Name: mesoscopic optical imaging of whole mouse heart
Uploaded: 2021-10-14T14:15:00.000+00:00
Duration: 8 min 53 s
Description: Watch this Scientific Journal Video about Mesoscopic Optical Imaging of Whole Mouse Heart at JoVE.com

This project has received fundings from the European Union&#8217;s Horizon 2020 research and innovation programme under grant agreement No 952166 (REPAIR), MUR under the FISR program, project FISR2019_00320 and Regione Toscana, Bando Ricerca Salute 2018, PERCARE project.

All animal handling and procedures were performed in accordance with the guidelines from Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes and conformed to the principles and regulations of the Italian Ministry of Health. The experimental protocol was approved by the Italian Ministry of Health (protocol number 647/2015-PR). All the animals were provided by ENVIGO, Italy. For these experiments, 5 male C57BL/6J mice of 6 months of age were used.
1. Solution preparation
<ol>
	<li>Prepare 4% Paraformaldehyde (PFA) in Phosphate-Buffered Saline (PBS) (pH 7.6) in a chemical hood. Store the 4% PFA aliquots at -20 &#176;C for several months.</li>
	<li>Prepare Hydrogel solution: Mix 4% Acrylamide, 0.05% Bis-acrylamide, 0.25% Initiatior AV-044 in 0.01 M PBS in a chemical hood. Keep the reagents and the solution on ice during the entire preparation. Store the hydrogel aliquots at -20 &#176;C for several months.</li>
	<li>Prepare Clearing solution: Mix 200 mM Boric acid, 4% Sodium Dodecyl-Sulfate (SDS) in deionized water; pH 8.6 in a chemical hood. Store the solution between 21-37 &#176;C to avoid SDS precipitation.</li>
	<li>Prepare fresh Tyrode solution on the day of the experiment: Add 10 mM Glucose, 10 mM HEPES, 113 mM NaCl, 1.2 mM MgCl2, and 4.7 mM KCl; titrate to pH 7.4 using 1 M NaOH.</li>
</ol>
2. Heart isolation
<ol>
	<li>Inject 0.1 mL of 500 I.U. Heparin subcutaneously 30 min before the heart isolation procedure.</li>
	<li>Fill a 30-mL syringe and three 6-cm Petri dishes with fresh Tyrode solution. Make a small rift (3-4 mm in depth) on the border of one of the Petri dishes and place it under a stereoscopic microscope.</li>
	<li>Fix a 1 mm-diameter cannula to the syringe and insert it in the rift of the Petri dish. Make sure there are no air bubbles in the syringe.</li>
	<li>Fill a 20-mL syringe with 4% PFA and keep it in the chemical hood. Prepare an empty Petri dish under the hood.</li>
	<li>Anesthetize the mouse with 3% Isoflurane/oxygen at a flow rate of 1.0 L/min and sacrifice it by cervical dislocation according to animal welfare rules in force.</li>
	<li>After the sacrifice, remove the fur over the chest and open the chest to have full access to the heart.</li>
	<li>Isolate the heart, immerse it in the Petri dish previously filled with 50 mL of Tyrode Solution. Use surgical scissors to cut the aorta immediately near the aortic arch to have the heart exposed.</li>
	<li>Transfer the heart under a stereoscopic microscope and carefully perform the cannulation. Do not insert the cannula too deep into the aorta (no more than 2 mm) to avoid tissue damage.</li>
	<li>Use a little clamp and a suture (size 5/0) to fix the heart to the cannula.</li>
	<li>Perfuse the heart with 30 mL of the Tyrode solution with a constant pressure of 10 mL/min to remove blood from the vessels.</li>
	<li>Detach the cannula from the syringe and place the heart in the Petri dish filled with Tyrode solution. Be careful not to have air bubbles in the cannula; otherwise, remove the air bubbles properly.</li>
	<li>Attach the 20-mL syringe filled with cold 4% PFA to the cannula and perfuse the heart at the same constant pressure.</li>
	<li>Incubate the heart in 10 mL of 4% PFA at 4 &#176;C overnight (O/N). To avoid tissue degradation, perform steps 2.6-2.13 in the shortest time possible.</li>
</ol>
3. Heart clearing
<ol>
	<li>The following day, wash the heart in 0.01 M PBS 3 times at 4 &#176;C for 15 min. 
	NOTE: After this step, the heart can be stored in PBS + 0.01% sodium azide (NaN3) at 4 &#176;C for several months.</li>
	<li>Incubate the heart in 30 mL of Hydrogel solution in shaking (15 rpm) at 4 &#176;C for 3 days.</li>
	<li>Degas the sample at room temperature using a dryer, a vacuum pump, and a tube system that connects the dryer to both the pump and a nitrogen pipeline.
	<ol>
		<li>Place the sample in the dryer and open the vial, keeping the cap on it.</li>
		<li>Close the dryer and remove the oxygen from the tube by opening the nitrogen pipeline.</li>
		<li>Turn on the vacuum&#160;pump to remove the oxygen from the dryer for 10 min.</li>
		<li>Turn off the pump and use the knob of the dryer to open the nitrogen pipeline. Once the pressure is equal to the atmospheric pressure, carefully open the dryer and quickly close the vial.</li>
	</ol>
	</li>
	<li>Keep the heart in the degassed Hydrogel solution at 37 &#176;C for 3 h at rest.</li>
	<li>When the Hydrogel is properly polymerized and appears entirely gelatinous, carefully remove the heart from it and place it in the sample holder.</li>
	<li>Insert the sample holder with the heart in one of the clearing chambers and close it properly to avoid leaks of the clearing solution.</li>
	<li>Switch on the water bath where the clearing solution container is placed and the peristaltic pump to start the recirculation of the clearing solution.</li>
	<li>Change the clearing solution in the container once a week to speed up the clarification procedure.</li>
</ol>
4. Cellular membrane staining
<ol>
	<li>Once the heart appears completely clarified, remove it from the sample holder and wash it in 50 mL of warmed-up PBS for 24 h. Wash again in 50 mL of PBS + 1% of Triton-X (PBS-T 1x) for 24 h.</li>
	<li>Incubate the sample in 0.01 mg/mL Wheat Germ Agglutinin (WGA) - Alexa Fluor 633 in 3 mL of PBS-T 1x in shaking (50 rpm) at room temperature for 7 days.</li>
	<li>After the 7-day incubation, wash the sample in 50 mL of PBS-T 1x at room temperature in shaking for 24 h.</li>
	<li>Incubate the sample in 4% PFA for 15 min and then wash it 3 times in PBS for 5 min each. 
	NOTE: After this step, the heart can be stored in PBS + 0.01% NaN3 at 4 &#176;C for several months.</li>
	<li>Incubate the heart in increasing concentrations of 2,2&#8242;-Thiodiethanol (TDE) in 0.01 M PBS (20% and 47% TDE/PBS) for 8 h each, up to the final concentration of 68% TDE in 0.01 M PBS to provide the required refractive index (RI = 1.46). This is the RI matching medium (RI-medium) to acquire images16&#8288;.</li>
</ol>
5. Heart mounting and acquisition
NOTE: All the components of the optical system are listed in detail in the Table of Materials.
<ol>
	<li>Gently fill about 80% of the external cuvette (quartz, 45 mm &#215; 45 mm &#215; 42.5 mm) with the RI-medium. 
	NOTE: Here, it is possible to use different non-volatile solutions that guarantee a RI of 1.46.</li>
	<li>Gently fill the internal cuvette (quartz, 45 mm &#215; 12.5 mm &#215; 12.5 mm) with the same RI-medium.</li>
	<li>Immerse the sample inside the internal cuvette. The sample incubations described above allow the sample to remain stable inside the RI-medium without being held.</li>
	<li>Gently move the sample to the bottom of the cuvette using thin tweezers and arrange the heart with its longitudinal axis parallel to the cuvette&#39;s main axis to minimize the excitation light path across the tissue during the scanning.</li>
	<li>Gently fix the tailored plug above the internal cuvette with two screws.</li>
	<li>Mount the sample to the microscope stage using the magnets.</li>
	<li>Translate the vertical sample stage manually to immerse the internal cuvette into the external one.</li>
	<li>Turn on the excitation light source (wavelength of 638 nm), setting a low power (in the order of 3 mW).</li>
	<li>Move the sample using the motorized translator to illuminate an inner plane of the tissue.</li>
	<li>Turn on the imaging software (HCImageLive) and set the camera Trigger on External&#160;(light-sheet) mode to drive the acquisition trigger of the camera by the custom software controlling the entire setup.</li>
	<li>Enable Autosave in the Scan Settings panel and set the output folder where the images need to be saved.</li>
	<li>Manually adjust the sample position in the XY plane with the linear translators to move the sample to the center of the FoV of the camera sensor.</li>
	<li>Move the sample along the Z-axis using the linear motorized translator to identify heart borders for tomographic reconstruction.</li>
	<li>Increase the laser power to ~20 mW, ready for the imaging session.</li>
	<li>Start the tomographic acquisition, click the Start button in the Sequence Panel&#160;of the imaging software, and at the same time move the sample along the Z-axis at the constant velocity of 6 &#181;m/s using the motorized translator.</li>
</ol>

<ol>
	<li>Cohn, J. N., Ferrari, R., Sharpe, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+remodeling-concepts+and+clinical+implications:+A+consensus+paper+from+an+International+Forum+on+Cardiac+Remodeling.">Cardiac remodeling-concepts and clinical implications: A consensus paper from an International Forum on Cardiac Remodeling.</a> Journal of the American College of Cardiology. 35, 569-582 (2000).</li><li>Finocchiaro, G., 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=Arrhythmogenic+potential+of+myocardial+disarray+in+hypertrophic+cardiomyopathy:+genetic+basis,+functional+consequences+and+relation+to+sudden+cardiac+death.">Arrhythmogenic potential of myocardial disarray in hypertrophic cardiomyopathy: genetic basis, functional consequences and relation to sudden cardiac death.</a> EP Europace. 2, 1-11 (2021).</li><li>Bishop, M. J., 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=Development+of+an+anatomically+detailed+MRI-derived+rabbit+ventricular+model+and+assessment+of+its+impact+on+simulations+of+electrophysiological+function.">Development of an anatomically detailed MRI-derived rabbit ventricular model and assessment of its impact on simulations of electrophysiological function.</a> American Journal of Physiology - Heart and Circulatory Physiology. 298 (2), 699-718 (2010).</li><li>Bishop, M. J., Boyle, P. M., Plank, G., Welsh, D. G., Vigmond, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modelling+the+role+of+the+coronary+vasculature+during+external+field+stimulation.">Modelling the role of the coronary vasculature during external field stimulation.</a> IEEE Transaction on Biomedical Engineering. 57, 2335-2345 (2010).</li><li>Tainaka, K., 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=Whole-body+imaging+with+single-cell+resolution+by+tissue+decolorization.">Whole-body imaging with single-cell resolution by tissue decolorization.</a> Cell. 159, 911-924 (2014).</li><li>Ueda, H. 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=Whole-brain+profiling+of+cells+and+circuits+in+mammals+by+tissue+clearing+and+light-sheet+microscopy.">Whole-brain profiling of cells and circuits in mammals by tissue clearing and light-sheet microscopy.</a> Neuron. 106, 369-387 (2020).</li><li>Richardson, D. S., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clarifying+tissue+clearing.">Clarifying tissue clearing.</a> Cell. 162, 246-257 (2015).</li><li>Silvestri, L., Costantini, I., Sacconi, L., Pavone, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clearing+of+fixed+tissue:+a+review+from+a+microscopist's+perspective.">Clearing of fixed tissue: a review from a microscopist's perspective.</a> Journal of Biomedical Optics. 21, 081205 (2016).</li><li>Chung, K., 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=Structural+and+molecular+interrogation+of+intact+biological+systems.">Structural and molecular interrogation of intact biological systems.</a> Nature. 497, 332-337 (2013).</li><li>Park, Y. G., 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=Protection+of+tissue+physicochemical+properties+using+polyfunctional+crosslinkers.">Protection of tissue physicochemical properties using polyfunctional crosslinkers.</a> Nature Biotechnology. 37, 73 (2019).</li><li>Ding, 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=Light-sheet+fluorescence+microscopy+for+the+study+of+the+murine+heart.">Light-sheet fluorescence microscopy for the study of the murine heart.</a> Journal of Visualized Experiments: JoVE. (139), e57769 (2018).</li><li>Olianti, C., 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=3D+imaging+and+morphometry+of+the+heart+capillary+system+in+spontaneously+hypertensive+rats+and+normotensive+controls.">3D imaging and morphometry of the heart capillary system in spontaneously hypertensive rats and normotensive controls.</a> Scientific Reports. 10, 1-9 (2020).</li><li>Pianca, N., 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=Cardiac+sympathetic+innervation+network+shapes+the+myocardium+by+locally+controlling+cardiomyocyte+size+through+the+cellular+proteolytic+machinery.">Cardiac sympathetic innervation network shapes the myocardium by locally controlling cardiomyocyte size through the cellular proteolytic machinery.</a> The Journal of Physiology. 597, 3639-3656 (2019).</li><li>Di Bona, A., Vita, V., Costantini, I., Zaglia, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Towards+a+clearer+view+of+sympathetic+innervation+of+cardiac+and+skeletal+muscles.">Towards a clearer view of sympathetic innervation of cardiac and skeletal muscles.</a> Progress in Biophysics and Molecular Biology. 154, 80-93 (2020).</li><li>Voigt, F. F., 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=The+mesoSPIM+initiative+-+open-source+light-sheet+microscopes+for+imaging+cleared+tissue.">The mesoSPIM initiative - open-source light-sheet microscopes for imaging cleared tissue.</a> Nature Methods. 16 (11), 1105-1108 (2019).</li><li>Costantini, I., 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+versatile+clearing+agent+for+multi-modal+brain+imaging.">A versatile clearing agent for multi-modal brain imaging.</a> Scientific Reports. 5, 9808 (2015).</li><li>Judd, J., Lovas, J., Huang, G. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation,+culture+and+transduction+of+adult+mouse+cardiomyocytes.">Isolation, culture and transduction of adult mouse cardiomyocytes.</a> Journal of Visualized Experiments: JoVE. (114), e54012 (2016).</li><li>Yi, F., 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=Microvessel+prediction+in+H&amp;E+stained+pathology+images+using+fully+convolutional+neural+networks.">Microvessel prediction in H&amp;E stained pathology images using fully convolutional neural networks.</a> BMC Bioinformatics. 19 (1), 64 (2018).</li><li>Susaki, E. A., 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=Versatile+whole-organ/body+staining+and+imaging+based+on+electrolyte-gel+properties+of+biological+tissues.">Versatile whole-organ/body staining and imaging based on electrolyte-gel properties of biological tissues.</a> Nature Communications. 11 (1), 1982 (2020).</li></ol>

Nothing to disclosure.

In this work, a successful approach to clear, stain, and image a whole mouse heart at high resolution was introduced. First, a tissue transformation protocol (CLARITY) was optimized and performed, slightly modified for its application on the cardiac tissue. Indeed, to obtain an efficient reconstruction in 3D of a whole heart, it is essential to prevent the phenomenon of light scattering. The CLARITY methodology allows us to obtain a highly transparent intact heart, but it requires long incubation times when performed passively (about 5 months). With respect to the brain, the cardiac tissue is not suitable for an active clearing, which takes advantage of an electric field. Even at low voltages, the electric field leads to damages and tissue breakages. Here, a passive clearing approach was optimized to obtain a completely cleared heart in about 3 months. After isolating and cannulating the heart through the proximal aorta, the CLARITY methodology was performed as described in section 3 of the protocol. To speed up the procedure, a homemade passive clearing setup was arranged (Figure 1), which ended up decreasing the timing of tissue clearing by about 40%. The setup is composed of a container for the clearing solution, a water bath, a peristaltic pump, several chambers containing different sample holders, capsule filters for each chamber, and a tubing system for the recirculation of the solution. The pump extracts and circulates the solution from the container in succession through each of the chambers, where the samples are held for clearing. Before entering the chambers, the solution flows through a capsule filter to trap the lipids flushed away from tissues during the clearing. The optimal temperature for the clearing solution, between 37-45 &#176;C, is maintained within the chambers during the recirculation by keeping the solution container in a water bath at 50 &#176;C. It is advised to change the clearing solution in the container once a week during the procedure. All components used are listed in detail in the Table of Materials. The optimized solution presented here allows us to obtain a whole passively cleared mouse heart in a significantly shorter time with respect to the standard passive clearing technique, thus reducing the required experimental time without damaging the organ. The staining approach was also optimized for homogeneous labeling of the cellular membranes and endothelium, using a fluorescent lectin (WGA - Alexa Fluor 633).
The heart cytoarchitecture has been reconstructed by developing a dedicated mesoSPIM that axially sweeps the light-sheet across the sample (https://mesospim.org). The custom-made fluorescence light-sheet microscope&#160;(Figure 3) was able to rapidly acquire images of a mesoscopic FoV (of the order of millimeters) with micrometric resolution. In this way, single cardiomyocytes can be resolved and mapped into a 3D reconstruction of the entire organ. The microscope illuminates the cleared sample with a light-sheet, dynamically generated by scanning a laser beam at 638 nm using a galvanometric mirror. A sCMOS camera characterizes the detection arm in a 2x magnification scheme which enables it to acquire the entire FoV in a single scan. The fluorescence signal was selected by placing a long-pass filter after the objective. The camera was set to work in rolling shutter mode: at any time, the lines of active camera pixels (i.e., exposed to the image) are synchronized with the in-plane shift of the focal band of the light-sheet, performed by an electrically tunable lens. This approach maximized the optical sectioning capability in the whole FoV by only acquiring images in the thinnest part of the focused light-sheet. This solution differs from conventional configurations, where acquisition involves the entire range of focal depth of the light-sheet, preventing peak optical sectioning resolution in large part of the FoV. An integrated sample stage supports cuvettes, thereby optimizing positioning and enabling axial movement of the sample during the imaging process. In this way, tomographic reconstructions are possible by acquiring consecutive internal sections. The images obtained have a mesoscopic FoV and a micrometric resolution, while the acquisition time required for a whole mouse heart is ~ 15 min. The synchronization between the camera rolling shutter and the excitation light beam sweeping the FoV allows acquiring the entire image plane with a high spatial resolution (Figure 4). This allows direct reconstruction of the sample in a single tomographic acquisition, without the necessity of sample radial displacement and multi-adjacent-stacks-based imaging. Notably, the microscope allowed the reconstruction of the entire organ of about (10 mm x 6 mm x 6 mm) in a single imaging session, with a near-isotropic voxel size and a sufficient signal-to-background ratio to resolve single cells across the whole organ potentially.
It is noteworthy that the proposed protocol presents some critical steps that must be performed carefully to achieve good results. In particular, the cannulation of the heart through the proximal aorta can be quite difficult, but it is an essential step to wash and fix the organ properly. Judd et al.17, showed how to perform this step effectively. Moreover, the degassing procedure needed by the CLARITY protocol is quite complex too, but it is essential for tissue preservation; if this step is not performed properly, the tissue could encounter damages and decay during the incubation in clearing solution.
Furthermore, although the presented experimental workflow is suitable for small fluorescent probes, the use of immunohistochemistry does not always provide good efficiency in the staining due to the higher molecular weight of the antibodies. Each immunostaining protocol requires proper optimization, and different approaches have been conceived to improve the antibody penetration, for example, tissue expansion18 and/or variations in pH and ionic strength19.
The mesoSPIM setup also presents two main limitations: i) the light-sheet preservation across the sample is strongly dependent on the tissue transparency, and ii) the dimension of the camera sensor limits the FoV. Guaranteeing a perfect refractive index matching inside the entire heart is very challenging, and small variations on the refractive index can produce light scattering, leading to degradation of the image quality. In this respect, a dual-side illumination scheme can be introduced. Two excitation arms can generate two distinct and aligned dynamic light sheets with maximally focused illumination by alternating the illumination from one side to the other of the specimen. Also, the FoV can be improved by using a new generation high-resolution back-illuminated sCMOS with very large sensors in combination with high numerical aperture telecentric lenses with low field distortion. This implementation would allow us to reconstruct bigger organs or expanded tissues maintaining the same optical section capability and thus producing micron-scale 3D images of centimeter-sized cleared samples.
Although the presented protocol still requires a long time for sample preparation and a high level of transparency to obtain a reliable cytoarchitecture reconstruction of the entire organ, the main significance of the approach resides in the improvements of the clearing protocol and the capability to perform mesoscopic reconstruction in a single scan at micrometric resolution. In the future, these advances can be combined with a multi-staining protocol to achieve whole-organ reconstruction integrating different biological structures.

Structural remodeling associated with cardiac diseases can affect electrical conduction and introduce electromechanical dysfunctions of the organ1,2. Current approaches used to predict functional alterations commonly employ MRI and DT-MRI to obtain an overall reconstruction of fibrosis deposition, vascular tree, and fiber distribution of the heart, and they are used to model preferential action potential propagation (APP) paths across the organ3,4. These strategies can provide a beautiful overview of the heart organization. However, their spatial resolution is insufficient to investigate the impact of structural remodeling on cardiac function at the cellular level.
Placing this framework at a different order of magnitude, where single cells can play individual roles on action potential propagation, is challenging. The main limitation is the inefficiency of standard imaging methods to perform high-resolution imaging (micrometric resolution) in massive (centimeter-sized) tissues. In fact, imaging biological tissues in 3D at high resolution is very complicated due to tissue opaqueness. The most common approach to perform 3D reconstructions in entire organs is to prepare thin sections. However, precise sectioning, assembling, and imaging require significant effort and time. An alternative approach that does not demand cutting the sample is to generate a transparent tissue. During the last years, several methodologies for clarifying tissues have been proposed5,6,7,8&#8288;. The challenge to produce massive, transparent, and fluorescently-labeled tissues has been recently achieved by developing true tissue transformation approaches (CLARITY9&#8288;, SHIELD10&#8288;). In particular, the CLARITY method is based on the transformation of an intact tissue into a nanoporous, hydrogel-hybridized, lipid-free form that enables to confer high transparency by the selective removal of membrane lipid bilayers. Notably, this method has been found successful also in cardiac preparation11,12,13,14&#8288;. However, since the heart is too fragile to be suitable for an active clearing, it must be cleared using the passive approach, which requires a long time to confer complete transparency.
In combination with advanced imaging techniques like light-sheet microscopy, CLARITY has the potential to image 3D massive heart tissues at micrometric resolution. In light-sheet microscopy, the illumination of the sample is performed with a thin sheet of light confined in the focal plane of the detection objective. The fluorescence emission is collected along an axis perpendicular to the illumination plane15. The detection architecture is similar to widefield microscopy, making the acquisition much faster than laser scanning microscopes. Moving the sample through the light sheet permits obtaining a complete tomography of big specimens, up to centimeter-sized samples. However, due to the intrinsic properties of the Gaussian beam, it is possible to obtain a very thin (of the order of a few microns) light-sheet only for a limited spatial extension, thus drastically limiting the field of view (FoV). Recently, a novel excitation scheme has been introduced to overcome this limitation and applied for brain imaging, allowing 3d reconstructions with isotropic resolution16.
In this paper, a passive clearing approach is presented, enabling a significant reduction of the clearing timing needed by the CLARITY protocol. The methodological framework described here allows reconstructing a whole mouse heart with micrometric resolution in a single tomographic scan with an acquisition time in the order of minutes.

<table><tbody><tr><td>2-2&amp;rsquo; Thiodiethanol</td><td>Sigma-Aldrich</td><td>166782</td><td></td></tr><tr><td>Acrylamide</td><td>Bio-Rad</td><td>61-0140</td><td></td></tr><tr><td>AV-044 Initiator</td><td>Wako Chemicals</td><td>AVP5874</td><td></td></tr><tr><td>Bis-Acrylamide</td><td>Bio-Rad</td><td>161-042</td><td></td></tr><tr><td>Boric Acid</td><td>Sigma-Aldrich</td><td>B7901</td><td></td></tr><tr><td>Camera</td><td>Hamamatsu</td><td>Orca flash 4.0 v3</td><td></td></tr><tr><td>Camera software</td><td>Hamamatsu</td><td>HC Image</td><td></td></tr><tr><td>Collimating lens</td><td>Thorlabs</td><td>AC254-050-A-ML</td><td></td></tr><tr><td>Detection arm</td><td>Integrated optics</td><td>0638L-15A-NI-PT-NF</td><td></td></tr><tr><td>Excitation lens</td><td>Nikon</td><td>91863</td><td></td></tr><tr><td>Extera&amp;igrave;nal quartz cuvette</td><td>Portmann Instruments</td><td>UQ-753</td><td></td></tr><tr><td>Fold mirrors</td><td>Thorlabs</td><td>BBE1-E02</td><td></td></tr><tr><td>Galvanometric mirror</td><td>Thorlabs</td><td>GVS211/M</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G8270</td><td></td></tr><tr><td>HCImage Live</td><td>Hamamatsu</td><td>4.6.1.19</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375</td><td></td></tr><tr><td>Internal quartz cuvette</td><td>Portmann Instruments</td><td>UQ-204</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P4504</td><td></td></tr><tr><td>Laser source</td><td>Integrated Optics</td><td>0638L-15A-NI-PT-NF</td><td></td></tr><tr><td>Long-pass filter</td><td>Thorlabs</td><td>FELH0650</td><td></td></tr><tr><td>Magnetic base</td><td>Thorlabs</td><td>KB25/M</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Chem-Lab</td><td>CI-1316-0250</td><td></td></tr><tr><td>Motorized traslator</td><td>Physisk Instrument</td><td>M-122.2DD</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>59888</td><td></td></tr><tr><td>Objective</td><td>Thorlabs</td><td>TL2X-SAP</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Agar Scientific</td><td>R1018</td><td></td></tr><tr><td>Phosphate Buffer Solution</td><td>Sigma-Aldrich</td><td>P4417</td><td></td></tr><tr><td>Polycap AS</td><td>Whatman</td><td>2606T</td><td></td></tr><tr><td>Relay lens</td><td>Qioptiq</td><td>G063200000</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate</td><td>Sigma-Aldrich</td><td>L3771</td><td></td></tr><tr><td>Tube lens</td><td>Thorlabs</td><td>ACT508-200-A-ML</td><td></td></tr><tr><td>Tunable lens</td><td>Optotune</td><td>EL-16-40-TC-VIS-5D-1-C</td><td></td></tr><tr><td>Vacuum pump</td><td>KNF Neuberger Inc</td><td>N86KT.18</td><td></td></tr><tr><td>Water bath</td><td>Memmert</td><td>WTB</td><td></td></tr></tbody></table>

mesoscopic optical imaging of whole mouse heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

The developed passive clearing setup allows to obtain a cleared adult mouse heart (with a dimension of the order 10 mm x 6 mm x 6 mm) in about 3 months. All the components of the setup are mounted as shown in Figure 1. The negligible temperature gradient between each clearing chamber (of the order of 3&#176;C) allows maintaining the temperature in a proper range across all chambers.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62795/62795fig01.jpg" /> 
Figure 1:&#160;Schematic of the passive clearing setup. The clearing solution (after being filtered) circulates in succession through the sample chambers with the help of the peristaltic pump. The maintenance of the solution container in a water bath set at 50 &#176;C allows the solution temperature to be between 37-45 &#176;C within the chambers. Image created with Biorender.com. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62795/62795fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure 2&#160;shows the result of the clearing process of an entire heart. As already reported by Costantini et al.16, the combination of the CLARITY methodology with TDE as RI-medium does not significantly change the sample&#39;s final volume nor leads to anisotropic deformation of the specimen.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62795/62795fig02.jpg" /> 
Figure 2: Representative image of a heart before (on the left) and after (on the right) the CLARITY protocol. The hearts become fully transparent and slightly oversized. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62795/62795fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Once the heart was cleared, cellular membranes were stained with an Alexa Fluor 633-conjugated WGA to perform the cytoarchitecture reconstruction of the entire organ. The custom-made fluorescence light-sheet microscope (Figure 3) was able to ensure 3D micron-scale resolution across the entire FoV.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62795/62795fig03.jpg" /> 
Figure 3: MesoSPIM. CAD rendering of the custom-made fluorescence light-sheet microscope. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62795/62795fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Considering the numerical aperture (NA = 0.1) of the detection optics, the radial (XY) Point Spread Function (PSF) of the system can be estimated in the order of 4-5 &#181;m. On the other hand, the excitation optics produce a light-sheet with a minimum waist of about 6 &#181;m (Full width half maximum, FWHM) that diverges up to 175 &#181;m at the edge of the FoV (Figure 4A-C). The synchronization of the camera rolling shutter with the axial scan of the laser beam ensured&#160;to collect the emission signal only in the sample portion excited with the waist of the light-sheet, resulting in an average FWHM of about 6.7 &#181;m along the entire FoV (Figure 4B-D).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62795/62795fig04.jpg" /> 
Figure 4: Light-sheet generation and characterization. (A) An excitation light-sheet generated with a laser source of 638 nm is focused on the center of the Field of View (FoV) and acquired with a pixel size of 3.25 &#181;m and an Exposure Time of 10 ms. Light intensity is normalized and reported with a colormap. The Full Width Half Maximum (FWHM) of the light intensity profile is evaluated in 15 different positions along the FoV. Results are shown in C. (B) Image of the excitation light-sheet generated by the synchronization between the camera rolling shutter operating at 1.92 Hz and the light beam position driven by the tunable lens. The FWHM of the light intensity profile is evaluated along the FoV and results are shown in D. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62795/62795fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The Z-PSF of the microscope was also estimated by a tomographic reconstruction of the fluorescent nanosphere (Figure 5). An FWHM of 6.4 &#181;m can be estimated by the fit, in good agreement with the previous assessment.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62795/62795fig05.jpg" /> 
Figure 5: Point Spread Function in the Z-axis. The Point Spread Function (PSF) of the optical system is estimated by imaging fluorescent sub-micron-scale nanospheres (excited with a light sheet with a wavelength of 638 nm) with a pixel size of 3.25 &#181;m &#215; 3.25 &#181;m &#215; 2.0 &#181;m. PSF intensity profile along the optical axis (Z) is represented as black dots. PSF profile is fitted with a Gaussian function with &#181; = 18.6 &#181;m and &#963; = 2.7 &#181;m. The FWHM of the PSF estimated by the fit is 6.4 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62795/62795fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Owing to the high transparency of the tissue, it was possible to illuminate the whole heart without significant distortion of the axially scanned light-sheet at an excitation wavelength of 638 nm. The fluorescence signal was collected by the sCMOS sensor operating at 500 ms of exposure time and a frame rate of 1.92 Hz. Based on previous quantification, the tomographic acquisition was performed using a Z-scan velocity of 6 &#181;m/s, and assuming a frame rate of 1.92 Hz, one frame every 3.12 &#181;m was acquired, oversampling the system Z-PSF by about two times. Two representative frames (on the coronal and transverse planes) of the left ventricle chamber are shown in Figure 6. This result confirms the potentiality of the system to resolve single cellular membranes in three dimensions with a sufficient Signal/Noise ratio in the entire organ (Figure 6).
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62795/62795fig06.jpg" /> 
Figure 6: Mouse heart tissue reconstruction. The clarified heart was stained with WGA conjugated to Alexa Fluor 633 and excited by a laser source with a wavelength of 638 nm. (A) Coronal and (B) transverse representative sections. (C-D) Tissue transformation produces high tissue transparency, allowing to resolve small structures in the wall depth. The optical system shows an axial resolution sufficient to resolve micrometric structures (panel. D). (E) 3D low-resolution heart rendering. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62795/62795fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Mesoscopic Optical Imaging of Whole Mouse Heart at JoVE.com

Mesoscopic Optical Imaging of Whole Mouse Heart

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

mesoscopic-optical-imaging-of-whole-mouse-heart

Nanyang Technological University

Research

JoVE Journal

Biology

2.0K Views.  European Laboratory for Non-Linear Spectroscopy. Deze methodologie combineert verbetering van het clearingprotocol met de oogcapaciteit van optische sectioning van de Mesos PIM-technologie, waardoor we een mesoscopische reconstructie met micrometrische resolutie kunnen uitvoeren. Het biedt de mogelijkheid om massieve hartweefsels of hele muizenharten in één scan in beeld te brengen zonder de oorspronkelijke driedimensionale weefselorganisatie te verliezen. Nadat het hart is gecannuleerd door proximale aorta gewassen in tyro-oplossing en g...

Video: Mesoscopic Optical Imaging of Whole Mouse Heart

Murine Echocardiography and Ultrasound Imaging

Rodent models of cardiac pathophysiology represent a valuable research tool to investigate mechanism of disease as well as test new therapeutics.1 Echocardiography provides a powerful, non-invasive tool to serially assess cardiac morphometry and function in a living animal.2 However, using this technique on mice poses unique challenges owing to the small size and rapid heart rate of these animals.3 Until recently, few ultrasound systems were capable of performing quality echocardiography on mice, and those generally lacked the image resolution and frame rate necessary to obtain truly quantitative measurements. Newly released systems such as the VisualSonics Vevo2100 provide new tools for researchers to carefully and non-invasively investigate cardiac function in mice. This system generates high resolution images and provides analysis capabilities similar to those used with human patients. Although color Doppler has been available for over 30 years in humans, this valuable technology has only recently been possible in rodent ultrasound.4,5 Color Doppler has broad applications for echocardiography, including the ability to quickly assess flow directionality in vessels and through valves, and to rapidly identify valve regurgitation. Strain analysis is a critical advance that is utilized to quantitatively measure regional myocardial function.6 This technique has the potential to detect changes in pathology, or resolution of pathology, earlier than conventional techniques. Coupled with the addition of three-dimensional image reconstruction, volumetric assessment of whole-organs is possible, including visualization and assessment of cardiac and vascular structures. Murine-compatible contrast imaging can also allow for volumetric measurements and tissue perfusion assessment.

This video demonstrates use of a rail-mounted high-frequency ultrasound probe to perform echocardiography on an anesthetized mouse. The methods describe both conventional two-dimensional and M-mode measurements of cardiac function in addition to newer, more powerful tools such as color Doppler, strain analysis, as well as general and targeted contrast imaging.

Rodent models of cardiac pathophysiology represent a valuable research tool to investigate mechanism of disease as well as test new therapeutics.1  ...

Medicine

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Bioengineering

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

Neuroscience

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. ...

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.

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 ...

Light-Sheet Microscopy and Tissue Clearing for Comprehensive Cardiac Imaging

Light-Sheet Imaging to Reveal Cardiac Structure in Rodent Hearts

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights into fundamental cardiac physiology and pathologic responses. We hereby delve into the development and implementation of the LSM technique to elucidate the micro-architecture of the heart in mouse models. The methodology integrates a customized LSM system with tissue clearing techniques, mitigating light scattering within cardiac tissues for volumetric imaging. The combination of conventional LSM with image stitching and multiview deconvolution approaches allows for the capture of the entire heart. To address the inherent trade-off between axial resolution and field of view (FOV), we further introduce an axially swept light-sheet microscopy (ASLM) method to minimize out-of-focus light and uniformly illuminate the heart across the propagation direction. In the meanwhile, tissue clearing methods such as iDISCO enhance light penetration, facilitating the visualization of deep structures and ensuring a comprehensive examination of the myocardium throughout the entire heart. The combination of the proposed LSM and tissue clearing methods presents a promising platform for researchers in resolving cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Author Spotlight: Customized Light-Sheet Imaging for Investigating Myocardial Structures in Rodent Hearts

The protocol utilizes advanced light-sheet microscopy along with adapted tissue clearing methods to investigate intricate cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights ...

Dynamic Measurement and Imaging of Capillaries, Arterioles, and Pericytes in Mouse Heart

Coronary arterial tone along with the opening or closing of the capillaries largely determine the blood flow to cardiomyocytes at constant perfusion pressure. However, it is difficult to monitor the dynamic changes of the coronary arterioles and the capillaries in the whole heart, primarily due to its motion and non-stop beating. Here we describe a method that enables monitoring of arterial perfusion rate, pressure and the diameter changes of the arterioles and capillaries in mouse right ventricular papillary muscles. The mouse septal artery is cannulated and perfused at a constant flow or pressure with the other dynamically measured. After perfusion with a fluorescently labeled lectin (e.g., Alexa Fluor-488 or -633 labeled Wheat-Germ Agglutinin, WGA), the arterioles and capillaries (and other vessels) in right ventricle papillary muscle and septum could be readily imaged. The vessel-diameter changes could then be measured in the presence or absence of heart contractions. When genetically encoded fluorescent proteins were expressed, specific features could be monitored. For examples, pericytes were visualized in mouse hearts that expressed NG2-DsRed. This method has provided a useful platform to study the physiological functions of capillary pericytes in heart. It is also suitable for studying the effect of reagents on the blood flow in heart by measuring the vascular/capillary diameter and the arterial luminal pressure simultaneously. This preparation, combined with a state-of-the-art optic imaging system, allows one to study the blood flow and its control at cellular and molecular level in the heart under near-physiological conditions.

Presented here is a protocol to study the coronary microcirculation in living murine heart tissue by ex vivo monitoring of the arterial perfusion pressure and flow that maintains the pressure, as well as vascular tree components including the capillary beds and pericytes, as the septal artery is cannulated and pressurized.

Coronary arterial tone along with the opening or closing of the capillaries largely determine the blood flow to cardiomyocytes at constant perfusion ...

High-resolution Optical Mapping of the Mouse Sino-atrial Node

Sino-atrial node (SAN) dysfunctions and associated complications constitute important causes of morbidity in patients with cardiac diseases. The development of novel pharmacological therapies to cure these patients relies on the thorough understanding of both normal physiology and pathophysiology of the SAN. Among the studies of cardiac pacemaking, the mouse SAN is widely used due to its feasibility for modifications in the expression of different genes that encode SAN ion channels or calcium handling proteins. Emerging evidence from electrophysiological and histological studies has also proved the representativeness and similarity of the mouse SAN structure and functions to larger mammals, including the presence of specialized conduction pathways from the SAN to the atrium and a complex pacemakers' hierarchy within the SAN. Recently, the technique of optical mapping has greatly facilitated the exploration and investigation of the origin of excitation and conduction within and from the mouse SAN, which in turn has extended the understanding of the SAN and benefited clinical treatments of SAN dysfunction associated diseases. In this manuscript, we have described in detail how to perform the optical mapping of the mouse SAN from the intact, Langendorff-perfused heart and from the isolated atrial preparation. This protocol is a useful tool to enhance the understanding of mouse SAN physiology and pathophysiology.

Here, we present a protocol for optical mapping of electrical activity from the mouse right atrium and especially the sino-atrial node, at a high spatial and temporal resolution.

Sino-atrial node (SAN) dysfunctions and associated complications constitute important causes of morbidity in patients with cardiac diseases. The ...

Cardiac Magnetic Resonance Imaging

Source: Frederick W. Damen and <a href="https://www-jove-com.remotexs.ntu.edu.sg/author/Craig+J._Goergen">Craig J. Goergen</a>, Weldon School of Biomedical Engineering, Purdue University, West Lafayette, Indiana
In this video, high field, small-bore magnetic resonance imaging (MRI) with physiological monitoring is demonstrated to acquire gated cine loops of the murine cardiovascular system. This procedure provides a basis for assessing left-ventricular function, visualizing vascular networks, and quantifying motion of organs due to respiration. Comparable small animal cardiovascular imaging modalities include high-frequency ultrasound and micro-computed tomography (CT); however, each modality is associated with trade-offs that should be considered. While ultrasound does provide high spatial and temporal resolution, imaging artifacts are common. For example, dense tissue (i.e., the sternum and ribs) can limit imaging penetration depth, and hyperechoic signal at the interface between gas and liquid (i.e., pleura surrounding the lungs) can blur contrast in nearby tissue. Micro-CT in contrast does not suffer from as many in-plane artifacts, but does have lower temporal resolution and limited soft-tissue contrast. Furthermore, micro-CT uses X-ray radiation and often requires the use of contrast agents to visualize vasculature, both of which are known to cause side effects at high doses including radiation damage&#160; and renal injury. Cardiovascular MRI provides a nice compromise between these techniques by negating the need for ionizing radiation and providing the user with the ability to image without contrast agents (although contrast agents are often used for MRI).
This data was acquired with a triggering Fast Low Angle SHot (FLASH) MRI sequence that was gated off of the R-peaks in the cardiac cycle and expiratory plateaus in respiration. These physiological events were monitored through subcutaneous electrodes and a pressure-sensitive pillow that was secured against the abdomen. To ensure the mouse was properly warmed, a rectal temperature probe was inserted and used to control the output of a MRI-safe heating fan. Once the animal was inserted into the bore of the MRI scanner and navigation sequences were run to confirm positioning, the gated FLASH imaging planes were prescribed and data acquired. Overall, high field MRI is a powerful research tool that can provide soft tissue contrast for the study of small animal disease models.

Cardiac MRI: Principle and Imaging

Source: Frederick W. Damen and Craig J. Goergen, Weldon School of Biomedical Engineering, Purdue University, West Lafayette, Indiana
In this video, high ...

Education

JoVE Science Education

Engineering

Biomedical Engineering

Mouse Echocardiography: A Non-invasive Technique to Assess Cardiac Morphology

Source: Baudouy, D., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/55843/" target="_blank">Echocardiographic and Histological Examination of Cardiac Morphology in the Mouse</a>. J. Vis. Exp. (2017).
This video describes a protocol to perform echocardiography on a laboratory mouse.

Source: Baudouy, D., et al. Echocardiographic and Histological Examination of Cardiac Morphology in the Mouse. J. Vis. Exp. (2017).
This video describes a ...

Mesoscopic Optical Imaging of Whole Mouse Heart

Samenvatting

Meer video's verkennen

Hoofdstukken in deze video