This work is supported by NIH grants, K08135443 (Y.B), 1R01HL132991 (X.A).

All animal care was in accordance with the guidelines of the Institutional Animal Care and Use Committee of Massachusetts General Hospital. Wild-type C57/B6 male mice, 8 weeks of age, were used for the present study.
1. Experimental preparation
<ol>
	<li>Prepare the working solution.
	<ol>
		<li>Prepare 1x Hank&#39;s Balanced Salt Solution (HBSS, with Ca2+ and Mg2+, and pH balanced with 20 mM HEPES, see Table of Materials). Use the HBSS solution to prepare and process the PCLS. Keep HBSS solution cold on ice when preparing PCLSs.</li>
	</ol>
	</li>
	<li>Prepare the culture medium of PCLS by supplementing Dulbecco&#39;s Modified Eagle Medium and F-12 (DMEM-F12) with an antibiotic-antimycotic agent (1:100 volume ratio, see Table of Materials).</li>
	<li>Prepare 1.5% agarose and 6% gelatin solution.
	<ol>
		<li>Mix low melting point (LMP) agarose or gelatin powder (see Table of Materials) with HBSS solution separately in 15 mL sterile centrifuge tubes (or 50 mL tubes if solution volume &#62;10 mL) to final concentrations. 
		NOTE: Total volume of agarose solution = 5 mL/mouse x number of mice; gelatin solution = 2 mL/mouse x number of mice.</li>
		<li>Heat the solution tubes in boiling water until the powder completely dissolves. Keep both agarose and gelatin solutions in a 42 &#176;C water bath. 
		NOTE: A heating lamp at the dissection table can be useful to keep the operating environment warm and prevent agarose solution from solidifying before being injected into the mouse lung.</li>
	</ol>
	</li>
	<li>Submerge all dissection tools, including a pair of dissecting scissors, two pairs of curved micro-dissecting forceps, and a pair of hemostatic forceps in 70% ethanol solution for at least 20 min for sterilization.</li>
	<li>Keep a vibratome ready (see Table of Materials) to section the lung tissue into thin slices.</li>
	<li>Keep an inverted phase-contrast microscope with a CCD camera to evaluate the airway or vascular contractile responses and a custom-built laser scanning confocal microscope (see Table of Materials) to assess the Ca2+ signaling in pulmonary SMCs18.</li>
</ol>
2. Inflation of mouse lungs with agarose and gelatin solution
<ol>
	<li>Euthanize the mouse.
	<ol>
		<li>Place the mouse in a plastic chamber (around 750 cm3) with 5% isoflurane. Keep the mouse in the chamber until it stops breathing for at least 1 min. 
		NOTE: Other primary euthanasia methods, such as intraperitoneal injection of ketamine (240 mg/Kg) and xylazine (32 mg/Kg)19 or pentobarbital (0.3 mL)20, can be applied as alternatives to inhaled isoflurane.</li>
		<li>Place the mouse body on a dissection board in the supine position. Fix the body in position by pinning down the tail, front paws, and head with 25 G syringe needles, and sanitize the body with 70% ethanol spray.</li>
		<li>Cut open the mouse&#39;s abdomen with surgical scissors (incision, ~2 cm long x 2 cm wide). Then, cut off the abdominal aorta to deplete the blood volume.</li>
	</ol>
	</li>
	<li>Inflate lung lobes following the steps below.
	<ol>
		<li>Open the chest cavity carefully along the sternum and bilateral inferior rib cage above the diaphragm, and observe the lung lobes collapse when the chest cavity opens.</li>
		<li>Remove part of bilateral ventral rib cages to expose the heart. Avoid lung damage by pointing the sharp scissor tip away from the lung tissue.</li>
		<li>Use forceps to separate the thymus and soft tissue in the mouse neck to expose the trachea.</li>
		<li>Cut a small hole (1.2 mm in diameter) in the upper end of the trachea allowing passage of the tip of a 20 G Y-shaped IV catheter (see Table of Materials).</li>
		<li>Connect one adaptor port of the Y-shaped catheter with a 3 mL syringe prefilled with 0.5 mL of air, and the other port with a 3 mL syringe prefilled with 2 mL of warm 1.5% agarose solution (42 &#176;C).</li>
		<li>Inject agarose solution to fill the catheter and then push the catheter through the pre-cut opening into the trachea for 5-8 mm in length.</li>
		<li>Slowly inject the agarose solution at around 1 mL/5 s. The lung will expand along the proximal-to-distal axis. Stop injection when the edge of each lung lobe is inflated. 
		NOTE: Do not overinflate the lung, as this could rupture it. The volume of agarose solution to inflate the whole lung of a young adult mouse is about 1.3 &#177; 0.1 mL.</li>
		<li>Inject 0.2-0.3 mL of air from the other syringe to push the residual agarose in the catheter and conductive airways to the distal alveoli space.</li>
		<li>Clip close the trachea with a pair of curved hemostatic forceps (see Table of Materials).</li>
	</ol>
	</li>
	<li>Infill the pulmonary vasculature.
	<ol>
		<li>Fill a 1 mL syringe with warm 6% gelatin. Connect to a needle scalp vein catheter, infill the catheter with gelatin solution, and then puncture the right ventricle close to the inferior wall with the needle.</li>
		<li>Push the needle into the right ventricle for 2-3 mm in length and point the needle tip to the main pulmonary artery.</li>
		<li>Inject 0.2 mL of gelatin solution slowly into the right ventricle and pulmonary arterial vessels. 
		NOTE: The lung lobes appear slightly paler with appropriate gelatin inflation.</li>
		<li>Keep the needle in place for 5 min after injection, cool the lung lobes by pouring ice-cold HBSS solution on the heart and lung, then place the body with the dissection board in a 4 &#176;C refrigerator or on ice for a total of 10 min.</li>
		<li>Remove the mouse lung and heart from surrounding connective tissues with scissors. Then, separate each lung lobe and keep them in HBSS solution on ice.</li>
	</ol>
	</li>
</ol>
3. Sectioning of lung lobes to thin slices
<ol>
	<li>Trim and attach the lung lobe on the top of the specimen column with superglue, and orient the lobe (Figure 1A,B) to allow the cutting direction to be perpendicular to most airways from the hila to the lung surface.</li>
	<li>Use a vibratome with a fresh thin razor blade to cut the lung lobe into 150 &#181;m slices. Collect the slices in 100 mm sterile Petri dishes prefilled with cold HBSS solution. 
	NOTE: Set appropriate blade moving speed and oscillation frequency before starting slicing. A typical setting for a Compresstom is Speed level 4 and Oscillation level 4.</li>
	<li>Transfer slices to Petri dishes filled with DMEM/F-12 culture medium (20 slices/15 mL medium/dish) and maintain them in a 37 &#176;C incubator overnight before experiments. 
	NOTE: Mouse PCLSs can be maintained in culture for about 7 days without losing airway contractile responses17. The longevity of pulmonary arteries has not been thoroughly evaluated. In our observation, they retain intact structure and vasoreaction for at least 3 days in DMEM/F12 medium. For long-term storage, mouse PCLSs can be placed in cryovials filled with DMEM/F12 medium containing 10% DMSO (5 slices in 1 mL medium per vial), frozen in an isopropyl alcohol-filled container at -80 &#176;C overnight, and cryopreserved in liquid nitrogen for weeks to months21.</li>
</ol>
4. Analyzing contractile responses of intrapulmonary airways and arteries
<ol>
	<li>Place PCLSs in an HBSS-filled 24-well culture plate, one in each well. Locate the PCLS in the middle of the well, then remove the HBSS solution with a pipette.</li>
	<li>Find the target airway and vessel in the slice under the microscope, then cover the slice using a nylon mesh with a pre-cut central hole (2-3 mm in diameter) to expose the target area.</li>
	<li>Lay down a hollow metal washer on top of the mesh to hold the slices in place (Figure 2A).</li>
	<li>Add 600 &#956;L of HBSS solution to submerge the PCLS. Rest the slice for 10 min, then record the baseline images of airways or vessels.</li>
	<li>Induce the airway or vascular contraction by cautiously suctioning out the blank HBSS solution with a pipette and adding 600 &#956;L of HBSS with an agonist, such as 1 &#181;M of methacholine (MCh) or 10 nM of endothelin (see Table of Materials). 
	NOTE: KCl stimulation as a tool-set standard is not required prior to methacholine or endothelin exposure. 100 mM of KCl stimulation in mouse airways only induces a small and irregular airway contraction (10%-15%)20. Likewise, a priming methacholine stimulation is not obligated as we have confirmed that the same agonist, for instance, methacholine or serotonin, triggers similar airway contraction on the first and repetitive exposures20,22.</li>
	<li>Observe the reaction under the microscope until the luminal area change reaches an equilibratory status, and then record the airway or vascular images for analysis.</li>
	<li>Remove MCh or endothelin solution with a pipette and add a new 600 &#181;L HBSS solution with the same agonist concentration and a relaxant; observe and record the airway or vascular relaxation.</li>
	<li>Quantify the airway or vascular reaction following the steps below.
	<ol>
		<li>Load the images to NIH-sponsored free software FIJI.</li>
		<li>Select the Polygon Selection in the toolbar to outline the airway or vascular lumen on each frame.</li>
		<li>Choose Analyze &#62; Set Measurements &#62; Area.</li>
		<li>Choose Analyze &#62; Measure to quantify the area of interest. Results are shown in a separate window for statistical analysis.</li>
	</ol>
	</li>
</ol>
5. Analyzing Ca2+ signaling of airway or vascular SMCs
<ol>
	<li>Prepare the Ca2+ dye loading buffer.
	<ol>
		<li>Dissolve Ca2+ dye, Oregon Green 488 BAPTA-1-AM (see Table of Materials), 50 &#181;g (one vial) with 10 &#181;L of DMSO.</li>
		<li>Dissolve 0.2 g of Pluronic F-12 powder (see Table of Materials) in 1 mL of DMSO to generate a 20% Pluronic solution.</li>
		<li>Mix 10 &#181;L of 20% Pluronic solution with 10 &#181;L of Ca2+ dye-DMSO solution.</li>
		<li>Make Ca2+ dye loading buffer by adding 20 &#181;L of the mixture to 2 mL of HBSS solution with 200 &#181;M of sulfobromophthalein (see Table of Materials).</li>
	</ol>
	</li>
	<li>Place 15 mouse PCLSs in 2 mL of Ca2+ loading buffer and incubate at 30 &#176;C for 1 h. Then move the slices to 2 mL of HBSS solution with 100 &#181;M of sulfobromophthalein for an additional 30 min for fluorescent dye de-esterification at room temperature.</li>
	<li>Detect Ca2+ signaling of SMCs.
	<ol>
		<li>Place Ca2+ dye-loaded slices on a large cover glass.</li>
		<li>Fill a 3 mL syringe with high vacuum silicone grease. Squeeze the grease through an 18 G blunted needle to draw two parallel lines across the cover glass above and below the slice.</li>
		<li>Cover the slice using a nylon mesh between two grease lines. Place the second cover glass on top of the mesh to generate a chamber sealed by grease on the sides (Figure 3A). 
		NOTE: The nylon mesh is essential to keep the slice attached to the bottom coverslip and thus stay within the working distance of a high magnitude inverted objective, such as 40x oil.</li>
		<li>Add HBSS or agonist solution into the chamber from one end by pipetting manually or via a perfusion system. Remove the fluid from the chamber by suctioning from the other end with tissue paper or vacuum in the case of continuous fluid perfusion.</li>
		<li>Put the PCLS chamber on the microscope stage. Detect the Ca2+ signaling of SMC23 with a custom-built resonant scanning confocal microscope with a laser scan rate of 15 or 30 images/s. 
		NOTE: Alternatively, a high-speed commercial laser scanning confocal microscope, widely available at present, can be applied in the Ca2+ signaling assay of pulmonary SMCs in PCLS.</li>
	</ol>
	</li>
	<li>Quantify the Ca2+ fluorescence in SMCs.
	<ol>
		<li>Load the recorded image sequence to FIJI and choose the Image &#62; Type &#62; 8-bit grayscale.</li>
		<li>Select the Rectangle Selection in the toolbar and define a 5 pixel x 5 pixel region of interest (ROI) in a smooth muscle cell.</li>
		<li>Choose Analyze &#62; Set Measurements &#62; Mean gray value, representing the Ca2+ fluorescence intensity.</li>
		<li>If the position of an ROI changes with SMC contraction, choose Analyze &#62; Measure the Ca2+ fluorescent intensity frame-by-frame.</li>
		<li>If the ROI of SMC stays in a similar position in a stack of images, choose Image &#62; Stacks &#62; Measure stack of the Ca2+ fluorescent intensity. 
		NOTE: A custom-written macro can be plugged in to track the ROI within the SMC when it moves with contraction in an image stack and automatically measures the Ca2+ intensity (gray value) of ROI frame-by-frame.</li>
	</ol>
	</li>
</ol>

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L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+rhinovirus+39+infection+on+airway+hyperresponsiveness+to+carbachol+in+human+airways+precision+cut+lung+slices.">Effects of rhinovirus 39 infection on airway hyperresponsiveness to carbachol in human airways precision cut lung slices.</a> The Journal of Allergy and Clinical Immunology. 141 (5), 1887-1890 (2018).</li><li>Bai, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreserved+Human+precision-cut+lung+slices+as+a+bioassay+for+live+tissue+banking.+a+viability+study+of+bronchodilation+with+bitter-taste+receptor+agonists.">Cryopreserved Human precision-cut lung slices as a bioassay for live tissue banking. a viability study of bronchodilation with bitter-taste receptor agonists.</a> American Journal of Respiratory Cell and Molecular Biology. 54 (5), 656-663 (2016).</li><li>Mondo&#241;edo, J. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+high-throughput+system+for+cyclic+stretching+of+precision-cut+lung+slices+during+acute+cigarette+smoke+extract+exposure.">A high-throughput system for cyclic stretching of precision-cut lung slices during acute cigarette smoke extract exposure.</a> Frontiers in Physiology. 11, 566 (2020).</li><li>Davidovich, N., Huang, J., Margulies, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reproducible+uniform+equibiaxial+stretch+of+precision-cut+lung+slices.">Reproducible uniform equibiaxial stretch of precision-cut lung slices.</a> American Journal of Physiology-Lung Cellular and Molecular Physiology. 304 (4), 210-220 (2013).</li><li>Ram-Mohan, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+traction+microscopy+to+quantify+muscle+contraction+within+precision-cut+lung+slices.">Tissue traction microscopy to quantify muscle contraction within precision-cut lung slices.</a> American Journal of Physiology-Lung Cellular and Molecular Physiology. 318 (2), 323-330 (2020).</li></ol>

The authors have nothing to disclose.

The preparation of PCLS involves several critical steps. First, it is essential to inflate the lung lobe homogeneously to avoid the variation of tissue stiffness from uneven agarose distribution. As the liquid agarose rapidly gels in thin catheters or airways at a temperature below 37 &#176;C, the resultant filling defect in the distal lung field could increase the disparity of lung tissue stiffness and cause tissue tearing during the vibratome section. Therefore, keeping the low-melting agarose solution at 42 &#176;C in...

Smooth muscle cell (SMC) is a major structural cell type in the lung, primarily residing in the media wall of airways and pulmonary vessels. SMCs contract to alter the luminal caliber, thus regulating air and blood flow1,2. Therefore, contractile regulation of SMCs is essential to maintain the homeostasis of air ventilation and pulmonary circulation. In contrast, aberrant SMC contractility provokes obstructive airway or pulmonary vascular diseases like asthma and pulmonary arterial hypertension. However, the functional assessment of lung SMCs has been challenged by limited access to the lung tissue, especially those small airways and microvessels in the distal part of the lung2,3. Current solutions resort to indirect assays, such as measuring airflow resistance by Flexivent to reflect airway constriction, and checking pulmonary arterial blood pressure by right heart catheterization to assess pulmonary vasocontraction4,5. However, these indirect assays have multiple disadvantages, such as being confounded by structural factors, failing to capture the spatial diversity of airway or vascular responses in the whole lung scale6,7, and unfitting for the mechanistic study of contractile regulation at the cellular level. Therefore, alternative approaches using isolated primary cells, trachea/bronchi muscle strips8,9, or large vascular segments10 have been applied for the SMC study in vitro. Nevertheless, these methods also have limitations. For example, a quick phenotypical adaptation of primary SMCs in the culture condition11,12 makes it problematic to extrapolate findings from cell culture to in vivo settings. In addition, the contractile phenotype of SMCs in the isolated proximal airway or vascular segments may not represent the SMCs in the distal lung6,7. Moreover, the muscle force measurement at the tissue level remains dissociated from molecular and cellular events that are essential for mechanistic insight into contractile regulation.
Precision-cut lung slice (PCLS), a live lung tissue section, provides an ideal ex vivo tool to characterize pulmonary SMCs in a near in vivo microenvironment (i.e., preserved multi-cellular architecture and interaction)13. Since Drs. Placke and Fisher first introduced the preparation of lung slices from agarose-inflated rat and hamster lungs in the 1980s14,15, this technique has been advanced continuously to provide PCLSs with higher quality and greater versatility for biomedical research. One significant improvement is the enhancement of pulmonary arterial preservation by gelatin infusion in addition to lung inflation with agarose via the trachea. As a result, both the airway and pulmonary arteries are kept intact in the PCLS for ex vivo assessement16. Furthermore, the PCLS is viable for a prolonged time in culture. For instance, mouse PCLSs had no significant change in cell viability and metabolism for a minimum of 12 days in culture, as well as, they retained airway contractility for up to 7 days17. In addition, PCLS keeps different-sized airways or vessels for contraction and relaxation assays. Moreover, intracellular Ca2+ signaling of SMCs, the determinant factor of cell contractility, can be assayed with Ca2+ reporter dyes imaged by a confocal or 2-photon microscope13.
Considering the extensive application of the mouse model in lung research, a detailed protocol is described here for preparing mouse PCLS with intact airways and intrapulmonary arteries for ex vivo lung research. Using the prepared PCLSs, we subsequently demonstrated how to evaluate the airway and pulmonary arterial responses to constrictive or relaxant stimuli. In addition, the method of loading the PCLS with Ca2+ reporter dye and then imaging Ca2+ signaling of SMCs associated with contractile or relaxant responses are also described.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL syringe</td>
			<td>BD</td>
			<td>309626</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL sterile centrifuge tubes</td>
			<td>Celltreat</td>
			<td>229411</td>
			<td></td>
		</tr>
		<tr>
			<td>3 mL syringe</td>
			<td>BD</td>
			<td>309585</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL sterile centrifuge tubes</td>
			<td>Celltreat</td>
			<td>229422</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetyl-beta-methacholine</td>
			<td>Millipore Sigma</td>
			<td>62-51-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-anitmycotic</td>
			<td>Thermo Fisher</td>
			<td>15240-062</td>
			<td></td>
		</tr>
		<tr>
			<td>CCD-camera</td>
			<td>Nikon</td>
			<td>Nikon Ds-Ri2 camera</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover glassess</td>
			<td>Fisher Scientific</td>
			<td>12-548-5CP; 12-548-5PP</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryogenic vials</td>
			<td>Fisher Scientific</td>
			<td>430488</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom-built laser scanning confocal microscope</td>
			<td></td>
			<td></td>
			<td>Details in Reference 18</td>
		</tr>
		<tr>
			<td>DMEM/F12</td>
			<td>Fisher Scientific</td>
			<td>MT-10-092-CM</td>
			<td></td>
		</tr>
		<tr>
			<td>Endothelin 1</td>
			<td>Millipore Sigma</td>
			<td>E7764</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine dissecting scissor</td>
			<td>Fisher Scientific</td>
			<td>NC9702861</td>
			<td></td>
		</tr>
		<tr>
			<td>Freezing container</td>
			<td>Sigma-Aldrich</td>
			<td>C1562</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin from porcine skin</td>
			<td>Sigma-Aldrich</td>
			<td>9000-70-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks&#39; Balanced Salt Solution (HBSS)</td>
			<td>Thermo Fisher</td>
			<td>14025092</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemostatic forcep</td>
			<td>Fisher Scientific</td>
			<td>16-100-117</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Thermo Fisher</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>High vaccum silicone grease</td>
			<td>Fisher Scientific</td>
			<td>146355d</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropyl alcohol</td>
			<td>Sigma-Aldrich</td>
			<td>W292907-1KG-K</td>
			<td></td>
		</tr>
		<tr>
			<td>Metal washers</td>
			<td>Home Depot Product Authority</td>
			<td>800442</td>
			<td>Everbilt Flat Washers #10</td>
		</tr>
		<tr>
			<td>Micro-dissecting forcep</td>
			<td>Sigma-Aldrich</td>
			<td>F4142</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle scalp vein set (25 G)</td>
			<td>EXELINT</td>
			<td>26708</td>
			<td></td>
		</tr>
		<tr>
			<td>NOC-5</td>
			<td>Cayman Chemical</td>
			<td>16534</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon mesh</td>
			<td>Component Supply</td>
			<td>U-CMN-300</td>
			<td></td>
		</tr>
		<tr>
			<td>Oregon green 488 BAPTA-1 AM</td>
			<td>Life Technologies</td>
			<td>o-6807</td>
			<td></td>
		</tr>
		<tr>
			<td>Phase-contrast microscope</td>
			<td>Nikon</td>
			<td>Nikon Eclipse TS 100</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Thermo Fisher</td>
			<td>P-6867</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor blades</td>
			<td>Personna</td>
			<td></td>
			<td>Personna Double Edge Razor Blades in White Wrapper 100 count</td>
		</tr>
		<tr>
			<td>Sulfobromophthalein</td>
			<td>Sigma-Aldrich</td>
			<td>S0252</td>
			<td></td>
		</tr>
		<tr>
			<td>Superglue</td>
			<td>Krazy Glue</td>
			<td></td>
			<td>Krazy Glue, All purpose</td>
		</tr>
		<tr>
			<td>Ultrapure low melting point agarose</td>
			<td>Thermo Fisher</td>
			<td>16520050</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Precisionary</td>
			<td>VF 310-0Z</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome chilling block</td>
			<td>Precisionary</td>
			<td>SKU-VM-CB12.5-NC</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome specimen tube</td>
			<td>Precisionary</td>
			<td>SKU VF-SPS-VM-12.5-NC</td>
			<td></td>
		</tr>
		<tr>
			<td>Y shaped IV catheter</td>
			<td>BD</td>
			<td>383336</td>
			<td>BD Saf-T-Intima closed IV catheter</td>
		</tr>
	</tbody>
</table>

utilizing the precision-cut lung slice to study the contractile regulation of airway and intrapulmonary arterial smooth muscle

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, respectively, hence playing a critical role in the homeostasis of the pulmonary system. Deregulation of SMC contractility contributes to several pulmonary diseases, including asthma and pulmonary hypertension. However, due to limited tissue access and a lack of culture systems to maintain in vivo SMC phenotypes, molecular mechanisms underlying the deregulated SMC contractility in these diseases remain fully identified. The precision-cut lung slice (PCLS) offers an ex vivo model that circumvents these technical difficulties. As a live, thin lung tissue section, the PCLS retains SMC in natural surroundings and allows in situ tracking of SMC contraction and intracellular Ca2+ signaling that regulates SMC contractility. Here, a detailed mouse PCLS preparation protocol is provided, which preserves intact airways and intrapulmonary arteries. This protocol involves two essential steps before subjecting the lung lobe to slicing: inflating the airway with low-melting-point agarose through the trachea and infilling pulmonary vessels with gelatin through the right ventricle. The PCLS prepared using this protocol can be used for bioassays to evaluate Ca2+-mediated contractile regulation of SMC in both the airway and the intrapulmonary arterial compartments. When applied to mouse models of respiratory diseases, this protocol enables the functional investigation of SMC, thereby providing insight into the underlying mechanism of SMC contractility deregulation in diseases.

Mouse PCLS preparation preserving intact intrapulmonary airways and arteries 
A 150 &#181;m thick PCLS was observed under the inverted phase-contrast microscope. In mouse lungs, conductive airways are accompanied by intrapulmonary arteries, running from the hilus to the peripheral lung. A representative pulmonary airway-artery bundle in a mouse PCLS is shown in Figure 2B. The airway can be easily identified by cuboidal epithelial cells with active cilial beating lining ...

Watch this Scientific Journal Video about Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle at JoVE.com

Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle

The present protocol describes preparing and utilizing mouse precision-cut lung slices to assess the airway and intrapulmonary arterial smooth muscle contractility in a nearly in vivo milieu.

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, ...

utilizing-precision-cut-lung-slice-to-study-contractile-regulation

Research

JoVE Journal

Biology

3.3K Views.  Massachusetts General Hospital and Harvard Medical School. The present protocol describes preparing and utilizing mouse precision-cut lung slices to assess the airway and intrapulmonary arterial smooth muscle contractility in a nearly in vivo milieu.

Video: Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

Badania wizualne Wrażliwość na prędkość i kierunek ruchu w Jaszczurki

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of ...

Computer-Generated Animal Model Stimuli

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In particular, understanding the constraints on visual signal design for communication has been of great interest. Traditional methods for investigating animal interactions have used basic observational techniques, staged encounters, or physical manipulation of morphology. Less intrusive methods have tried to simulate conspecifics using crude playback tools, such as mirrors, still images, or models. As technology has become more advanced, video playback has emerged as another tool in which to examine visual communication (Rosenthal, 2000). However, to move one step further, the application of computer-animation now allows researchers to specifically isolate critical components necessary to elicit social responses from conspecifics, and manipulate these features to control interactions. Here, I provide detail on how to create an animation using the Jacky dragon as a model, but this process may be adaptable for other species. In building the animation, I elected to use Lightwave  3D to alter object morphology, add texture, install bones, and provide comparable weight shading that prevents exaggerated movement. The animation is then matched to select motor patterns to replicate critical movement features. Finally, the sequence must rendered into an individual clip for presentation. Although there are other adaptable techniques, this particular method had been demonstrated to be effective in eliciting both conspicuous and social responses in staged interactions.

Computer-generated stimuli using the Jacky dragon as a model.

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In ...

Focal Ca2+ Transient Detection in Smooth Muscle

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ from internal stores, and allows multiple cells to be monitored simultaneously to assess, for example, coupling in syncytial tissue. Subcellular Ca2+ transients are common in smooth muscle, yet are difficult to measure accurately because of the problems caused by their stochastic occurrence, over an often wide field of view, in an organ that it prone to contract. To overcome this problem, we've developed a series of imaging protocols and analysis routines to acquire and then analyse, in an automated fashion, the frequency, location and amplitude of such events. While this approach may be applied in other contexts, our own work involves the detection of local purinergic Ca2+ transients for locating transmitter release with submicron resolution. 
ATP is released as a cotransmitter from autonomic nerves, where it binds to P2X1 receptors on the smooth muscle of the detrusor and vas deferens. Ca2+ enters the smooth muscle, resulting in purinergic neuroeffector Ca2+ transients (NCTs). The focal Ca2+ transients allow the optical monitoring of neurotransmitter release in a manner that has many advantages over electrophysiology. Apart from the greatly improved spatial resolution, optical recording has the additional advantage of allowing the recording of transmitter release from many distinguishable sites simultaneously. Furthermore, the optical plane of focus is easier to maintain or correct during long recording series than is the repositioning of an intracellular sharp microelectrode. 
In summary, a method for imaging of Ca2+ fluorescence is outlined which details the preparation of tissue, and the acquisition and analysis of data. We outline the use of several scripts for the analysis of such Ca2+ transients.

Focal Ca2+ Transient Detection in Smooth Muscle

Details methods for high-resolution Ca2+ imaging of smooth muscle within isolated organs, including: preparation of the tissue, image acquisition and data analysis.

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ ...

Heterotopic and Orthotopic Tracheal Transplantation in Mice used as Models to Study the Development of  Obliterative Airway Disease

Obliterative airway disease (OAD) is the major complication after lung transplantations that limits long term survival (1-7).
To study the pathophysiology, treatment and prevention of OAD, different animal models of tracheal transplantation in rodents have been developed (1-7). Here, we use two established models of trachea transplantation, the heterotopic and orthotopic model and demonstrate their advantages and limitations.
For the heterotopic model, the donor trachea is wrapped into the greater omentum of the recipient, whereas the donor trachea is anastomosed by end-to-end anastomosis in the orthotopic model.
In both models, the development of obliterative lesions histological similar to clinical OAD has been demonstrated (1-7).
This video shows how to perform both, the heterotopic as well as the orthotopic tracheal transplantation technique in mice, and compares the time course of OAD development in both models using histology.

This video shows and compares two experimental models to study the development of obliterative airway disease (OAD) in mice, the heterotopic and orthotopic tracheal transplantation model.

Obliterative airway disease (OAD) is the major complication after lung transplantations that limits long term survival (1-7).
To study the ...

Isolation of Human Umbilical Arterial Smooth Muscle Cells (HUASMC)

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded and their collection does not imply any added risk to the newborn or mother s health. Moreover no ethical concerns are raised. The UC is composed by one vein and two arteries from which both endothelial cells (ECs) 1 and smooth muscle cells (SMCs) 2, two of the main cellular components of blood vessels, can be isolated. In this project the SMCs were obtained after enzymatic treatment of the UC arteries accordingly the experimental procedure previously described by Jaffe et al 3. After cell isolation they were kept in t-flash with DMEM-F12 supplemented with 5% of fetal bovine serum and were cultured for several passages. Cells maintained their morphological and other phenotypic characteristics in the different generations. The aim of this study was to isolate smooth muscle cells in order to use them as models for future assays with constrictor drugs, isolate and structurally characterize L-type calcium channels, to study cellular and molecular aspects of the vascular function 4 and to use them in tissue engineering.

Isolation of Human Umbilical Arterial Smooth Muscle Cells HUASMC

The umbilical cords are used to isolate smooth muscle cells by different ways. In this work we used the enzymatic treatment to isolated smooth muscle cells.

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded ...

Exploring Arterial Smooth Muscle Kv7 Potassium Channel Function using Patch Clamp Electrophysiology and Pressure Myography

Contraction or relaxation of smooth muscle cells within the walls of resistance arteries determines the artery diameter and thereby controls flow of blood through the vessel and contributes to systemic blood pressure. The contraction process is regulated primarily by cytosolic calcium concentration ([Ca2+]cyt), which is in turn controlled by a variety of ion transporters and channels. Ion channels are common intermediates in signal transduction pathways activated by vasoactive hormones to effect vasoconstriction or vasodilation. And ion channels are often targeted by therapeutic agents either intentionally (e.g. calcium channel blockers used to induce vasodilation and lower blood pressure) or unintentionally (e.g. to induce unwanted cardiovascular side effects).
Kv7 (KCNQ) voltage-activated potassium channels have recently been implicated as important physiological and therapeutic targets for regulation of smooth muscle contraction. To elucidate the specific roles of Kv7 channels in both physiological signal transduction and in the actions of therapeutic agents, we need to study how their activity is modulated at the cellular level as well as evaluate their contribution in the context of the intact artery.
The rat mesenteric arteries provide a useful model system. The arteries can be easily dissected, cleaned of connective tissue, and used to prepare isolated arterial myocytes for patch clamp electrophysiology, or cannulated and pressurized for measurements of vasoconstrictor/vasodilator responses under relatively physiological conditions. Here we describe the methods used for both types of measurements and provide some examples of how the experimental design can be integrated to provide a clearer understanding of the roles of these ion channels in the regulation of vascular tone.

Measurements of Kv7 (KCNQ) potassium channel activity in isolated arterial myocytes (using patch clamp electrophysiological techniques) in parallel with measurements of constrictor/dilator responses (using pressure myography) can reveal important information about the roles of Kv7 channels in vascular smooth muscle physiology and pharmacology.

Contraction  or relaxation of smooth muscle cells within the walls of resistance arteries  determines the artery diameter and thereby controls flow of ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Measurement of Smooth Muscle Function in the Isolated Tissue Bath-applications to Pharmacology Research

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues. While this technique has been implemented for over 100 years, the versatility, simplicity and reproducibility of this assay helps it to remain an indispensable tool for pharmacologists and physiologists alike. Tissue bath systems are available in a wide array of shapes and sizes, allowing a scientist to evaluate samples as small as murine mesenteric arteries and as large as porcine ileum &#8211; if not larger. Central to the isolated tissue bath assay is the ability to measure concentration-dependent changes to isometric contraction, and how the efficacy and potency of contractile agonists can be manipulated by increasing concentrations of antagonists or inhibitors. Even though the general principles remain relatively similar, recent technological advances allow even more versatility to the tissue bath assay by incorporating computer-based data recording and analysis software. This video will demonstrate the function of the isolated tissue bath to measure the isometric contraction of an isolated smooth muscle (in this case rat thoracic aorta rings), and share the types of knowledge that can be created with this technique. Included are detailed descriptions of aortic tissue dissection and preparation, placement of aortic rings in the tissue bath and proper tissue equilibration prior to experimentation, tests of tissue viability, experimental design and implementation, and data quantitation. Aorta will be connected to isometric force transducers, the data from which will be captured using a commercially available analog-to-digital converter and bridge amplifier specifically designed for use in these experiments. The accompanying software to this system will be used to visualize the experiment and analyze captured data.

This protocol describes the measurement of isometric contraction in an isolated smooth muscle preparation, using an isolated tissue bath system and computer-based data acquisition.

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues.  ...

Isolation of Murine Coronary Vascular Smooth Muscle Cells

While the isolation and culture of vascular smooth muscle cells (VSMCs) from large vessels is well established, we sought to isolate and culture VSMCs from the coronary circulation. Hearts with intact aortic arches were removed and perfused via retrograde Langendorff with digestion solution containing 300 Units/ml of collagenase type II, 0.1 mg/ml soybean trypsin inhibitor and 1 M CaCl2. The perfusates were collected at 15 min intervals for 90 min, pelleted by centrifugation, resuspended in plating media, and plated on tissue culture dishes. VSMCs were characterized by presence of SM22&#945;, &#945;-SMA, and vimentin. One of the main advantages of using this technique is the ability to isolate VSMCs from the coronary circulation of mice. Although the small number of cells obtained can limit some of the applications for which the cells can be utilized, isolated coronary VSMCs can be used in a variety of well-established cell culture techniques and assays. Studies investigating VSMCs from genetically modified mice can provide further information about structure-function and signaling processes associated with vascular pathologies.

The purpose of this protocol is to demonstrate the isolation and culture techniques of murine primary vascular smooth muscle cells (VSMCs) from the coronary circulation. Once VSMCs have been isolated, they can be used for many standard culture techniques.

While the isolation and culture of vascular smooth muscle cells (VSMCs) from large vessels is well established, we sought to isolate and culture VSMCs ...

Measuring Proliferation of Vascular Smooth Muscle Cells Using Click Chemistry

The ability of a cell to proliferate is integral to the normal function of most cells, and dysregulation of proliferation is at the heart of many disease processes. For these reasons, measuring proliferation is a common tool used to assess cell function. Cell proliferation can be measured simply by counting; however, this is an indirect means of measuring proliferation. One common means of directly detecting cells preparing to divide is by incorporation of labeled nucleoside analogs. These include the radioactive nucleoside analog 3H-thymidine plus non-radioactive nucleoside analogs such as 5-bromo-2&#8217; deoxyuridine (BrdU) and 5-ethynyl-2&#8242;-deoxyuridine (EdU). Incorporation of EdU is detected by click chemistry, which has several advantages when compared to BrdU. In this report, we provide a protocol for measuring proliferation by the incorporation of EdU. This protocol includes options for various readouts, along with the advantages and disadvantages of each. We also discuss places where the protocol can be optimized or altered to meet the specific needs of the experiment planned. Finally, we touch on the ways that this basic protocol can be modified for measuring other cell metabolites.

Proliferation is a critical part of cellular function, and a common readout used to assess potential toxicity of new drugs. Measuring proliferation is, therefore, a frequently used assay in cell biology. Here we present a simple, versatile method of measuring proliferation that can be used in adherent and non-adherent cells.

The ability of a cell to proliferate is integral to the normal function of most cells, and dysregulation of proliferation is at the heart of many disease ...