Wire Myography for Dorsal Aorta and Mesenteric Resistance Arteries from Mice and Rats

Summary

We provide a detailed protocol for side-by-side wire myography to assess vascular reactivity in conduit versus resistance arteries isolated from mice and rats.

Abstract

Wire myography is a widely used ex vivo technique for the quantitative assessment of vascular function in isolated arteries. Because large conduit and small resistance arteries differ fundamentally in structure and physiological role, analyzing both vessel classes in parallel using arteries harvested from the same mouse or rat enables paired comparisons that reduce inter-animal variability and reveal vessel-specific functional changes. This article presents a step-by-step protocol for assessing vascular reactivity in isolated murine conduit and resistance arteries using isometric wire myography. We describe the preparation of fresh, oxygenated physiological buffer; careful dissection, cleaning, and mounting of arterial segments; and normalization to set vessels near their optimal length for active force generation. A troubleshooting guide and an example experimental sequence for generating cumulative concentration-response curves to standard vasoconstrictors and vasodilators are provided. This protocol allows quantitative measurement of both endothelial and smooth muscle function and facilitates detection of vessel-specific alterations in vascular reactivity associated with vascular disease, such as hypertension.

Introduction

Hypertension is characterized by altered vascular reactivity, such as endothelial dysfunction and vascular smooth muscle hyperreactivity to vasoconstrictors¹. These functional changes, together with microvascular remodeling and rarefaction, contribute to impaired tissue perfusion and dysregulated blood pressure^2,3,4. Ex vivo assays of isolated vessels are powerful tools for dissecting the cellular and molecular mechanisms that control vascular tone in health and disease, as they allow precise control of the vascular microenvironment and selective pharmacological manipulation of smooth muscle and endothelial signaling pathways.

Vascular reactivity in isolated arteries is commonly assessed using two complementary ex vivo techniques: wire myography and pressure myography^5,6. In wire myography, vessel rings or segments are mounted on pins or wires, and isometric force is recorded^7,8. In pressure myography, intact vessel segments are cannulated and pressurized to a defined intraluminal pressure, and vessel diameter is tracked by video microscopy⁹. Unlike wire myography, pressure myography permits control of transmural pressure and intraluminal flow. Thus, it more closely recapitulates the in vivo hemodynamic environment of resistance arteries and is commonly used to study myogenic tone, flow-mediated responses, and vascular structural or mechanical properties^9,10. Conversely, wire myography is technically easier to implement (no cannulation or video microscopy), supports higher throughput using commercial multi-chamber systems, and is well-suited for generating reproducible pharmacological concentration-response curves in small-to-medium arteries with diameters on the order of tens to several hundred micrometers^5,6. Accordingly, wire myography is often preferred for side-by-side pharmacological profiling of vascular reactivity in conduit and resistance arteries isolated from mice or rats.

Conduit (large elastic) and resistance (small muscular) arteries differ fundamentally in structure and function, and these differences dictate distinct experimental considerations and endpoints. Conduit arteries (e.g., descending thoracic aorta) contain large amounts of elastic tissue and substantial smooth muscle mass; resistance arteries (e.g., mesenteric arteries) have relatively greater smooth-muscle-to-lumen ratios and are major determinants of peripheral resistance and local autoregulation^11,12. Because the same agonist can activate different molecular responses in these vascular beds, side-by-side comparison of conduit and resistance vessels from the same animal provides useful complementary information. For instance, in conduit arteries, endothelium-derived nitric oxide release is the predominant relaxant pathway by which acetylcholine (ACh) induces vasorelaxation^13,14. In contrast, in small resistance arteries, ACh induces relaxation via nitric oxide, prostacyclin, and, importantly, endothelium-derived hyperpolarizing factors (EDHF)^15,16. Accordingly, side-by-side wire myography of conduit and resistance arteries harvested from the same mouse or rat enables paired comparisons that facilitate detection of vessel-specific functional changes while reducing inter-animal variability.

Accurate interpretation of wire myography data depends on careful attention to a set of technical variables: composition and oxygenation of the physiological salt solution (PSS), tissue handling and dissection, mounting technique, optimal resting pretension (normalization), bath temperature, and selection of appropriate pharmacological controls⁶. Normalization refers to the technical procedure of setting the mechanical operating point of the vessel so that active force development is optimized and approximates physiological operating conditions⁵. Practically, for aortic rings, it is common practice to use a fixed pretension, often determined empirically to avoid overdistension while giving stable responses. For small resistance arteries, normalization is performed by first determining the internal circumference corresponding to an equivalent transmural pressure of 100 mmHg (IC₁₀₀). The vessel is then set to a defined fraction of this value to establish the normalized internal circumference (IC₁). In practice, many protocols set IC₁ near 0.9 × IC₁₀₀ for rat MRA and around 1.0 × IC₁₀₀ for mouse MRA, although these values may be adjusted based on vessel type, species, and experimental goals^5,17.

Protocol

The following protocol adheres to the guidelines set and has been approved by The University of Toledo Health Science Campus Animal Care and Use Committee.

CAUTION Before you begin, please follow Institutional Animal Care and Use Committee (IACUC) and Institutional Biosafety Committee (IBC) guidelines at all times; use appropriate personal protective equipment (PPE); exercise care when using surgical or other sharp instruments; consult and comply with relevant Safety Data Sheets (SDS) when handling chemicals and biological tissues; handle carbogen as a pressurized gas cylinder using approved regulators and secure storage; and dispose of biological and chemical waste according to institutional procedures.

1. Preparation of PSS/modified Krebs bicarbonate (Krebs) buffer

NOTE: Krebs buffer composition (final concentrations, pH 7.4): 130 mM NaCl, 4.7 mM KCl, 1.17 mM MgSO₄, 1.18 mM KH₂PO₄, 1.6 mM CaCl₂, 25 mM NaHCO₃, 5.5 mM glucose, and 0.03 mM EDTA.

1. Prepare large-volume stock: dissolve all constituents except NaHCO_3, glucose, and CaCl₂ in double-distilled water to make a stock. Store this stock at 4 °C.
2. On the day of the experiment, aliquot the required volume of stock (e.g., 1 L), sequentially add NaHCO₃, glucose, and CaCl₂, and continuously oxygenate with carbogen (95% O₂/5% CO₂).
   NOTE: To minimize calcium salt precipitation, prepare and store a 1 M CaCl₂ stock at 4 °C and add the required volume slowly to the buffer only after NaHCO₃ is fully dissolved, while stirring continuously. Maintain carbogen bubbling during CaCl₂ addition (see step 1.3) to reduce calcium carbonate formation. If visible precipitation occurs, discard and prepare fresh buffer.
3. For pH equilibration, bubble the buffer with carbogen for at least 20 min, then measure pH and adjust to pH 7.4 if necessary.
   NOTE: Typical prebubbling pH ≈ 7.7; continuous bubbling reduces pH to physiological ≈7.4.
4. Prepare myograph chambers by filling them with freshly prepared Krebs buffer and begin gentle bubbling with carbogen. Adjust the bubbling rate to ensure adequate gas exchange without vigorous agitation.
5. Reserve an aliquot of freshly prepared, oxygenated Krebs buffer and chill it on ice for tissue dissection. Warm the remaining oxygenated buffer to 37 °C (with continuous carbogen bubbling) for use during experiments.

2. Dissection of the descending thoracic aorta and mesenteric arcade

1. Euthanize the animal according to IACUC guidelines. Standardize the timing of animal euthanasia and tissue harvest within a 2 h window to minimize potential circadian effects.
2. Immediately open the thoracic and abdominal cavities via a midline incision and reflect the chest wall laterally to expose the heart and great vessels.
   NOTE: Do not perform systemic perfusion under standard conditions, as this may inadvertently cause endothelial damage. Furthermore, retained blood in the mesenteric vessels enhances the visual discrimination between arteries and veins and facilitates assessment of arterial integrity. However, in some pathological models (e.g., thrombotic disorders), gentle, low-pressure perfusion with heparinized, ice-cold physiological buffer may be advisable to clear obstructing thrombi while minimizing endothelial injury.
3. Excise the descending thoracic aorta. To facilitate dissection, remove the lungs and esophagus to improve visualization. Do not directly grasp the descending thoracic aorta; instead, gently hold the aortic arch or periadventitial fat, lift slightly, and carefully sever the intercostal branch vessels from below the aorta. Excise the descending thoracic aorta by cutting just before it passes through the diaphragm and place it in a Sylgard-coated Petri dish containing ice-cold oxygenated Krebs buffer.
   NOTE: Maintain cold temperature to minimize spontaneous tone and preserve viability. Black Sylgard increases contrast for vessel dissection.
4. Excise the whole mesenteric arcade. Lift the mesenteric arcade gently to locate the superior mesenteric artery branching from the abdominal aorta caudal to the diaphragm. Cut the superior mesenteric artery proximally and free the mesenteric arcade from the surrounding connective tissue without direct vessel manipulation. Place the mesenteric arcade in a separate Sylgard-coated Petri dish containing ice-cold oxygenated Krebs buffer.

3. Isolation and mounting of aortic rings on pin myographs

1. Pin the proximal and distal ends of the descending thoracic aorta with dissection pins to straighten the vessel. Avoid overstretching to prevent mechanical damage.
   NOTE: Place pins in the periadventitial fat rather than through the vessel wall to minimize damage.
2. Under a stereomicroscope with an external light source, carefully remove periadventitial fat and connective tissue using fine Vannas scissors and forceps. Avoid damage to the vessel wall.
   NOTE: It is common practice to remove the periadventitial fat unless the goal is to study the role of perivascular adventitial tissues in the regulation of vascular tone.
3. Cut 2 mm aortic rings, making clean, perpendicular cuts. Discard rings that are punctured or visibly damaged. Whenever possible, sample comparable locations across animals to minimize variability.
4. Disconnect one chamber containing oxygenated Krebs buffer from the myography machine and carefully transport it to the stereomicroscope.
   NOTE: Jostling the chamber can affect the transducer and may alter results, so hold the chamber carefully and place it down gently.
5. Under the stereomicroscope, use the micrometer to bring the myograph pins together and slide the aortic ring onto the pins, minimizing manipulation of the luminal surface.
6. Note the micrometer reading with the ring fully relaxed; apply a slight stretch to secure the tissue for transport.
7. Connect the chamber to the instrument and restore the vessel to the original micrometer reading.
8. After all rings are mounted, heat the chambers to 37 °C and monitor temperature with the supplied probe (typically 20-30 min to reach 37 °C).
9. Zero the force readings on the user interface, launch the data acquisition software, and begin recording.
10. Incrementally apply pretension to aortic rings to a vessel-appropriate fixed value using the micrometer.
    NOTE: A typical pretension value for mouse aortic rings is 5 milliNewtons (mN)⁵. While reported pretension values for rat aortic rings vary across studies, prior optimization supports a pretension of 10 mN in this protocol¹⁸.
11. Equilibrate aortic rings for 45-60 min at the selected pretension, with repeated buffer changes using 37 °C Krebs.
    NOTE: It is common for aortic rings to partly lose tension and thus require fine readjustment of pretension several times using the micrometer before the tension stabilizes.
12. Zero the force readings on the user interface before beginning experimental protocols.

4. Isolation and mounting of MRA on wire myographs

1. Pin the mesenteric arcade in a clockwise pattern to straighten the mesenteric branches. Begin by securing the cecum, then pin successive stretches of small intestine clockwise to gently tension the mesenteric vasculature without overstretching. Replace the buffer with fresh Krebs as needed to remove blood or debris.
2. Identify the superior mesenteric artery at the center of the dish where arcades radiate.
3. Select an appropriate MRA for isolation; typical targets: internal diameter 200-250 µm (determined later during normalization).
   NOTE: In adult experimental animals, this generally corresponds to 2^nd-order branches in mice and 3^rd-4^th-order branches in rats. Branch order is defined by sequential bifurcations from the superior mesenteric artery. 1^st-order branches arise directly from the superior mesenteric artery, 2^nd-order branches arise from 1^st-order branches, and so forth.
4. Under a stereomicroscope, dissect away the accompanying vein and remove periadventitial fat and connective tissue using Vannas scissors and fine forceps.
   NOTE: MRAs can be distinguished from veins by their thicker walls and smaller lumens, which are readily visualized by the presence of residual intraluminal blood.
5. Relax the nearby gut-pin(s) that are stretching the gut wall adjacent to the selected vessel before excising the artery to prevent vessel recoil.
6. Excise the whole length of the selected MRA (should be >2 mm long), making clean, perpendicular cuts. Using fine forceps, grasp one end of the artery and transfer it to a clean Petri dish containing ice-cold Krebs buffer (regular, non-Sylgard-coated dishes facilitate subsequent mounting). Inspect the artery for damage; the presence of intraluminal blood facilitates visual assessment of integrity.
7. Excise an approximately 2 mm segment from an undamaged region. Discard any areas that were handled during transport or show visible damage.
   NOTE: Using transparent, non-Sylgard-coated dishes allows for the placement of a ruler underneath the dish to accurately measure the vessel length.
8. Prepare two 2.5 cm lengths of 40 µm stainless steel wire for mounting.
9. Using fine forceps, carefully thread the wires, one at a time, through the lumen so there is sufficient wire projecting from both ends for mounting. Minimize endothelial trauma. After inserting both wires, record the exact length with a ruler for use in the normalization procedure (see step 5.2).
   NOTE: This is one of the most technically challenging and time-consuming steps of the protocol and requires patience and repeated practice to minimize endothelial damage. After inserting the first wire, it is often easier to clamp both wires together and advance them through the vessel.
10. Disconnect one chamber containing oxygenated Krebs buffer from the myography machine and carefully transport it to the stereomicroscope.
11. Under the stereomicroscope, transfer the MRA (holding wires at both ends) to the myograph chamber. Using the micrometer, bring the myograph jaws together to clamp the wire ends between the jaws so the artery is suspended freely between them.
    NOTE: At this stage, it is advisable to clamp the wires such that they lie on top of one another. This secures the wires in place and prevents them from twisting.
12. Secure the top wire under the mounting screws at the near micrometer side by wrapping the ends in a clockwise fashion; repeat for the second wire on the transducer side.
    NOTE: Tighten the screws gently but firmly. If the screws are too loose, the vessel may slip; if over-tightened, the wire or transducer may be damaged. After fastening the first end, slightly open the gap between the myograph jaws before securing the opposite end to ensure even wire tension.
13. Slightly move the jaws apart and align the wires to lie parallel and coplanar, nearly touching.
    NOTE: Perform all wire adjustments away from the mounted vessel to minimize damage.
14. Repeat mounting for all MRAs.
15. Heat chambers to 37 °C and monitor temperature (20-30 min typical).
16. Zero the force readings on the user interface before initiating the normalization procedure.

5. Normalization procedure for the MRA

NOTE: Aortic rings are mounted at a fixed, predefined pretension. Therefore, a separate normalization step is not performed.

1. Set normalization parameters in the data acquisition software: eyepiece calibration = 1; target pressure = 13.3 kPa ( ≈ 100 mmHg); normalized internal circumference (IC₁) = 0.9 for rat MRAs and 1.0 for mouse MRAs; averaging time = 2 s; delay = 60 s.
2. Enter the vessel-specific parameters into the normalization module as follows:
   1. For tissue endpoints (mm): set a1 = 0 and a2 = the measured length of the MRA in millimeters (the exact length recorded during excision).
   2. Enter wire diameter: 40 µm.
   3. Micrometer reading: enter the baseline micrometer value recorded with the vessel mounted and fully relaxed.
3. Start the normalization procedure by stepwise stretching the mounted MRA in 50 µm increments. After each stretch, enter the new micrometer reading into the normalization module and wait 60 s between the increments.
   NOTE: Sudden drops in tension after a micrometer adjustment usually indicate vessel slippage. Mounting screws should be tightened gently but firmly.
4. Wait for the normalization module to plot resting wall tension (mN/mm) versus internal circumference (µm) and fit an exponential curve. Using the micrometer, continue stepwise stretching until the tension-circumference curve intersects the 100 mmHg isobar. When near the intersection, reduce the step size to 10-30 µm to avoid overstretching the vessel.
5. When the curve intersects the 100 mmHg isobar, note the software-calculated internal circumference and set the micrometer to the corresponding value calculated by the module so the preparation rests at the target normalized internal circumference (IC₁).
   NOTE: Poor curve fitting usually indicates incorrect parameters entered into the software, such as tissue length, wire diameter, or micrometer readings.
6. Repeat the normalization procedure for all mounted MRAs, then allow vessels to equilibrate for 30-45 min with repeated buffer changes using 37 °C Krebs.
7. Zero the force readings on the user interface immediately before beginning experiments.
   ​NOTE: Key parameter differences between mice and rats are summarized in Table 1.

Parameter	Mouse	Rat
Pretension Value for Aortic Rings	5 mN	10 mN
Target IC1 Ratio for MRA	1	0.9
Typical MRA Branch Order for 200-250 μm Internal Diameter	2nd	3rd-4th

Table 1: Key parameter differences between mice and rats.

6. Experimental protocol (example sequence)

1. Viability check: Elicit a contractile response with a depolarizing high-K⁺ challenge (e.g., 120 mM KCl).
   NOTE: The high-K⁺ buffer (KPSS) is prepared by equimolar substitution of NaCl with KCl from the standard Krebs buffer to maintain osmolarity and is continuously oxygenated with 95% O₂/5% CO₂. To avoid cold shock, warm the KPSS to 37 °C before use. KPSS buffer composition (final concentrations, pH 7.4): 120 mM KCl, 14.7 mM NaCl, 1.17 mM MgSO₄, 1.18 mM KH₂PO₄, 1.6 mM CaCl₂, 25 mM NaHCO₃, 5.5 mM glucose, and 0.03 mM EDTA.
2. Perform at least three consecutive washes with fresh 37 °C oxygenated Krebs buffer until a stable passive basal tension is restored.
   NOTE: All wash steps are performed by completely removing the bathing solution and replacing it with fresh 37 °C oxygenated Krebs buffer. Commercial myographs commonly have a built-in wash function that drains the myography chambers.
3. Generate a cumulative concentration-response curve (CRC) to phenylephrine (PE), adding increasing concentrations (example range 10⁻⁹ to 10⁻⁴ M). Allow each concentration to plateau before adding the next concentration until a stable maximal plateau is achieved.
4. Perform at least three consecutive washes with fresh 37 °C oxygenated Krebs buffer until a stable passive basal tension is restored.
5. Assess endothelial function: Preconstrict with a submaximal concentration of PE, allow response to stabilize, and then assess endothelium-dependent relaxation with ACh (example range 10⁻⁹ to 10⁻⁵ M). Allow each concentration to plateau before adding the next concentration until a stable maximal relaxation is achieved. Record percent relaxation relative to the preconstricted tone (See step 7.4). Preparations exhibiting <70% relaxation are excluded.
   NOTE: Vascular wall diseases, such as hypertension, are commonly associated with endothelial dysfunction, which often leads to impaired endothelium-dependent vasorelaxation response to ACh¹⁹. Under these conditions, preparations exhibiting <70 % relaxation should not be excluded automatically, as this may represent true disease related endothelial dysfunction rather than technical failure.
6. Perform at least three consecutive washes with fresh 37 °C oxygenated Krebs buffer until a stable passive basal tension is restored.
7. Test other vasoactive agents, with and without pathway-specific inhibitors, as required by the experimental question_.
8. At the end of the experiment, remove all the mounted vessels, clean all chambers with 8% acetic acid, followed by three washes of water, and finally turn off the gas tank and the machine.
9. Dispose of biological and chemical waste according to institutional procedures.

7. Data analysis and reporting

1. Baseline definition and recording. Record the baseline tension immediately before the start of any vasoconstrictor challenge (i.e., high-K⁺ challenge, vasoconstrictor CRC, or preconstriction before a vasodilator CRC).
   NOTE: For subsequent analysis, it is practically easier to zero the force readings on the user interface after adequate washing of any previous stimulation, immediately before the start of any vasoconstrictor challenge (i.e., displayed baseline tension = 0).
2. High-K⁺ challenge. Compute the active force as: Active Force = Maximum Tone after KCl - Baseline tension (units: mN). Express the active force normalized to either unit length (mN/mm) or cross-sectional area (mN/mm²) of the mounted arterial segment. Measurement of cross-sectional area requires subsequent histological analysis of the mounted artery segments to determine wall thickness.
   NOTE: Because aortic rings often produce a non-plateauing response, measure the maximum high-K⁺ response within a pre-specified time (e.g., 10 min) across all preparations, to minimize variability. In MRAs, maximum high-K⁺ response is usually achieved within seconds after stimulation.
3. Contractile CRCs. For each concentration in a contractile cumulative CRC (e.g., PE), calculate the active force as follows. Express the active force at each concentration as a percentage of the response elicited by the high-K⁺ challenge. Use these % values for fitting contractile CRCs (See step 7.5).
   Active Force = Maximum Tone after drug application - Baseline tension (units: mN)
   NOTE: Certain conditions may cause variability in the maximum high-K⁺ responses across preparations (e.g., changes in smooth muscle mass or differentiation state resulting from pathology). Under these circumstances, express the active force at each concentration normalized to either unit length (mN/mm) or cross-sectional area (mN/mm²) of the mounted arterial segment.
4. Relaxant CRCs. For relaxation CRCs after a submaximal preconstriction (preconstriction tone), compute percent relaxation at each concentration as follows. Use these % values for fitting relaxation CRCs (See step 7.5).
   Relaxation = (Preconstriction tone - Tone after drug application) / (Preconstriction tone - Baseline tension) × 100.
5. Curve fitting. For either contractile or relaxant cumulative CRCs, plot log [agonist concentration] on the x-axis versus the normalized response on the y-axis. Fit the plotted curve with a four-parameter logistic model using non-linear regression. Extract E_max (maximal effect) and pEC₅₀ (−logEC₅₀) for statistical comparisons, where EC₅₀ is the agonist concentration producing 50% of E_max.
6. Data presentation: Present the normalized CRCs. For each of the data points on the curve, use group mean ± standard error of the mean (SEM) or standard deviation (SD). Include representative original traces and provide a parameter table summarizing the E_max and pEC₅₀ values (mean ± SEM or SD).
7. Statistical analysis:
   1. Compare CRCs between experimental groups using two-way repeated measures analysis of variance (ANOVA) with an appropriate post-hoc test (e.g., Bonferroni). However, check assumptions (sphericity, normality) and use alternative approaches (e.g., mixed effects models) when data are missing or unbalanced.
   2. Compare pEC₅₀ and E_max between two experimental groups using an unpaired t-test, or one-way ANOVA if comparing more than two experimental groups. Alternatively, use non-parametric tests when variance homogeneity and normality assumptions are violated. Set statistical significance at p < 0.05.

Results

A schematic depicting the overall workflow of wire myography in conduit and resistance arteries isolated from mice or rats is shown in Figure 1. Representative images of isolation and mounting of a mouse thoracic aortic ring and a 2^nd-order MRA are shown in Figure 2 and Figure 3, respectively. Software screenshots depicting pretensioning and normalization procedures using a rat aortic ring and an MRA are shown in

Discussion

Wire myography remains a powerful, high-resolution approach for interrogating vascular function ex vivo. When executed carefully, this method provides reliable quantitative endpoints that address distinct biological questions while allowing rigorous comparison of macro- and microvascular beds within the same animal. Conduit artery studies reveal changes in global arterial mechanics and large-vessel reactivity relevant to pulse pressure and aortic stiffness, whereas resistance artery experiments probe microvascul...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
15 mL Conical Centrifuge Tubes	Thermo Fisher Scientific	14-959-70C
50 mL Conical Centrifuge Tubes	Thermo Fisher Scientific	14-959-49A
Acetic acid	Thermo Fisher Scientific	A490-212
Acetylcholine	Millipore Sigma	A6625-100G
Analytical Balance	Mettler Toledo	MX304
CaCl2	Thermo Fisher Scientific	C79-500
Carbogen (95% O2/5% CO2)	Airgas	CD USP50
Dissection Microscope	ZEISS	ZEISS Stemi 305
Dissection petri dish	Living Systems Instrumentation	DD-90-S-BLK
Dissection pins	Danish Myo Technology	400626
Dissection tool kit	Danish Myo Technology	101017	Kit consists of dissection scissor (1), multi-purpose scissor (1), Petri dishes (5), forceps (3) and dissection pins.
Dumont #5 - Fine Forceps	Fine Science Tools	11254-20
EDTA	Thermo Fisher Scientific	BP2482-500
Freezer (-20 °C)	Thermo Fisher Scientific	TSG2325FA
Freezer Hinged Boxes	USA Scientific	2310-5848
Gloves (Large)	Thermo Fisher Scientific	19-041-171D
Gloves (Medium)	Thermo Fisher Scientific	19-041-171C
Gloves (Small)	Thermo Fisher Scientific	19-041-171B
Glucose	Thermo Fisher Scientific	D16-500
Graduated Cylinder	Thermo Fisher Scientific	08-572G
GraphPad Prism	GraphPad Software	N/A	Data analysis and graphical presentation
Hydrochloric Acid Solution	Thermo Fisher Scientific	SA48-1
Ice Bucket	Thermo Fisher Scientific	03-395-156
KCl	Thermo Fisher Scientific	P217-500
KH2PO4	Thermo Fisher Scientific	P380-500
LabChart 8 Software	AD Instruments	N/A	Data capture software
Laboratory Refrigerator	Thermo Fisher Scientific	FBG49RPGA
LED Light Source	Thermo Fisher Scientific	AMPSILED21
Magnetic Stirrer	Thermo Fisher Scientific	FB30786159
Magnetic Stirring Bars (Large)	Thermo Fisher Scientific	14-513-68
Magnetic Stirring Bars (Medium)	Thermo Fisher Scientific	14-513-61
MgSO4	Thermo Fisher Scientific	M63-500
Micro Spoon and Spatula Sampling Set	Thermo Fisher Scientific	S43322
Microcentrifuge Tubes	USA Scientific	1615-5510
Mini Microcentrifuge	Thermo Fisher Scientific	07-203-954
Mini-screwdriver	Thermo Fisher Scientific	NC1349489
Multi wire myograph system	Danish Myo Technology	DMT 620M
NaCl	Thermo Fisher Scientific	S271-3
NaHCO3	Thermo Fisher Scientific	S233-500
pH Meter	Thermo Fisher Scientific	01-912-351	Includes pH meter, LE420 sensor, installation QuickGuide, and pH buffer starter pack
Phenylephrine	Millipore Sigma	P6126-5G
Pipet Controller	Thermo Fisher Scientific	01-001-023
Pipette Tips in Racks (10 µL)	USA Scientific	1111-3810
Pipette Tips in Racks (1000 µL)	USA Scientific	1111-2830
Pipette Tips in Racks (200 µL)	USA Scientific	1111-1810
PYREX Reusable Media Storage Bottles	Thermo Fisher Scientific	06-414-1D
Serological Pipettes (10 mL)	Thermo Fisher Scientific	02-923-204
Serological Pipettes (25 mL)	Thermo Fisher Scientific	02-923-205
Single-Channel micropipette Multi-Pack	Thermo Fisher Scientific	13-684-250	3x pipettes (0.5-10 μL, 10-100 μL, 100-1000 μL)
Stainless steel wire (40 μm)	Danish Myo Technology	400447
Timer	Thermo Fisher Scientific	02-261-840
Ultra Pure Water System	Sartorius	H2O-MM-UV-SET-US
Vannas Spring Scissors	Fine Science Tools	15000-00
Volumetric Flask	Thermo Fisher Scientific	13-756-400
Vortex Mixer	Thermo Fisher Scientific	10-320-807
Water bath	Thermo Fisher Scientific	07-202-155
Weighing Dishes	Thermo Fisher Scientific	02-202-101