We thank Dana Morgenstern for careful proofreading of the manuscript.

All methods and procedures described in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of Case Western Reserve University.
1. Preparation of Solutions, Instruments, and Coverslips
NOTE: 1x solutions can be used for up to a month.
<ol>
	<li>Make 10x Krebs-Henseleit buffer HEPES buffer without calcium (KHB-HB) by adding 68.96 g of NaCl, 3.57 g of KCl, 59.58 g of HEPES, 2.18 g of K2HPO4, 3.08 g of MgSO4 and 19.82 g of glucose to 800 mL of double distilled water in a 1,000 mL flask. After contents are fully dissolved, bring up to volume in a 1,000 mL volumetric flask. 
	NOTE: Traditional Krebs Henseleit solution uses sodium bicarbonate as a buffer and the solution in this protocol uses Krebs Henseleit solution with HEPES buffer. Solution is stable for 6 months if sterile filtered.</li>
	<li>Make 10x Tyrode&#39;s solution by adding 86.51 g of NaCl, 0.552 g of NaH2PO4, 2.03 g of MgCl2, 9.91 g of glucose, 4.03 g of KCl, 2.65 g of CaCl2, and 35.76 g of HEPES to 800 mL of double distilled water in a 1000 mL flask. After contents are fully dissolved, bring up to volume in a 1000 mL volumetric flask. 
	NOTE: Solution is stable for 6 months if sterile filtered.</li>
	<li>Make 1x KHB-HB by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Place flask in 37 &#176;C water bath. Once the solution has reached 37 &#176;C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1000 mL volumetric flask. Sterile filter the solution using a vacuum filtration system.</li>
	<li>Make 1x Tyrode&#39;s solution by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Place flask in 37 &#176;C water bath. Once the solution has reached 37 &#176;C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1,000 mL volumetric flask. Sterile filter using a vacuum filtration system.</li>
	<li>Make 1x modified Tyrode&#39;s solution by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Dissolve 3.07 g of L-glutathione reduced in flask. Place flask in 37 &#176;C water bath. Once the solution has reached 37 &#176;C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1,000 mL volumetric flask. Sterile filter the solution using a vacuum filtration system.</li>
	<li>Make 100 mM blebbistatin stock solution by adding 855 &#181;L of dimethyl sulfoxide (DMSO) to 25 mg of powder. Aliquot out in 20 &#181;L increments and store in a -80 &#176;C freezer for up to six months.</li>
	<li>Make stopping buffer by adding 2 g of bovine serum albumin (BSA) and 1 vial of aliquoted blebbistatin stock to 100 mL of 1x KHB-HB and sterile filter the solution using a vacuum filtration system.</li>
	<li>Make plating buffer by adding 5 mL of fetal bovine serum and 1 vial of the aliquoted blebbistatin stock to 95 mL of M199 HEPES. Sterile filter the solution using a vacuum filtration system.</li>
	<li>Make myocyte culture buffer by adding one 1 vial of the aliquoted blebbistatin stock and 4 mls penicillin-streptomycin to 396 mL of M199 (25 mM HEPES). Sterile filter the solution using a vacuum filtration system.</li>
	<li>Autoclave 2 pairs of Dumont tweezers, 2 pairs of Iris curved scissors, 2 hemostats, one pair of plastic surgery forceps, 6 black braided silk 4-0 sutures arranged to be used as a surgical double-throw knot, and four 100 mL beakers.</li>
	<li>Sterilize 22 x 22 mm2 glass coverslips. First, place a single coverslip in each well of a six well plate. Afterwards, with the lid removed, turn on the UV lamp of the biosafety cabinet and expose the coverslips to UV light for 1 h.</li>
	<li>Make working laminin stock solution by first thawing the bottle on ice. Add contents of one bottle to enough cold sterile phosphate-buffered saline (PBS) to reach a final concentration of 0.04 mg/mL. Aliquot out 1.3 &#181;L into autoclaved 1.5 mL centrifuge tubes. Store at -80 &#176;C. 
	NOTE: Each tube has enough laminin for a single six well plate. Avoid multiple freeze thaw cycles.</li>
	<li>Coat sterilized coverslip by first thawing the working laminin solution on ice. Using a P1000 pipette, aspirate 200 &#181;L of laminin. Gently drag the pipette tip along one edge of the coverslip to allow capillary action to pull out a minuscule amount of laminin to facilitate coverslip attachment to the six well plate.</li>
	<li>Then, expel the remaining laminin in the center of the coverslip. In a circular motion, spread the laminin droplet across the coverslip. Place in a 37 &#176;C incubator at least 1 h and up to 24 h before the isolation.</li>
</ol>
2. Preparation of the Langendorff Apparatus
NOTE: The individual components of the Langendorff apparatus used in this protocol are listed in Table of Materials.
<ol>
	<li>Turn on the circulating water bath. Set temperature so that perfusate has a temperature of 37 &#176;C. 
	NOTE: With the solution reservoirs set to a height of 60 cm, the circulating waterbirth needs to be set to 41 &#176;C to have the perfusate be 37 &#176;C. Unlike previously reported protocols, the height of the reservoir does not need to be changed.</li>
	<li>Rinse the Langendorff apparatus with 70% ethanol followed by two rinses with autoclaved double distilled water. After rinsing, fill reservoir with KHB-HB and oxygenate with 100% oxygen.</li>
	<li>Prime the system by allowing oxygenated KHB-HB to first flow into a 100 mL beaker. Once 50 mL of solution has flowed into the beaker, switch the 3-way stop-cock position to stop flow from the KHB-HB reservoir. Pour 50 mL of oxygenated KHB-HB from the beaker into the collagenase reservoir.</li>
	<li>Let the KHB-HB drain from the digestion reservoir until 5 mL remains in collagenase reservoir. While priming collagenase reservoir, switch the 3-way stop cock repeatedly between reservoirs to allow the lines to degas. After the system is primed, remember to use the degassing trap located on top of the heating coil to allow any remaining air to exit the system.</li>
	<li>Make the collagenase solution. For rats combine 100 mg of type II collagenase, 100 mL of oxygenated KHB-HB, and 2 vials of the blebbistain stock. For mice, combine 100 mg of type II collagenase, 40 mL of oxygenated KHB-HB, and 2 vials of the blebbistain stock. Once mixed, the solution should be stable for 1 h. 
	NOTE: Myocyte viability can vary between type II collagenase lots. Take advantage of a collagenase sampling program to test a lot before bulk ordering.</li>
</ol>
3. Myocyte Isolation
<ol>
	<li>Inject the animal with 1,000 U of heparin. Wait 5 min. 
	NOTE: Mice and rats of any age can be used. However, in general the older or more diseased the animal, the lower the myocyte yield.</li>
	<li>Sacrifice the animal by first anesthetizing it with isoflurane using the open-drop method (1 cc of Isoflurane per 500 cc volume) before euthanizing the animal with a pentobarbital mixture (150 mg/kg intraperitoneal).</li>
	<li>Rapidly excise the heart by first grabbing the fur above the xiphoid process. With the iris scissors, make a small incision immediately below the xiphoid process and pull the fur upwards toward the head exposing the skin.</li>
	<li>Grab the xiphoid process and cut the diaphragm exposing the thoracic cavity. Make a trap door incision, pull the sternum back using a hemostat, and use the curved forceps to excise the heart above the ascending aorta and place in cold KHB-HB.</li>
	<li>Cannulate the heart using a stereo microscope and number 5 forceps. Make sure the heart is submerged and the cannula was primed before heart excision to prevent emboli. Confirm proper positioning of the cannula by visualizing the tip of the cannula approximately 1 mm above aortic insertion into the ventricle. 
	NOTE: The faster the cannulation time, the better the myocyte yield.</li>
	<li>Start the flow of KHB-HB by rotating the stopcock on the Langendorff. Connect the cannula to the Langendorff. Perfuse the heart for 5 min. 
	NOTE: Since perfusion is supplied by a gravity-based system, flow through the heart will be a function of coronary artery compliance.</li>
	<li>Switch perfusion from the KHB-HB reservoir to the digestion buffer reservoir. Once the digestion buffer reaches the heart, set a timer (5 min for mouse or 15 min for rat). Make sure to collect the perfusate in a sterile 100 mL beaker. Refill the digestion buffer reservoir as needed with the perfusate until the digestion time has expired.</li>
	<li>After digestion, separate the chambers of the heart with forceps and the iris scissors in a sterile 100 mL beaker. Place each chamber into a separate well of a six well plate. Pour 5 mL of collagenase solution into each well.</li>
	<li>Immediately start mincing the heart tissue using scissors. Tissue chunks should be approximately 1 mm3. Using sterile transfer pipettes, gently triturate the minced heart tissue. The solution should turn cloudy.</li>
	<li>Once the tissue chunks become white and feathery, examine the cells using an inverted microscope. If the number of viable cells is greater than 80%, proceed to strain the cells into a 50 mL conical tube using a 100 &#181;m cell strainer. Use a different tube and strainer for each chamber of the heart.
	<ol>
		<li>If the number of viable cells is less than 80%, check the time it took to cannulate. If the cannulation time is over 5 min, try another heart. If not, assay new collagenase lots through the collagenase sampling program.</li>
	</ol>
	</li>
	<li>Pellet the cells by centrifuging at 215 x g for 2 min. The pellet should be compact and not loose. If the pellet is loose, the preparation contains many dead cells. In a tissue culture hood, resuspend the pellet in 10 mL of stopping buffer.</li>
	<li>Pellet the cells by centrifuging at 215 x g for 2 min. The pellet should be compact and not loose. If the pellet is loose, the preparation contains many dead cells.</li>
	<li>Resuspend the cells in 5 mL of plating buffer. Perform a cell count. Adjust the milliliters of plating buffer to reach a final myocyte concentration of 2 x 104 cells per mL.</li>
	<li>Remove the laminin-coated coverslips from the incubator. Aspirate the laminin droplet.</li>
	<li>Plate 200 &#181;L of myocyte suspension on each coverslip. Place in a 37 &#176;C incubator (21% O2, 5% CO2) for 2 h to allow attachment. After 2 h, aspirate the unattached cells, add 2 mL of culture media, and culture for up to 4 days.</li>
</ol>
4. Fura-2 Dye Loading
<ol>
	<li>Make a 2 mM fura-2 acetoxymethyl ester (fura-2 AM) stock solution by adding 25 &#181;L of DMSO to 50 &#181;g of fura-2 AM powder (1 vial). Aliquot out into 6 &#181;L aliquots. Take 1 aliquot of fura-2 AM and add to 6 mL of plating medium. Vortex to mix.</li>
	<li>Remove 1 six well plate of myocytes from the incubator. Aspirate media. Add 1 mL of fura-2 media mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.</li>
	<li>Aspirate fura-2 media mixture and add 1 mL of Tyrode&#39;s solution to each well. Cover with foil. Wait 20 min at room temperature to allow for dye washout before imaging.</li>
</ol>
5. Fluo-4 Dye Loading
<ol>
	<li>Make a 1.82 mM fluo-4 acetoxymethyl ester (fluo-4 AM) stock solution by adding 25 &#181;L of DMSO to 50 &#181;g of fluo-4 AM powder (1 vial). Aliquot out into 8.333 &#181;L aliquots. Take 1 aliquot of fluo-4 AM stock and add to 6 mL of plating medium. Vortex to mix.</li>
	<li>Remove 1 six well plate of myocytes from the incubator. Aspirate media. Add 1 mL of fluo-4 AM media mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.</li>
	<li>Aspirate fluo-4 AM media mixture and add 1 mL of Tyrode&#39;s solution to each well. Cover with foil. Wait 20 min at room temperature to allow for dye washout before imaging.</li>
</ol>
6. Membrane Potential Dye Loading
<ol>
	<li>Remove component A and component B from the membrane potential kit. In a 15 mL conical tube, combine 50 &#181;L of component B and 5 &#181;L of component A. Vortex to mix. Add 10 mL of plating media to the 15 mL conical tube containing the voltage dye mixture. Vortex to mix.</li>
	<li>Remove 1 six well plate of myocytes from the incubator. Aspirate the media. Add 800 &#181;L of the membrane potential dye mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.</li>
	<li>Aspirate dye media mixture and add 1 mL of modified-Tyrode&#39;s solution to each well. Cover with foil.</li>
</ol>
7. Photometry and Charge Coupled Device Recordings
<ol>
	<li>Turn on the equipment in the following order: microscope, arc lamp, hyperswitch, fluorescence interface system, Myocam power supply, field stimulator, and computer.</li>
	<li>Make sure the excitation/emission filter sets are appropriate for the imaging dye. 
	NOTE: Fura-2 is excited at 340 nm and 380 nm of light. It emits at 510 nm of light. Fluo-4 and the voltage membrane dye are excited at 485 nm of light and emit at 520 nm of light.</li>
	<li>Prime the system by turning on the vacuum, fully opening the hose clamp, and gently plunging each 60 mL syringe being used in the manifold. For calcium recordings use Tyrode&#39;s solution. For voltage recordings use modified Tyrode&#39;s solution.</li>
	<li>Turn heater on and set flow by adjusting the roller clamp on the perfusion tubing. Make recordings at 36 &#177; 1 &#176;C.</li>
	<li>Open the acquisition software. Make sure the parameters are set for the correct imaging dye.</li>
	<li>In the dark, remove the foil from the six well plate and place a coverslip in the pacing chamber. Make sure the stimulator is off during this step. Focus on the myocytes using the 10x objective.</li>
	<li>Once in focus, start pacing by field stimulating at 1 Hz, 0.2 V. Gradually increase the voltage until 1:1 pacing is obtained. Then increase the voltage until 1.5x the threshold is reached. 
	NOTE: Because excitation-contraction coupling is temperature dependent, make sure the cells have been perfused for 15 min before recording. This allows for myocytes to recover from the shock of going from room temperature back to 37 &#176;C as well as loosely attached cells to float away.</li>
	<li>Switch from the 10x objective to the 40x objective. Focus in on a cell that is following a 1:1 pacing. Adjust the plastic shades so only one cell is in the field of view.</li>
	<li>Using the software, place the area of interest box on well-defined sarcomeres. Start the acquisition software to initiate the excitation light. Using the neutral density filters, adjust the intensity setting accordingly to obtain an adequate SNR.</li>
</ol>

<ol>
	<li>Hagen, B. M., Boyman, L., Kao, J. P., Lederer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparative+assessment+of+fluo+Ca2++indicators+in+rat+ventricular+myocytes.">A comparative assessment of fluo Ca2+ indicators in rat ventricular myocytes.</a> Cell Calcium. 52 (2), 170-181 (2012).</li><li>Schaffer, P., Ahammer, H., Muller, W., Koidl, B., Windisch, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Di-4-ANEPPS+causes+photodynamic+damage+to+isolated+cardiomyocytes.">Di-4-ANEPPS causes photodynamic damage to isolated cardiomyocytes.</a> Pfl&#252;gers Archiv: European Journal of Physiology. 426 (6), 548-551 (1994).</li><li>Hardy, M. E., Lawrence, C. L., Standen, N. B., Rodrigo, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Can+optical+recordings+of+membrane+potential+be+used+to+screen+for+drug-induced+action+potential+prolongation+in+single+cardiac+myocytes.">Can optical recordings of membrane potential be used to screen for drug-induced action potential prolongation in single cardiac myocytes.</a> Journal of Pharmacological and Toxicological Methods. 54 (2), 173-182 (2006).</li><li>Tian, Q., 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=Optical+action+potential+screening+on+adult+ventricular+myocytes+as+an+alternative+QT-screen.">Optical action potential screening on adult ventricular myocytes as an alternative QT-screen.</a> Cellular Physiology and Biochemistry. 27 (3-4), 281-290 (2011).</li><li>Warren, M., 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=High-precision+recording+of+the+action+potential+in+isolated+cardiomyocytes+using+the+near-infrared+fluorescent+dye+di-4-ANBDQBS.">High-precision recording of the action potential in isolated cardiomyocytes using the near-infrared fluorescent dye di-4-ANBDQBS.</a> The American Journal of Physiology-Heart and Circulatory Physiology. 299 (4), 1271-1281 (2010).</li><li>FluoVolt&#8482; Membrane Potential Kit. ThermoFisher Scientific Available from: <a target="_blank" href="https://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0009668_FluoVolt_Membrane_Potential_Kit_UG.pdf">https://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0009668_FluoVolt_Membrane_Potential_Kit_UG.pdf</a> (2018)</li><li>Fluorescence SpectraViewer. ThermoFisher Scientific Available from: <a target="_blank" href="https://www.thermofisher.com/us/en/home/life-science/cell-analysis/labeling-chemistry/fluorescence-spectraviewer.html">https://www.thermofisher.com/us/en/home/life-science/cell-analysis/labeling-chemistry/fluorescence-spectraviewer.html</a> (2019)</li><li>Paredes, R. M., Etzler, J. C., Watts, L. T., Zheng, W., Lechleiter, J. 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The authors have nothing to disclose.

Being able to isolate cardiac myocytes is a powerful method that can be used to understand cardiac physiology, pathology, and toxicology. In the above protocol, we described a method that utilizes a constant gravity pressure Langendorff apparatus to obtain single cardiac myocytes. Afterwards, using the fluorescence photometry system, we describe how to simultaneously acquire either calcium and shortening or voltage and shortening traces.
Because of the different kinetics between calcium dyes, care must be taken on which dye to select. For this protocol, both the fura-2 and fluo-4 used were engineered with AM esters necessitating a wash step to allow for intracellular esterases time to cleave the AM group and trap the dye in the cell. While both fura-2 and fluo-4 are considered high affinity calcium dyes, the Kd for fura-2 is 145 nM compared to the 345 nM for fluo-49. Further, fura-2 is ratiometric. Because of this, it can be used to quantify intracellular calcium levels9,12. Fluo-4 on the other hand is a single wave calcium probe. The advantage of using fluo-4 is it produces a brighter fluorescence signal. Regardless of which calcium dye is used, compared to the calcium dye, membrane voltage probes have a lower SNR.
As shown in Figure 4 and Figure 5, voltage traces compared to calcium traces are smaller in amplitude. Using the software&#39;s digital trace filtering, it is possible to increase the SNR and quantify the data (Figure 4 and Figure 7). Once quantified, both calcium transients and optical APDs demonstrate restitution, shortening their duration at faster pacing frequencies (Figure 2, Figure 3, Figure 6, and Figure 7). Shorter APDs during faster pacing cycles are necessary to allow enough time for ventricular filling during diastole. Alterations in this phenomenon is thought to be indicative of an increase in the risk of arrythmias13,14,15,16. While alterations in APD can be caused by disease, they can also be caused by chemicals. As shown in Figure 7, when the predominant murine repolarizing potassium current, Ito, is blocked, the optical APD becomes longer.
Still, as reported previously with voltage sensitive dyes, light intensity and duration can alter the APD2,5,17. This is believed to be the result of the generation of reactive oxidative species (ROS)5. Previously, it has been shown that the addition of antioxidants to the recording solution can prevent voltage sensitive dye cytotoxicity5. As a result, we added the antioxidant L-glutathione (10 mM), to Tyrode&#39;s solution. Shown in Figure 8 is the last 11 s of a 20 s recording obtained at 1 Hz pacing. As indicated by the red arrows, alterations in the APD did not occur until 15 s into the recording; therefore, while the modified Tyrode&#39;s solution did not prevent phototoxicity it delayed it significantly. Using modified Tyrode&#39;s solution, using a low light intensity setting and keeping the duration of the recording to under 5 s, it is possible to avoid any dye induced alterations in APD. This is important because without taking care to avoid phototoxicity, the data could be misinterpreted as causing early or delayed after depolarizations. In addition to limiting the exposure to blue light, there are additional precautions that can be taken to prevent misinterpretation of the data.
The first is to only record from cells that follow one to one pacing and have a resting sarcomere length greater than or equal to 1.75 &#181;m. The 1.75 &#181;m cutoff is taken from the observation by Gordon et al.18 that tension rapidly declines once the sarcomere length is below this amount. Nonetheless, certain pathologies may result in significant alterations in resting sarcomere length. To be sure that the phenotype is real and not an artifact of the isolation, the following trouble shooting approaches should be taken.
If myocytes are consistently not following 1:1 pacing, have sarcomere lengths below 1.75 &#181;m, heavy membrane blebbing, or do not survive the isolation, the first thing to check is the time it took to cannulate the heart. The longer the cannulation time, the lower the yield will be. If a long cannulation time is required, viability can be improved by placing the heart in a cardioplegic solution19. Nonetheless, because the collagenase is an enzyme, the activity and specificity of a specific lot change over time. If the overall yields progressively become worse despite good cannulation times, new lots should be assayed. While our protocol was optimized for 5 s recordings, if longer voltage traces are needed, additional neutral density filters will need to be purchased. The system described in the protocol comes with neutral density filters that reduce the transmitted light by 37%, 50%, 75%, 90%, and 95%.
In summary, we described a methodology that allowed for the isolation of adult murine ventricular myocytes that were used for calcium, voltage, and sarcomere shortening measurements.

In order to study the heart during healthy and pathological states, it is often useful to examine the phenotype at the single cell level. While scientific advances have permitted the robust measurement of single cell calcium dynamics, single cell optical voltage measurements have remained scarce1. Arguably, this is because of the low signal to noise ratio (SNR), phototoxicity, and photobleaching of traditional potentiometric dyes2,3. Nonetheless, isolated myocyte optical action potentials have been obtained2,3,4. Further, with advances in the chemistry and the physics of voltage sensitive dyes, the SNR has improved5. Newer membrane potential probes (Table of Materials) respond to changes in membrane potential in sub-milliseconds and have a fluorogenic response range of approximately 25% per 100 mV. Further, the excitation/emission of the membrane potential kit (e.g., FluoVolt; Table of Materials) used in this protocol works with standard fluorescein isothiocyanate (FITC) or green fluorescent protein (GFP) settings6.
The FITC and GFP excitation/emission spectra overlap with the fluo-4 calcium bound spectra7. Simultaneous acquisition of fluorescence photometry with digital cell geometry measurements traditionally has been used for the simultaneous acquisition of calcium and cellular shortening measurements8. This protocol describes in detail how to isolate murine myocytes and how to record calcium or voltage signals using standard FITC settings. Additionally, it describes how a simple switch in excitation/emission filters on the imaging workstation can be used to obtain calcium and shortening measurements using the ratio metric calcium dye fura-2. Compared to fluo-4, fura-2 has a higher affinity for calcium and is relatively resistant to photobleaching9. Consequently, using a single workstation this protocol allows for a thorough examination of singly myocyte excitation-contraction coupling.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.25 Liter Water Jacketed Reservoir</td>
			<td>Radnoti, LLC</td>
			<td>120142-025</td>
			<td></td>
		</tr>
		<tr>
			<td>1 liter volumetric flask</td>
			<td>Fisher Scientific</td>
			<td>10-205F</td>
			<td></td>
		</tr>
		<tr>
			<td>100 ml beaker</td>
			<td>Fisher Scientific</td>
			<td>FB-100-100</td>
			<td></td>
		</tr>
		<tr>
			<td>100 ml graduated cylinder</td>
			<td>Fisher Scientific</td>
			<td>08 562 5C</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 ml flask</td>
			<td>Fisher Scientific</td>
			<td>FB-500-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>2-Bar Lab Stand with Stabilizer Bar and 24&#34; Stainless Steel Rods</td>
			<td>Radnoti, LLC</td>
			<td>159951-2</td>
			<td></td>
		</tr>
		<tr>
			<td>4-Aminopyridine</td>
			<td>Sigma-Aldrich</td>
			<td>275875</td>
			<td></td>
		</tr>
		<tr>
			<td>40X Oly UApo/340 Non-Immersion Objective (NA 0.9, WD 0.2mm)</td>
			<td>IonOptix</td>
			<td>MSCP1-40 (b)</td>
			<td></td>
		</tr>
		<tr>
			<td>60-mL syringe, BD Luer-Lok tip</td>
			<td>BD</td>
			<td>309650</td>
			<td></td>
		</tr>
		<tr>
			<td>Aortic Metal Cannulae</td>
			<td>Harvard Apparatus</td>
			<td>73-0112</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Fisher Scientific</td>
			<td>BP9703-100</td>
			<td></td>
		</tr>
		<tr>
			<td>C-6 Standard Heating Circulator</td>
			<td>Chemyx</td>
			<td>A30006</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Fisher Scientific</td>
			<td>BP510500</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell framing adapter</td>
			<td>IonOptix</td>
			<td>CFA300</td>
			<td></td>
		</tr>
		<tr>
			<td>CellPr0 Vacuum Filtration System, 1 liter, 0.22&#181;m,Cs/12</td>
			<td>Labratory Product Sales, Inc</td>
			<td>V100022</td>
			<td></td>
		</tr>
		<tr>
			<td>CellPro Vacuum Filtration System, 250mL, 0.22&#181;m,Cs/12</td>
			<td>Labratory Product Sales, Inc</td>
			<td>V25022</td>
			<td></td>
		</tr>
		<tr>
			<td>CellPro Vacuum Filtration System, 500mL, 0.22&#181;m,12/CS</td>
			<td>Labratory Product Sales, Inc</td>
			<td>V50022</td>
			<td></td>
		</tr>
		<tr>
			<td>CMC (mTCII) Temp Control w/ inline flow heater</td>
			<td>IonOptix</td>
			<td>TEMPC2</td>
			<td></td>
		</tr>
		<tr>
			<td>Cole-Parmer Large-bore 3-way, male-lock, stopcocks</td>
			<td>Cole-Parmer</td>
			<td>EW-30600-23</td>
			<td></td>
		</tr>
		<tr>
			<td>Cole-Parmer Luer fittings, Large-bore stopcocks, male lock, 4-way</td>
			<td>Cole-Parmer</td>
			<td>EW-30600-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Cole-Parmer Stopcocks with Luer Connections; 1-way; male slip</td>
			<td>Cole-Parmer</td>
			<td>EW-30600-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase Type II</td>
			<td>Worthington</td>
			<td>LS004177</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Sterile Cell Strainers</td>
			<td>Fisher Scientific</td>
			<td>07-201-432</td>
			<td></td>
		</tr>
		<tr>
			<td>Dell Optiplex 790 mini-tower, 4G RAM, 250G HD, Windows 7 Pro</td>
			<td>IonOptix</td>
			<td>CPUD7M</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Fisher Scientific</td>
			<td>50980367</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont Tweezers Style 5</td>
			<td>Amazon</td>
			<td>B00F70ZDEQ</td>
			<td></td>
		</tr>
		<tr>
			<td>FHD Rapid Change Stimulation Chamber</td>
			<td>IonOptix</td>
			<td>FHDRCC1</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluo-3/4 Optics Package</td>
			<td>IonOptix</td>
			<td>IonOP-Fluo</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence system interface &#8211; (w PCI-I/O card)</td>
			<td>IonOptix</td>
			<td>FSI700</td>
			<td></td>
		</tr>
		<tr>
			<td>Gibco Penicillin-Streptomycin (10,000 U/mL)</td>
			<td>Fisher Scientific</td>
			<td>15-140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Fisher Scientific</td>
			<td>D16-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemostat, Curved 5-1/2&#34;</td>
			<td>Amazon</td>
			<td>B00GGAAPD0</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Fisher Scientific</td>
			<td>BP310500</td>
			<td></td>
		</tr>
		<tr>
			<td>HyperSwitch dual excitation light source</td>
			<td>IonOptix</td>
			<td>HSW400</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Motic Fluorescence Microscope</td>
			<td>IonOptix</td>
			<td>MSCP1-40 (a)</td>
			<td></td>
		</tr>
		<tr>
			<td>IonWizard Core + Analysis</td>
			<td>IonOptix</td>
			<td>IONWIZ</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris Scissors, curved</td>
			<td>Amazon</td>
			<td>B018KRRMY6</td>
			<td></td>
		</tr>
		<tr>
			<td>K2HPO4</td>
			<td>Fisher Scientific</td>
			<td>P288-100</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher Scientific</td>
			<td>BP3661</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutathione reduced</td>
			<td>Sigma-Aldrich</td>
			<td>G4251</td>
			<td></td>
		</tr>
		<tr>
			<td>LOOK Silk Spool, Black Braided, 4-0, 100yds</td>
			<td>SouthernAnesthesiaSurgical Inc.</td>
			<td>SP116-EA</td>
			<td></td>
		</tr>
		<tr>
			<td>M199 Media</td>
			<td>Fisher Scientific</td>
			<td>12 340 030</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Fisher Scientific</td>
			<td>MP021914215</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>Fisher Scientific</td>
			<td>BP2131</td>
			<td></td>
		</tr>
		<tr>
			<td>MyoCam-S Digital CCD video system</td>
			<td>IonOptix</td>
			<td>MCS100</td>
			<td></td>
		</tr>
		<tr>
			<td>MyoPacer Field Stimulator</td>
			<td>IonOptix</td>
			<td>MYP100</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Fisher Scientific</td>
			<td>BP358212</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Fisher Scientific</td>
			<td>56-754-9250GM</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygenator Bubbler with Fluid Inlet for 0.25 Liter</td>
			<td>Radnoti, LLC</td>
			<td>140143-025</td>
			<td></td>
		</tr>
		<tr>
			<td>Photomultiplier sub-system</td>
			<td>IonOptix</td>
			<td>PMT400</td>
			<td></td>
		</tr>
		<tr>
			<td>PMT Acquisition add-on</td>
			<td>IonOptix</td>
			<td>PMTACQ</td>
			<td></td>
		</tr>
		<tr>
			<td>Radnoti Heating Coil 5 mL with Degasing Trap</td>
			<td>Radnoti, LLC</td>
			<td>158830</td>
			<td></td>
		</tr>
		<tr>
			<td>Ring Clamp 60 &#8211; 80mm Dia. for 250ml Reservoir</td>
			<td>Radnoti, LLC</td>
			<td>120141-025</td>
			<td></td>
		</tr>
		<tr>
			<td>Ring Clamp for Bubble Trap Compliance Chamber</td>
			<td>Radnoti, LLC</td>
			<td>120149RC</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF000010 5/32 in.9/32 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-214</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF000013 3/16 in.3/8 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-217</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF000016 1/4 in.5/16 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-219</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF000025 5/16 in.5/8 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-226</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF00003 1/16 in.3/16 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-209</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF00005 1/16 in.3/32 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-210</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain ACF00009 5/32 in.7/32 in.</td>
			<td>Fisher Scientific</td>
			<td>14-171-213</td>
			<td></td>
		</tr>
		<tr>
			<td>Sarcomere Length Recording add-on</td>
			<td>IonOptix</td>
			<td>SARACQ</td>
			<td></td>
		</tr>
		<tr>
			<td>T/C Adson Tissue Platic Surgery Forceps 4.75&#34;</td>
			<td>Amazon</td>
			<td>B00JDWRBGC</td>
			<td></td>
		</tr>
		<tr>
			<td>VETUS Anti-Static Curved Tip Tweezers</td>
			<td>Amazon</td>
			<td>B07QMZC94J</td>
			<td></td>
		</tr>
		<tr>
			<td>Vistek 3200 Motic Vibration Isolation Platform</td>
			<td>IonOptix</td>
			<td>ISO100</td>
			<td></td>
		</tr>
	</tbody>
</table>

optical imaging of isolated murine ventricular myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

Figure 1A shows the Langendorff apparatus. The oxygenator is in the KHB-HB reservoir. The collagenase solution is in the middle 60 mL syringe reservoir. The degassing line is connected to the empty 60 mL syringe reservoir. After a successful isolation, most of the cells should be rod shaped and striated. Under a 40x objective, most myocytes should have clear striations visible. Figure 1B,C shows examples of healthy rat myocytes. Once isolated, cells can be cultured up to 4 days while maintaining their morphology and electrical properties.
To measure excitation-contraction coupling, the cells are then placed in a heated pacing chamber. Because myocytes are sensitive to changes in temperature, it is important to allow the coverslip to equilibrate for 15 min in the chamber before recording. For fluorescence recordings, the excitation wavelength is generated by a 75 W xenon-arc bulb. Xenon-arc bulbs produce a light spectrum that mimics natural sunlight. The intensity of the light and the wavelength are controlled by neutral density/emission filers. The excitation light then passes through the objective to the myocyte. The emission wavelength is then collected by a photomultiplier tube. Using the system described here, both the excitation and emission filters need to be changed manually.
Shortening on the other hand is obtained by a charge coupled device sensor. Measuring in real time up to 1,000 times per second, the acquisition software performs an average of the lines within an area of interest to create a well resolved striation pattern. A fast Fourier transform (FFT) is then calculated. The peak within the power spectrum represents the average sarcomere spacing. Changes in the sarcomere spacing during pacing are then plotted and subsequently quantified.
Figure 2 shows calcium and shortening traces recorded from a C57/B6 mouse myocyte loaded with the calcium dye fura-2. The pacing protocol is a modification of pacing protocols described previously10,11. Healthy mouse myocytes should be able to be paced at their resting heart rate 10 Hz. Figure 3 is quantification of ensembled averaged data obtained from a C57/B6 mice and their transgenic (TG) littermates who had a point mutation introduced into a potassium channel. Notice there is no difference between the groups except for the relaxation time at 10 Hz pacing.
Unlike fura-2 which is a dual excitation dye, the voltage dye and fluo-4 are single wavelength excitation dyes whose excitation/emission work with standard FITC excitation and emission spectrum (494/506 nm). Therefore, recordings of calcium and sarcomere shortening or voltage and sarcomere shortening can be obtained using this filter set.
Figure 4A shows a voltage tracing recorded from a C57/B6 mouse myocyte paced at 10 Hz. Compared to calcium signals, single cell voltage tracings are smaller in amplitude and need post-processing to obtain a useable signal. Figure 4B shows an ensembled averaged action potential (AP) made from the APs in Figure 4A. Figure 4C,D shows an ensembled average AP after a low pass Butterworth or a Savitzky-Golay digital filter was applied. Care must be taken when filtering the signal as not to distort the real data. Notice the subtle differences in the shape of the APs in Figure 4B-D.
Figure 5 shows traces recorded from rat myocytes paced at 1 Hz. In addition to the voltage signal being lower than the calcium signal, the contraction kinetics are different as well. This is because calcium dyes buffer calcium while voltage dyes do not.
As with the calcium transient (Figure 3), myocytes demonstrated pacing dependent changes in their optical action potential duration (APD) as well (Figure 6). While the fura-2 traces were ensembled averaged before being quantified, the voltage traces were filtered with a Savitzky-Golay polynomial smoothing filter (width 5, order 2) before being ensembled averaged and quantified.
As quantified in Figure 6 and Figure 7, in addition to demonstrating pacing induced changes in APD, they also demonstrated drug induced prolongation of the AP. At 4 Hz pacing, concentration dependent blockade of the transient outward current (Ito) with 4-aminopyridine resulted in prolongation of the APD.
Finally, care must be taken to avoid cytotoxicity. Figure 8 is the last 11 s of a 20 s recording. Indicated by the red arrows in Figure 8, prolonged exposure of myocytes to blue light leads to triggered activity.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60196/60196fig1a.jpg" /> 
Figure 1: Constant pressure Langendorff apparatus. (A) The Langendorff Apparatus with each component labeled in white lettering. (B) Isolated Sprague-Dawley rat myocytes viewed through a 10x objective. (C) Isolated rat myocytes viewed through a 40x objective. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60196/60196fig2a.jpg" /> 
Figure 2: Representative calcium and sarcomere shortening traces recorded from C57/B6 myoyctes using fura-2. Calcium and sarcomere shortening traces recorded at 1, 2, 4, 10, 0.5 and 0.75 Hz. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60196/60196fig3a.jpg" /> 
Figure 3: Quantification of sarcomere shortening, peak calcium, relaxation time, and reuptake time recorded from a C57/B6 wild type (WT) and transgenic (TG) mice. (A) Sarcomere shortening. (B) Peak calcium. (C) Relaxation time defined as 90% return to baseline of the shortening trace. (D) Reuptake time defined as 90% return to baseline of the calcium trace. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60196/60196fig4a.jpg" /> 
Figure 4: Optical action potential recorded from a C57/B6 mouse myocyte paced at 10 Hz. (A) 1 second unfiltered trace. (B) Ensembled averaged optical action potential. (C) Ensembled averaged optical action potential after a lowpass Butterworth filter was applied. (D) Ensembled averaged optical action potential after a Savitzky-Golay polynomial smoothing filter was applied. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60196/60196fig5a.jpg" /> 
Figure 5: Representative calcium, voltage, and sarcomere shortening traces recorded from Sprague-Dawley rat myocytes paced at 1 Hz. (A) Calcium and sarcomere shortening traces recorded at 1 Hz pacing using fluo-4. (B) Voltage and sarcomere shortening traces recorded at 1 Hz pacing using the voltage dye. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60196/60196fig6a.jpg" /> 
Figure 6: Optical action potentials recorded from Sprague-Dawley rat myocytes paced at 1, 2, and 4 Hz pacing. (A) Filtered trace recorded at 1 Hz pacing. (B) Filtered trace recorded at 2 Hz pacing. (C) Filtered trace recorded at 4 Hz pacing. (D) Action potential duration 10, measured as 10% return to baseline. (E) Action potential duration 50, measured as 50% return to baseline. (F) Action potential duration 90, measured as 90% return to baseline. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60196/60196fig7a.jpg" /> 
Figure 7: The Effects of 4-aminopyridine on Sprague-Dawley rat optical action potentials recorded at 4 Hz pacing. (A) Ensembled averaged trace recorded at 4 Hz pacing with no 4-Aminopyridine in the solution. (B) Ensembled averaged trace recorded at 4 Hz pacing with 1 &#181;M 4-Aminopyridine in the solution. (C) Ensembled averaged trace recorded at 4 Hz pacing with 10 &#181;M 4-Aminopyridine in the solution. (D) Action potential duration 10, measured as 10% return to baseline. (E) Action potential duration 50, measured as 50% return to baseline. (F) Action potential duration 90, measured as 90% return to baseline. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/60196/60196fig8a.jpg" /> 
Figure 8: Voltage dye induced phototoxicity in Sprague-Dawley rat myocytes after 20 s of continuous light exposure. Red arrows indicate cytotoxic events. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Optical Imaging of Isolated Murine Ventricular Myocytes at JoVE.com

Optical Imaging of Isolated Murine Ventricular Myocytes

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

optical-imaging-of-isolated-murine-ventricular-myocytes

Research

JoVE Journal

Bioengineering

6.1K Views.  University of Science and Technology of China, Hefei, Anhui, China. We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

Video: Optical Imaging of Isolated Murine Ventricular Myocytes

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

In vitro Synthesis of Native, Fibrous Long Spacing and Segmental Long Spacing Collagen

Collagen fibrils are present in the extracellular matrix of animal tissue to provide structural scaffolding and mechanical strength. These native collagen fibrils have a characteristic banding periodicity of ~67 nm and are formed in vivo through the hierarchical assembly of Type I collagen monomers, which are 300 nm in length and 1.4 nm in diameter. In vitro, by varying the conditions to which the monomer building blocks are exposed, unique structures ranging in length scales up to 50 microns can be constructed, including not only native type fibrils, but also fibrous long spacing and segmental long spacing collagen. Herein, we present procedures for forming the three different collagen structures from a common commercially available collagen monomer. Using the protocols that we and others have published in the past to make these three types typically lead to mixtures of structures. In particular, unbanded fibrils were commonly found when making native collagen, and native fibrils were often present when making fibrous long spacing collagen. These new procedures have the advantage of producing the desired collagen fibril type almost exclusively. The formation of the desired structures is verified by imaging using an atomic force microscope.

In vitro Synthesis of Native, Fibrous Long Spacing and Segmental Long Spacing Collagen

Simple and reproducible procedures are described for making three structurally distinct collagen assemblies from a common commercially available Type I collagen monomer. Native type, fibrous long spacing or segmental long spacing collagen can be constructed by varying the conditions to which the 300 nm long and 1.4 nm diameter monomer building block is exposed.

Collagen fibrils are present in the  extracellular matrix of animal tissue to provide structural scaffolding and  mechanical strength. These native ...

Wideband Optical Detector of Ultrasound for Medical Imaging Applications

Optical sensors of ultrasound are a promising alternative to piezoelectric techniques, as has been recently demonstrated in the field of optoacoustic imaging. In medical applications, one of the major limitations of optical sensing technology is its susceptibility to environmental conditions, e.g. changes in pressure and temperature, which may saturate the detection. Additionally, the clinical environment often imposes stringent limits on the size and robustness of the sensor. In this work, the combination of pulse interferometry and fiber-based optical sensing is demonstrated for ultrasound detection. Pulse interferometry enables robust performance of the readout system in the presence of rapid variations in the environmental conditions, whereas the use of all-fiber technology leads to a mechanically flexible sensing element compatible with highly demanding medical applications such as intravascular imaging. In order to achieve a short sensor length, a pi-phase-shifted fiber Bragg grating is used, which acts as a resonator trapping light over an effective length of 350 &#181;m. To enable high bandwidth, the sensor is used for sideway detection of ultrasound, which is highly beneficial in circumferential imaging geometries such as intravascular imaging. An optoacoustic imaging setup is used to determine the response of the sensor for acoustic point sources at different positions.

Optical detection of ultrasound is impractical in many imaging scenarios because it often requires stable environmental conditions. We demonstrate an optical technique for ultrasound sensing in volatile environments with miniaturization and sensitivity levels appropriate for optoacoustic imaging in restrictive scenarios, e.g. intravascular applications.

Optical sensors of ultrasound are a promising alternative to piezoelectric techniques, as has been recently demonstrated in the field of optoacoustic ...

Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. While the outer surface membrane and associated TATS membrane components appear to be continuous, there are substantial differences in lipid and protein content. In ventricular myocytes (VMs), certain TATS components are highly abundant contributing to rectilinear tubule networks and regular branching 3D architectures. It is thought that peripheral TATS components propagate action potentials from the cell surface to thousands of remote intracellular sarcoendoplasmic reticulum (SER) membrane contact domains, thereby activating intracellular Ca2+ release units (CRUs). In contrast to VMs, the organization and functional role of TATS membranes in atrial myocytes (AMs) is significantly different and much less understood. Taken together, quantitative structural characterization of TATS membrane networks in healthy and diseased myocytes is an essential prerequisite towards better understanding of functional plasticity and pathophysiological reorganization. Here, we present a strategic combination of protocols for direct quantitative analysis of TATS membrane networks in living VMs and AMs. For this, we accompany primary cell isolations of mouse VMs and/or AMs with critical quality control steps and direct membrane staining protocols for fluorescence imaging of TATS membranes. Using an optimized workflow for confocal or superresolution TATS image processing, binarized and skeletonized data are generated for quantitative analysis of the TATS network and its components. Unlike previously published indirect regional aggregate image analysis strategies, our protocols enable direct characterization of specific components and derive complex physiological properties of TATS membrane networks in living myocytes with high throughput and open access software tools. In summary, the combined protocol strategy can be readily applied for quantitative TATS network studies during physiological myocyte adaptation or disease changes, comparison of different cardiac or skeletal muscle cell types, phenotyping of transgenic models, and pharmacological or therapeutic interventions.

In cardiac myocytes, tubular membrane structures form intracellular networks. We describe optimized protocols for i) isolation of myocytes from mouse heart including quality control, ii) live cell staining for state-of-the-art fluorescence microscopy, and iii) direct image analysis to quantify the component complexity and the plasticity of intracellular membrane networks.

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Computed Tomography and Optical Imaging of Osteogenesis-angiogenesis Coupling to Assess Integration of Cranial Bone Autografts and Allografts

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that implantation of a bone autograft would induce greater bone regeneration by abundant blood vessel formation. To investigate the effect of the graft on neovascularization at the defect site, we developed a micro&#8211;computed tomography (&#181;CT) approach to characterize newly forming blood vessels, which involves systemic perfusion of the animal with a polymerizing contrast agent. This method enables detailed vascular analysis of an organ in its entirety. Additionally, blood perfusion was assessed using fluorescence imaging (FLI) of a blood-borne fluorescent agent. Bone formation was quantified by FLI using a hydroxyapatite-targeted probe and &#181;CT analysis. Stem cell recruitment was monitored by bioluminescence imaging (BLI) of transgenic mice that express luciferase under the control of the osteocalcin promoter. Here we describe and demonstrate preparation of the allograft, calvarial defect surgery, &#181;CT scanning protocols for the neovascularization study and bone formation analysis (including the in vivo perfusion of contrast agent), and the protocol for data analysis.
The 3D high-resolution analysis of vasculature demonstrated significantly greater angiogenesis in animals with implanted autografts, especially with respect to arteriole formation. Accordingly, blood perfusion was significantly higher in the autograft group by the 7th day after surgery. We observed superior bone mineralization and measured greater bone formation in animals that received autografts. Autograft implantation induced resident stem cell recruitment to the graft-host bone suture, where the cells differentiated into bone-forming cells between the 7th and 10th postoperative day. This finding means that enhanced bone formation may be attributed to the augmented vascular feeding that characterizes autograft implantation. The methods depicted may serve as an optimal tool to study bone regeneration in terms of tightly bounded bone formation and neovascularization.

Implantation of autologous and allogeneic bone grafts constitute accepted approaches to treat major craniofacial bone loss. Yet the effect of graft composition on the interplay among neovascularization, cell differentiation and bone formation is unclear. We present a multimodal imaging protocol aimed to elucidate the angiogenesis-osteogenesis interdependence at the graft proximity.

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that ...

Medical-grade Sterilizable Target for Fluid-immersed Fetoscope Optical Distortion Calibration

We have developed a calibration target for use with fluid-immersed endoscopes within the context of the GIFT-Surg (Guided Instrumentation for Fetal Therapy and Surgery) project. One of the aims of this project is to engineer novel, real-time image processing methods for intra-operative use in the treatment of congenital birth defects, such as spina bifida and the twin-to-twin transfusion syndrome. The developed target allows for the sterility-preserving optical distortion calibration of endoscopes within a few minutes. Good optical distortion calibration and compensation are important for mitigating undesirable effects like radial distortions, which not only hamper accurate imaging using existing endoscopic technology during fetal surgery, but also make acquired images less suitable for potentially very useful image computing applications, like real-time mosaicing. In this paper proposes a novel fabrication method to create an affordable, sterilizable calibration target suitable for use in a clinical setup. This method involves etching a calibration pattern by laser cutting a sandblasted stainless steel sheet. This target was validated using the camera calibration module provided by OpenCV, a state-of-the-art software library popular in the computer vision community.

This article describes the design and development of a sterilizable custom camera optical distortion calibration target for the peri-operative, fluid-immersed calibration of endoscopes during endoscopic interventions.

We have developed a calibration target for use with fluid-immersed endoscopes within the context of the GIFT-Surg (Guided Instrumentation for Fetal ...

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...

Light-sheet Fluorescence Microscopy for the Study of the Murine Heart

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. Traditional light-sheet techniques involve the use of a single illumination beam directed orthogonally at sample tissue. Images of large samples that are produced using a single illumination beam contain stripes or artifacts and suffer from a reduced resolution due to the scattering and absorption of light by the tissue. This study uses a dual-sided illumination beam and a simplified CLARITY optical clearing technique for the murine heart. These techniques allow for deeper imaging by removing lipids from the heart and produce a large field of imaging, greater than 10 x 10 x 10 mm3. As a result, this strategy enables us to quantify the ventricular dimensions, track the cardiac lineage, and localize the spatial distribution of cardiac-specific proteins and ion-channels from the post-natal to adult mouse hearts with sufficient contrast and resolution.

This study uses a dual-sided illumination light-sheet fluorescence microscopy (LSFM) technique combined with optical clearing to study the murine heart.

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. ...