Medicine
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Quantitative Analysis of Cellular Composition in Advanced Atherosclerotic Lesions of Smooth Muscle Cell Lineage-Tracing Mice
Summary February 20th, 2019
We propose a standardized protocol to characterize the cellular composition of late-stage murine atherosclerotic lesions including systematic methods of animal dissection, tissue embedding, sectioning, staining, and analysis of brachiocephalic arteries from atheroprone smooth muscle cell lineage tracing mice.
Transcript
Our protocol helps investigators to implement standardized experimental pipelines for performing phenotypic characterization of similar populations by immunofluorescent staining. With this technique, researchers can rigorously analyze the similar composition of complex disease tissues and perform qualitative and quantitative analyses of the phenotypes of the cell types of interest. Demonstrating the procedure will be Sidney Mahan, a technician from my laboratory.
To prepare the animal for brachiocephalic artery harvest, cut the peritoneum up to the sternum, taking care not to damage the abdominal tissues. And make two cuts at the mid-axillary line through the thorax, taking care not to damage the heart and lungs. Then, cut the diaphragm to partially expose the heart.
To profuse the mouse, install a gravity profusion system such that the pressure of the profusion fluid is equivalent to the average murine blood pressure to allow a consistent profusion and maintenance of the vessel morphology. Then, profuse the animal with five milliliters of room temperature PBS, 10 milliliters of 4%room temperature paraformaldehyde, and five additional milliliters of room temperature PBS. Connect a 23 or 25 gauge butterfly needle to the gravity profusion system, and run PBS through the needle to expel the air from the tubes before inserting the needle into the left ventricle.
After securing the needle in place, use iris scissors to make a less than two centimeter incision in the right atrium. Harvest the tissues of interest into fresh 4%paraformaldehyde. To expose the brachiocephalic artery, use scissors to make an incision down the midline of the sternum through the manubrium and use forceps to pull both sides of the ribcage to fully open the chest cavity.
Next, place the animal under a dissecting microscope to partially visualize the carotids, and use fine tweezers to pull the muscles, connective tissue, and fat until the right carotid, brachiocephalic artery, subclavian bifurcation, and aortic arch are cleaned and isolated of the surrounding connective tissue and fat. Next, use forceps to grab the right carotid below its bifurcation and make an initial cut above the forceps. Still, holding the carotid, make a second cut through the subclavian artery and make the final two cuts through the aortic arch on either side of the brachiocephalic artery.
Then, collect any other vascular tissues of interest and place the brachiocephalic artery and the other tissues in 4%paraformaldehyde solution overnight at room temperature. For immunofluorescent staining of the brachiocephalic artery, use a microtome to obtain 10 micrometer sections through the paraffin-embedded tissue sample until it can be confirmed by the light microscopy that the aortic arch has been sectioned through and the brachiocephalic artery is visible. Adjust the orientation of the block until the tissue is completely perpendicular to the microtome blade and obtain three serial 10 micrometer thick sections of the brachiocephalic artery per glass slide, until the subclavian bifurcation is reached.
When all of the sections have been collected, hydrate the tissue samples under a chemical hood with two five minute xylene immersions, followed by a five minute immersion ethanol series as indicated, and two five-minute deionized water immersions. Next, incubate slides in antigen retrieval solution, according to standard protocols, followed by heating in a microwave for 20 minutes. After a one hour cool down at room temperature, use a hydrophobic pen to encircle each tissue section.
And block any non-specific binding with blocking buffer for one hour at room temperature. At the end of the incubation, label the tissue samples overnight at four degrees Celsius with the primary antibody cocktail of interest, or isotype matched IgG control antibodies. The next morning, wash slides three times in PBS for five minutes per wash, followed by a one hour incubation in the appropriate fluorescence conjugated secondary antibody cocktail and DAPI for nucleus visualization.
Then, use mounting medium suitable for fluorescence microscopy to mount coverslips onto the slides. For confocal microscopy imaging of the labeled tissue sections, use sections stained with the IgG control and primary antibodies to set the detector's sensitivity, laser power, and offset for each individual channel. Then, set the upper and lower positions for the z-stack acquisition and determine the thickness and number of stacks to be imaged.
When all of the images have been acquired, open the images in ImageJ and open the channel tools panel and pseudo color the different staining channels. Merge the color channels. All of the channels should become visible on the same image.
Turn individual color channels on and off within the channel tools panel, and right-click on the counting icon to select the multi-point tool. Double click on the counting icon to open the point tool panel and select the type and size of the items used to count the cells. Check show all and turn on the DAPI channel only in the channel tools panel.
Then, select counter channel zero and click on individual nuclei to be tagged, scrolling through the z-stacks to count all of the nuclei stained within the region of interest. The number of quantified single-cell events will be indicated below in the point tool panel. When all of the nuclei have been tagged, turn on another staining channel and select counter channel one to tag the cells in which there is a colocalization between DAPI and the antibody staining.
For cytoplasmic staining, select cells with staining surrounding the nucleus throughout the entire depth of the nucleus, checking multiple z-stacks as necessary. The results can then be expressed as the number of cells to the total number of DAPI positive cells, or the number of cells per area within the region of interest. Immunofluorescent staining with antibodies targeting phenotypic markers, as demonstrated, allows images of each individual marker staining and differential interference contrast to be acquired for the delineation of regions of interest for single cell counting.
Single cell counting to determine the abundance of different smooth muscle cell-derived populations can then be performed using ImageJ. For example, single cell counting analysis in the fibrous cap region of these representative cross sections revealed remarkable differences in the cellular composition of the fibrous cap area between mice treated with anti IL-1 beta antibody and mice treated with an isotype matched IgG control. Indeed, the inhibition of IL-1 beta was associated with a decrease in YFP-positive smooth muscle cells, and an increase in galectin-3-positive cells.
Further, a decrease in the number of YFP-positive smooth muscle aortic alpha actin-positive smooth muscle cells was observed. Whereas the relative number of smooth muscle cell-derived YFP-positive galectin-3-positive macrophages was significantly increased at both brachiocephalic artery locations. Notably, the inhibition of IL-1 beta did not impact the overall number of osteochondrogenic cells within the lesion, or the proportion of smooth muscle cell-derived or macrophage-derived chondrogenic cells.
This protocol has been designed to enhance the rigor and reproducibility in atherosclerotic lesion analysis by standardizing key steps such as tissue collection, processing and sectioning, as well as single cell counting. This protocol can be used to analyze additional vascular beds prone to presenting atherosclerotic lesions including the aorta and the aortic root.
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