contractile force) development.
9. At this point, add acetylcholine to the myograph chamber to achieve a final concentration of 500 nM (this is the sub-maximum concentration of acetylcholine that was established during the experiments described in section 5. of the protocol).
10. Wait for a few minutes to register any possible changes in force development until the formed plateau is stable (Figure 7).
    NOTE: Due to the inhibitory action of L-NAME on the endothelial layer's ability to produce NO in response to acetylcholine, it is not expected to see any changes in registered force generation in response to phenylephrine, as the aortic endothelial NOS (eNOS) is not able to generate NO in response to acetylcholine application.
11. Drain the chambers and wash the tissue with warm, aerated HEPES-PSS solution 3x.

8. Contribution of the endothelial layer to aortic vasorelaxation

1. In order to underscore the role of the endothelial layer in aortic vasorelaxation, test acetylcholine-induced vasorelaxation in endothelium-denuded aortic segments, in which the endothelial layers are removed.
2. Position the myograph chamber under the microscope and gently pass a small wire (the same wire that is used for wire myography can be used here) through the lumen of the aorta. Gently move the wire through the lumen (gentle circular movement) for a short period of time. This is enough to remove the endothelium or intima.
3. Cut the aorta into 2 mm aortic rings and mount those rings onto the myograph chamber as described before.
4. Set the optimal tension at 6 mN and allow the tissue to rest for 30 min in aerated, warm HEPES-PSS buffer
5. At the end of the 30 min equilibration period, drain the chambers and add 5 mL of fresh, warm, aerated HEPES-PSS solution to each chamber.
6. Zero the force for all myograph chambers by turning the micrometer in a counter-clockwise direction.
7. Slowly rotate the micrometer counter-clockwise to increase the distance between the pins until the registered force reaches the desired optimal tension for the mouse aorta (6 mN for C57BL/6J mouse aorta). Rest the tissue for another 15-20 min at the optimal tension (6 mN).
8. Drain the chambers 1x more before adding 5 mL of high K⁺ solution to each chamber.
9. Drain the chambers again as soon as the contraction response to high K⁺ solution reaches a plateau and wash the tissue with HEPES-PSS solution 3x. Rest the tissue for 15-20 min
10. Pre-contract the aortic segments with the vasoconstrictor agent phenylephrine (10 µM final concentration in the chamber).
    NOTE: Due to the removal of the endothelium, the basal NO production in the aortic wall is significantly diminished; therefore, the peak for phenylephrine-induced contraction (the peak of force generation) is expected to be higher in aortic tissue lacking the endothelial layer.
11. When the phenylephrine-induced force reaches a plateau, add the sub-maximum concentration of acetylcholine to the myograph chamber to achieve a final concentration of 500 nM.
    NOTE: If the removal of the endothelial layer is done properly, there will not be any changes in the registered phenylephrine-induced force, as the acetylcholine-induced NO production by the endothelium will have been diminished due to the removal of the endothelial layer. The recorded trace will be very similar to the one observed in the presence of L-NAME (section 7.).
12. Wait for 3 min to ensure that the registered plateau for phenylephrine-induced force development is not changing before ending the experiment (Figure 8).
    NOTE: If the application of acetylcholine causes any drop in phenylephrine-induced contraction while in the plateau, this is an indication that the endothelial layer has not been completely removed.

Results

The tensometric small chamber myography protocol explained here is the standard method for measuring vascular reactivity in small and large arteries and allows for simultaneous measurements of vascular reactivity in up to four blood vessel segments from the same experimental small laboratory animal. In this report, we specifically use the system to measure endothelial function in the isolated mouse aorta (Figure 1). In this protocol, isolated aortic segments are mounted onto a small organ ch...

Discussion

The field of vascular biology heavily relies on tools that help researchers to assess the functional and structural integrity of the blood vessel wall. It also demands special attention on the direct and indirect interactions between the three layers of blood vessels: the intima, media, and adventitia. Among those three layers, the intima is formed by a monolayer of endothelial cells and has a very important function in regulating vascular health and hemostasis.

It is well established that any...

Materials

Name	Company	Catalog Number	Comments
Acetylcholine	SigmaAldrich	A6625-100G
CaCl2	SigmaAldrich	C4901-1KG
Carbogen gas	Matheson	H103847
Dissecting scissors	FST	91460-11
DMT 620 Multi chamber myograph system	DMT	DMT 620	Multi chamber myograph system
Dumont forceps	FST	91150-20
EDTA	SigmaAldrich	E5134-10G
Glucose	SigmaAldrich	G8270-1KG
HEPES	SigmaAldrich	H7006-1KG
KCl	SigmaAldrich	P9541-1KG
KH2PO4	SigmaAldrich	P5655-1KG
LabChart	ADI instruments		Data acquisition software
Light source	Volpi	14363
L-Name	Fischer Scientific	50-200-7725
MgSO4	SigmaAldrich	M2643-500G
Microscope	Leica	S6D	stereo zoom microscope
NaCl	SigmaAldrich	S5886-5KG
NaHCO3	SigmaAldrich	S5761-500G
Organ bath system	DMT	720MO
Phenylephrine	SigmaAldrich	P6126-10G
Pump	Welch	2546B-01
Software	ADI instruments	LabChart 8.1.20
Spring Scissors	FST	15003-08
Sylgard 184 Kit	Electron Microscopy Services	24236-10	silicone elastomer kit
Tank Regulator	Fischer Scientific	10575147
Water bath system	Fischer Scientific	15-462-10