Name: measurement of endothelium-dependent vasorelaxation in the mouse thoracic aorta using tensometric small volume chamber myography
Uploaded: 2022-08-12T13:00:00
Description: Watch this Scientific Journal Video about Measurement of Endothelium-Dependent Vasorelaxation in the Mouse Thoracic Aorta Using Tensometric Small Volume Chamber Myography at JoVE.com

This work was supported by funding from the National Institutes of Health (R15HL145646) and Midwestern University College of Graduate Studies.

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 &#176;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 exp...

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X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optimal+initial+tension+for+rat+basilar+artery+in+wire+myography.">An optimal initial tension for rat basilar artery in wire myography.</a> Microvascular Research. 97, 156-158 (2015).</li><li>Chung, A. W., Yang, H. H., Yeung, K. 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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...

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 literature2,5.
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 &#181;m) 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.

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

measurement of endothelium-dependent vasorelaxation in the mouse thoracic aorta using tensometric small volume chamber myography

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.

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...

Watch this Scientific Journal Video about Measurement of Endothelium-Dependent Vasorelaxation in the Mouse Thoracic Aorta Using Tensometric Small Volume Chamber Myography at JoVE.com

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

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.

Small volume chamber tensometric myography is a commonly used technique to evaluate the vascular contractility of small and large blood vessels in ...

We demonstrate the isometric or tensometric myography as the gold standard protocol to measure the reactivity of small resistance or large conduit blood vessels in a well-controlled physiological condition. The measurements are highly reproducible and reliable. The technique is very valuable in allowing researchers to study different layers of blood vessels separately or investigate interaction between different layers of the blood vessel wall simultaneously.Demonstrating the procedure will be Robert Folk, the research lab specialist from my laboratory. Begin by turning on the water bath and setting it to 37 degrees Celsius. Place two beakers labeled appropriately into the water bath, one with 600 milliliters of HEPES-PSS solution and one with 150 milliliters of high-potassium solution.Aerate a beaker containing 300 milliliters of HEPES-PSS solution with carbogen gas for at least 10 minutes. Add 30 milliliters of the aerated HEPES-PSS solution to a 50-milliliter centrifuge tube, label appropriately, and place it on ice. Keep the remaining aerated HEPES-PSS solution on ice to be used during the aortic dissection process.Turn on the four-chamber myograph unit at 5 milliliters of HEPES-PSS solution to each myograph chamber and set the heat to 37 degrees Celsius. Allow the solution in each chamber to be aerated for 30 minutes and check to ensure that the chambers have reached the desired 37 degrees Celsius. Turn on the myograph data acquisition hardware and computer.Transfer the thoracic cage excised from the euthanized mice to a clear silicone elastomer-coated Petri dish filled with ice cold aerated HEPES-PSS buffer and pin it on both sides. Under the microscope, gently cut and remove the heart and attached aorta from the rib cage and transfer to a clean silicone elastomer-coated dish, then gently remove fat connective tissue and clotted blood from the aorta. Using sharp small scissors, dissect and isolate the whole aorta starting from the arch area to the bottom of the descending aorta.Cut the dissected aorta into four segments of 2 millimeters each. Use the mini-ruler within the dish as a reference. In chambers already containing 5 milliliters of warmed and aerated HEPES-PSS buffer, carefully slide each of the 2-millimeter aortic segments onto the two mounting pins using forceps.When placing the chamber back in the myograph unit, slowly move the pins apart by rotating the micrometer counterclockwise so the aortic segment does not slide off the pins. Return each myograph chamber to the unit and start aerating the chambers at 37 degrees Celsius for 20 minutes. Drain the chambers and add 5 milliliters of fresh, warm, aerated HEPES-PSS solution to each chamber.If needed, readjust the force measurements to read the optimal tension of 6 millinewtons. Rest the tissue for another 15 to 20 minutes. Open the data acquisition software.Change the tracking numbers from 50 to 1 to 500 to 1 and press Start. Before starting a new experiment, be sure to add the appropriate label on the data acquisition software. Drain the chambers one more time before adding 5 milliliters of high-potassium solution to each chamber.Drain the chambers again as soon as the contractile response to the high-potassium solution reaches a plateau for force generation. Wash the tissue with HEPES-PSS solution thrice. Add 5 milliliters of high-potassium solution to each chamber.Drain the chambers as soon as the contractile response to high-potassium solution reaches a plateau for force generation and wash the tissue thrice with HEPES-PSS, then rest the tissue for 15 to 20 minutes. Pre-contract the aortic segments with the vasoconstrictor agent PE at a sub-maximum dose. Allow the PE-induced contraction curve to reach a plateau for tension development.Add a plateau of tension development to PE at 5 microliters of increasing doses of acetylcholine working stock solutions in 3-minute intervals to establish final concentrations of acetylcholine from 50 picomolar to 10 micromolar in the myograph chamber as described in the text manuscript. After completing the dose response experiment, drain the chambers, wash the aortic segments with warm and aerated HEPES-PSS solution thrice and rest the tissues for 30 minutes. After resting the aortic segments for 30 minutes, drain the chambers and add fresh, warm high-potassium solution to each chamber, then drain the chambers and wash the tissue with HEPES-PSS solution when the contraction response to high-potassium solution reaches an F plateau as demonstrated earlier.Pre-incubate the aortic segments with L-NAME by adding 10 microliters of the prepared 100-millimolar working stock solution to assess the contribution of nitric oxide production to the observed acetylcholine-induced vasorelaxation. Without removing the L name, add the sub-maximum dose of PE to the chamber to induce aortic contraction. Wait until the PE-induced contraction curve reaches a plateau for tension development, then add acetylcholine to the myograph chamber to achieve a final concentration of 500 nanomolar.Wait for a few minutes to register any possible changes in force development until the formed plateau is stable. Drain the chambers and wash the tissue with warm aerated HEPES-PSS solution thrice. Position the myograph chamber under the microscope and gently pass a small wire through the lumen of the aorta.Gently move the wire through the lumen for a short period. Cut the aorta into 2-millimeter aortic rings and mount those rings onto the myograph chamber. Set the optimal tension at 6 millinewtons and allow the tissue to rest for 30 minutes in aerated warm HEPES-PSS buffer, then drain the chambers and add 5 milliliters of fresh warm aerated HEPES-PSS solution to each chamber.Zero the force for all myograph chambers and slowly rotate the micrometer counterclockwise to increase the distance between the pins until the registered force reaches the desired optimal tension for the mouse aorta. Press the tissue for another 15 to 20 minutes at the optimal tension, then drain the chambers once more before adding 5 milliliters of high-potassium solution to each chamber. Drain the chambers again and wash the tissue with HEPES-PSS solution thrice once the contraction response to the high-potassium ion solution reaches a plateau.Rest the tissue for 15 to 20 minutes, then pre-contract the aortic segments with PE, the vasoconstrictor agent. When the PE-induced force reaches a plateau, add the sub-maximum concentration of acetylcholine to the myograph chamber to achieve a final concentration of 500 nanomolar. Wait 3 minutes to ensure that the registered plateau for PE-induced force development is not changing before ending the experiment.The integrity and viability of each isolated vessel were assessed by recording the contractile force generation. The peak of the recorded force was later used to normalize the force generation for the same segment in response to the agonists. The endothelium-mediated vasorelaxation measurements depicted smooth muscle mediated contraction and force generation.When the PE-induced contraction reached a plateau, increasing doses of acetylcholine achieved the maximum vasorelaxation in the isolated segment. The nitric oxide inhibitor L-NAME completely blocked the acetylcholine-induced vasorelaxation in the pre-contracted aorta indicating that acetylcholine induces aortic vasorelaxation through increasing nitric oxide production. Moreover, removing the endothelial layer from the aortic segments also blocked acetylcholine-induced vasorelaxation underscoring the role of the endothelium in blood vessel relaxation.Ensure to be gentle while handling, cleaning, and dissecting the blood vessel. It is critical to add the pharmacological agonists and inhibitors with care to not disturb the blood vessel rings mounted onto the myograph chambers. The information can help design experiments to explore potential therapeutic approaches to address the dysfunction of a specific blood vessel's layers.

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Small Volume (1-3L) Filtration of Coastal Seawater Samples

The workflow begins with the collection of coastal marine waters for downstream microbial community, nutrient and trace gas analyses. For today s demonstration samples were collected from the deck of the HMS John Strickland operating in Saanich Inlet. This video documents small volume (~1 L) filtration of microbial biomass from the water column. The protocol is an extension of the large volume sampling protocol described earlier, with one major difference: here, there is no pre-filtration step, so all size classes of biomass are collected down to the 0.22 &mu;m filter cut-off. Samples collected this way are ideal for nucleic acid analysis. The set-up, filtration, and clean-up steps each take about 20-30 minutes. If using two peristaltic pumps simultaneously, up to 8 samples may be filtered at the same time. To prevent biofilm formation between sampling trips, all filtration equipment must be rinsed with dilute HCl and deionized water and autoclaved immediately after use.

Small Volume 1-3L Filtration of Coastal Seawater Samples

This video documents small volume (~1 L) filtration of microbial biomass from the water column.

The workflow begins with the collection of coastal marine waters for downstream microbial community, nutrient and trace gas analyses. For today s ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with a molecular mass smaller than 1.5 kDa. Although the definitive molecular identity of the pore is still under debate, proteins such as cyclophilin D, VDAC and ANT contribute to mtPTP formation. While the involvement of mtPTP opening in cell death is well established1, accumulating evidence indicates that the mtPTP serves a physiologic role during mitochondrial Ca2+ homeostasis2, bioenergetics and redox signaling 3.
mtPTP opening is triggered by matrix Ca2+ but its activity can be modulated by several other factors such as oxidative stress, adenine nucleotide depletion, high concentrations of Pi, mitochondrial membrane depolarization or uncoupling, and long chain fatty acids4. In vitro, mtPTP opening can be achieved by increasing Ca2+ concentration inside the mitochondrial matrix through exogenous additions of Ca2+ (calcium retention capacity). When Ca2+ levels inside mitochondria reach a certain threshold, the mtPTP opens and facilitates Ca2+ release, dissipation of the proton motive force, membrane potential collapse and an increase in mitochondrial matrix volume (swelling) that ultimately leads to the rupture of the outer mitochondrial membrane and irreversible loss of organelle function.
Here we describe a fluorometric assay that allows for a comprehensive characterization of mtPTP opening in isolated mouse heart mitochondria. The assay involves the simultaneous measurement of 3 mitochondrial parameters that are altered when mtPTP opening occurs: mitochondrial Ca2+ handling (uptake and release, as measured by Ca2+ concentration in the assay medium), mitochondrial membrane potential, and mitochondrial volume. The dyes employed for Ca2+ measurement in the assay medium and mitochondrial membrane potential are Fura FF, a membrane impermeant, ratiometric indicator which undergoes a shift in the excitation wavelength in the presence of Ca2+, and JC-1, a cationic, ratiometric indicator which forms green monomers or red aggregates at low and high membrane potential, respectively. Changes in mitochondrial volume are measured by recording light scattering by the mitochondrial suspension. Since high-quality, functional mitochondria are required for the mtPTP opening assay, we also describe the steps necessary to obtain intact, highly coupled and functional isolated heart mitochondria.

A spectrofluorometric protocol for the measurement of the mitochondrial permeability transition pore opening in isolated mouse heart mitochondria is presented here. The assay involves the simultaneous measurement of mitochondria Ca2+ handling, mitochondrial membrane potential and mitochondrial volume. The procedure for obtaining high-quality and functional heart mitochondria is also described.

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with ...

Assessing Murine Resistance Artery Function Using Pressure Myography

Pressure myograph systems are exquisitely useful in the functional assessment of small arteries, pressurized to a suitable transmural pressure. The near physiological condition achieved in pressure myography permits in-depth characterization of intrinsic responses to pharmacological and physiological stimuli, which can be extrapolated to the in vivo behavior of the vascular bed. Pressure myograph has several advantages over conventional wire myographs. For example, smaller resistance vessels can be studied at tightly controlled and physiologically relevant intraluminal pressures. Here, we study the ability of 3rd order mesenteric arteries (3-4 mm long), preconstricted with phenylephrine, to vaso-relax in response to acetylcholine. Mesenteric arteries are mounted on two cannulas connected to a pressurized and sealed system that is maintained at constant pressure of 60 mmHg. The lumen and outer diameter of the vessel are continuously recorded using a video camera, allowing real time quantification of the vasoconstriction and vasorelaxation in response to phenylephrine and acetylcholine, respectively.
	To demonstrate the applicability of pressure myography to study the etiology of cardiovascular disease, we assessed endothelium-dependent vascular function in a murine model of systemic hypertension. Mice deficient in the &alpha;1 subunit of soluble guanylate cyclase (sGC&alpha;1-/-) are hypertensive when on a 129S6 (S6) background (sGC&alpha;1-/-S6) but not when on a C57BL/6 (B6) background (sGC&alpha;1-/-B6). Using pressure myography, we demonstrate that sGC&alpha;1-deficiency results in impaired endothelium-dependent vasorelaxation. The vascular dysfunction is more pronounced in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice, likely contributing to the higher blood pressure in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice. 
	Pressure myography is a relatively simple, but sensitive and mechanistically useful technique that can be used to assess the effect of various stimuli on vascular contraction and relaxation, thereby augmenting our insight into the mechanisms underlying cardiovascular disease.

In pressure myography, an intact small segment of a vessel is mounted onto two small cannulas and pressurized to a suitable luminal pressure. Here, we describe the method to measure vasorelaxation response of the mouse 3rd order mesenteric arteries in c57 and sGC&alpha;1-/- mice using pressure myography.

Pressure myograph systems are  exquisitely useful in the functional assessment of small arteries, pressurized  to a suitable transmural pressure. The  ...

Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular systolic and diastolic function. Measures of ventricular contractility and compliance are obtained through examination of cardiac response to changes in afterload and preload. These techniques were originally developed nearly three decades ago to measure cardiac function in large mammals and humans. The application of these analyses to small mammals, such as mice, has been accomplished through the optimization of microsurgical techniques and creation of conductance catheters. Conductance catheters allow for estimation of the blood pool by exploiting the relationship between electrical conductance and volume. When properly performed, these techniques allow for testing of cardiac function in genetic mutant mouse models or in drug treatment studies. The accuracy and precision of these studies are dependent on careful attention to the calibration of instruments, systematic conduct of hemodynamic measurements and data analyses. We will review the methods of conducting pressure-volume loop experiments using a conductance catheter in mice.

Cardiac pressure-volume loop analysis is the most comprehensive way to measure cardiac function in the intact heart. We describe a technique to perform and analyze cardiac pressure volume loops, using conductance catheters.

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular ...

Measuring Pressure Volume Loops in the Mouse

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the mouse provides great potential to explore cardiac function at the molecular level. The mouse&#39;s small size does present some challenges in regards to performing detailed cardiac phenotyping. Miniaturization and other advancements in technology have made many methods of cardiac assessment possible in the mouse. Of these, the simultaneous collection of pressure and volume data provides a detailed picture of cardiac function that is not available through any other modality. Here a detailed procedure for the collection of pressure-volume loop data is described. Included is a discussion of the principles underlying the measurements and the potential sources of error. Anesthetic management and surgical approaches are discussed in great detail as they are both critical to obtaining high quality hemodynamic measurements. The principles of hemodynamic protocol development and relevant aspects of data analysis are also addressed.

This manuscript describes a detailed protocol for the collection of pressure-volume data from the mouse.

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Volume Segmentation and Analysis of Biological Materials Using SuRVoS (Super-region Volume Segmentation) Workbench

Segmentation is the process of isolating specific regions or objects within an imaged volume, so that further study can be undertaken on these areas of interest. When considering the analysis of complex biological systems, the segmentation of three-dimensional image data is a time consuming and labor intensive step. With the increased availability of many imaging modalities and with automated data collection schemes, this poses an increased challenge for the modern experimental biologist to move from data to knowledge. This publication describes the use of SuRVoS Workbench, a program designed to address these issues by providing methods to semi-automatically segment complex biological volumetric data. Three datasets of differing magnification and imaging modalities are presented here, each highlighting different strategies of segmenting with SuRVoS. Phase contrast X-ray tomography (microCT) of the fruiting body of a plant is used to demonstrate segmentation using model training, cryo electron tomography (cryoET) of human platelets is used to demonstrate segmentation using super- and megavoxels, and cryo soft X-ray tomography (cryoSXT) of a mammalian cell line is used to demonstrate the label splitting tools. Strategies and parameters for each datatype are also presented. By blending a selection of semi-automatic processes into a single interactive tool, SuRVoS provides several benefits. Overall time to segment volumetric data is reduced by a factor of five when compared to manual segmentation, a mainstay in many image processing fields. This is a significant savings when full manual segmentation can take weeks of effort. Additionally, subjectivity is addressed through the use of computationally identified boundaries, and splitting complex collections of objects by their calculated properties rather than on a case-by-case basis.

Volume Segmentation and Analysis of Biological Materials Using SuRVoS Super-region Volume Segmentation Workbench

Segmentation of three-dimensional data from many imaging techniques is a major bottleneck in analysis of complex biological systems. Here, we describe the use of SuRVoS Workbench to semi-automatically segment volumetric data at various length-scales using example datasets from cryo-electron tomography, cryo soft X-ray tomography, and phase contrast X-ray tomography techniques.

Segmentation is the process of isolating specific regions or objects within an imaged volume, so that further study can be undertaken on these areas of ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...