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

The main advantage of this technology is that it allows for rapid imaging of either the mouse heart. And it can be used to observe the presence of air channels after gene therapy. Although this method can provide insight into development cardiology, it can also be applied to other systems, such as neurology and pulmonology.

Generally, individuals will struggle with this technique because mounting an imaging of the adult mouse heart can be difficult, and it's different from other, traditional techniques, such as confocal and inverted microscopy. Place a continuous wave laser with three wavelengths of 405 nanometers, 473 nanometers, and 532 nanometers. Then, affix mirror one and align it with the mirror plane at 45 degrees to the beam.

This will direct the laser toward the beam splitter, which forms the dual-sided illumination setup. Next, pass the beam through a 50 millimeter diameter neutral density filter, a beam expander, and a 25 millimeter diameter pin hole, all positioned 150 millimeters from each other. Pass the beam through a 50-50 beam splitter, placed 150 millimeters from the pinhole.

Next, place mirror two 150 millimeters from the beam splitter, at 90 degrees to the forward beam, and align it so that its mirror plane is at a 45 degree angle to the beam. Then, place mirror three 100 millimeters from mirror two, and align it so that it's mirror plane is at a 45 degree angle to the beam reflected from mirror two. Use this reflected beam to form one side of the dual illumination light sheet.

On the far side of the beam splitter, place mirror five so that it's mirror plane is at 45 degrees to the beam that is emitted in a forward direction. Use the beam emitted from mirror five to form the second side of the dual illumination light sheet. Next, set up the dual-sided illumination system in systemic fashion.

Place cylindrical lens two 150 millimeters away from mirror three, and another identical cylindrical lens 150 millimeters away from mirror five, on the other side of the dual illumination setup. Next, place two mirrors, each in line with the cylindrical lenses, at distances of 50 millimeters to reflect the beam at 90 degrees. On both sides of the light sheet, form an achromatic doublet from a pair of lenses, with the first lens being placed 100 millimeters from the previous mirror, and having a one inch diameter and a focal length of 100 millimeters.

The second lens should be placed 160 millimeters from lens one, with a diameter of one inch, and a focal length of 60 millimeters. Then, place the illumination objectives one and two 150 millimeters from the previous achromatic doublets, so that they are in line with the beam. The beam emitted from the objectives forms the light sheet for imaging the samples.

First, prepare the fluorescent bead sample by diluting a 0.53 micrometer bead solution, one to 150, 000, in a pre-warmed refractive index matching solution, with one percent low melt pointing agarose. Next, use a piece of borosilicate glass tubing with an inner diameter of 12 millimeters, and an outer diameter of 18 millimeters, to a length of 30 millimeters. Pipette the bead agarose solution in the borosilicate tubing and allow the agarose to solidify at room temperature.

Now, fill a 3-D printed ABS chamber with a 99.5 percent gylcerol solution. Place the borosilicate glass tubing, containing the beads, inside the chamber. Attach a 3-D motorized translational stage to the borosilicate glass tubing to control the movement and orientation of the sample within the ABS chamber.

Then, use custom designed software to acquire images using the sCMOS camera, at a rate of 30 frames per second. Using the motor controller, move the sample one millimeter in the lateral direction, and acquire images at each one millimeter increment, until the entire sample has been imaged. Stack the acquired images using a visualization software, and measure the point spread function of the system using these bead images.

Use a refractive index matching solution, with a pH of 7.5, and dissolve one percent agarose into the matching solution. Place a cleared adult mouse heart sample into the refractive index matching solution, with one percent agarose dissolved. Then, insert the sample into borosilicate glass tubing, and allow the agarose to solidify at room temperature.

Attach a 3-D motorized translational stage to the borosilicate glass tubing to control the movement and orientation of the sample within a 3-D printed ABS chamber. Position the sample so that it is in the center of the Gaussian beam, created by the dual illumination system. Then, set the acquisition rate of the sCMOS camera to 30 frames per second.

Now, using the motor controller, move the sample one millimeter in the axial direction and acquire images at each one millimeter increment. Continue until the entire sample has been imaged. Stack the acquired images using a visualization software.

Develop the 3-D images using these image stacks. To accomplish this, deconvolve the point spread function determined previously, and utilize it for the acquired image stacks. Finally, set a pixel threshold intensity value to observe the contours of the heart, and add pseudo-color to the images, based on this gray-scale intensity.

At day one post-natal, one can visualize the valves, atrium, ventricle, pectinate muscle, and trabeculation, among other features. At day seven post-natal, the features are even more defined. The atrium, ventricle, ventricular cavity dimensions, and ventricle wall thickness are all evident.

To study cardiomyocyte differentiation within the heart, heterozygous knock-in mice were cre-labeled to show cardiomyocytes. The labeled cells are shown here, in each plane direction. The three planes were merged, to create a 3-D rendering of the heart.

The red circled region within the inset shows two translucent mouse hearts after undergoing the clarity technique and being inserted into the borosilicate glass tubing. In addition to studying post-natal mice, the imaging system can also be used to study the adult mouse heart. Here, GFP tagged ROMK channels are shown at 7.5 months.

These were mainly found in the ventricular wall. Once mastered, this technique can be done in one to two minutes. When attempting this procedure, it is important to remove all the bubbles around the sample in the tube.

Other methods, such as auto-segmentation of the image stacks, can be performed to answer additional questions related to cardiovascular injury and regeneration. After its development, this technique was used by researchers to study the cardiac architecture in post-natal and adult developmental stages in mice.