The overall goal of this experiment is to determine the relative tumor burden of intra-peritoneal ovarian cancer metastasis in a syngeneic orthotopic xenograft murine model. This novel method to image and quantify relative tumor burden can help us answer key questions in regulation of ovarian cancer metastasis. The fundamental advantage of this technique is that by un-mixing of fluorescent spectra, tissue auto fluorescence is removed from the images, enabling the measurement of standardized, relative tumor burden.

This technique lends insights in the human ovarian cancer metastasis with implications towards monitoring the efficacy of therapeutics. Visual demonstration of this method is critical because we are coupling a robust imaging technique with an analytical procedure to extract quantitative data from the images. Demonstrating the procedure today will be Yueying Liu, the laboratory Program Manager for the Stack Lab.

After 10 weeks of tumor cell growth, place the body of each anesthetized mouse into a small animal optical imaging system, one at a time. Scan all mice in the cohort at each time point. To observe fluorescently labeled tumor cells, execute a multi-spectral acquisition to collect a total of five images.

Select a standard exposure of 15 seconds, two by two binning, a field of view of 16 centimeters, and an F stop of 1.1. Next, acquire a reflectance image of the entire animal using white light from an open excitation filter, an open emission filter, a standard exposure of 0.2 seconds with two by two binning, and a field of view of 16 centimeters, with an F stop of 2.8. Following euthanasia, use pointed dissection scissors to cut the skin anteriorly along the ventral mid-line of the mouse, from the top of the thighs to the bottom of the rib cage.

Gently separate the skin from the peritoneal tissue, being sure not to puncture the peritoneal wall. Then cut laterally, both along the bottom of the rib cage, and the top of the thighs, to create two flaps of skin. Place the mouse on its ventral side in the scanner, so that the peritoneal cavity is facing downward with the skin flaps pulled open.

Scan the mouse with its organs in situ, using the same parameters that were previously used for the live mouse. Now, continue dissecting the mouse by extracting the liver, the omentum, the pancreas, the stomach, the diaphragm, and the small and large intestines using standard techniques. Also, remove the left and right peritoneum, the ovaries, the kidneys, the mesentery, and lastly, the fat pad.

Observe, enumerate, and record each of the visible tumors on the organs. Use a caliper to measure the diameter of each tumor. Designate tumors that are less than two millimeters in diameter as small, in between two and five millimeters as medium, and greater than five millimeters as large.

Next, place the organs on the organ scanning template, found in Figure Four of the accompanying text protocol, and use it to organize the location of each organ during the next scan. Use PBS to keep the organs moist. Scan each sheet of organs using the small animal optical imaging system and the parameters that were previously used.

After scanning, place the organs in 10%formaldehyde, and process them for histology using standard techniques. In order to reduce the amount of auto fluorescence in each multi-spectral scan, first load the red fluorescent protein in vivo spectral file followed by the auto floor in vivo red file in the unmixed images window, and select un-mix. Then export the dot bip files in a 16 bit unscaled dot tif format, and load them into Image J, by going to file, open, and then selecting the correct 16 bit dot tif file.

Next, go to the image menu, and select adjust, then brightness contrast, and set the minimum displayed value to zero, and the maximum displayed value to plus 35353. Then under image menu again, go to look up tables, and select red hot. Various animal models will require different brightness levels, but it is important to keep the brightness consistent throughout a given study.

Using the free hand selection tool, draw a free form region of interest selection around each organ. The free form region of interest should be drawn close to the organ borders. Then, type control and M to use the measure tool to calculate the surface area of each organ.

Record this data in a spread sheet. With the region of interest that was drawn around each organ, right click and select duplicate to include only the organ. Then, under image, go to adjust, then threshold, and set the lower threshold level to plus 1, 200 and the upper threshold level to plus 1, 700.

Also, check the dark background, and then select okay. This will select only the most brightly fluorescing regions. The images may require manual manipulation of the threshold sliders in Image J so that the previously brightest areas are selected for the analysis step, as shown above.

Different disease models will require different threshold constraints, but it is important to keep the threshold levels consistent throughout a given study. Under analyze, select analyze particles, and change the size of the pixel squared to 10 to infinity. Choose to show outlines, and check to select only display results, and then click okay.

This will measure both the areas, and raw integrated densities of the bright regions deemed tumors. Record the values of the surface area and the raw integrated density of each tumor selection together in the same spread sheet containing the organ surface areas. Then, under analyze, select tools, and go to calibration bar.

Add a calibration scale bar to the images of the organs. Montages can contain greater or fewer numbers of organs as dictated by a given study. Next, under file, go to save as, and save the montage as both a dot tif and a dot jpeg.

Then, open the dot jpeg file of the organs, and go to the insert menu to add a text box to each image. Add organ labels by creating a text box below each of the three rows of organs. Open each of the 16 bit dot tif files of the full body scans into Image J in order of mouse number.

Then, go to the image menu, and select adjust, and then brightness contrast. Set the minimum displayed value to zero, and the maximum displayed value to plus 35353. Next, go back to the image menu, and to look up tables, and then select red hot.

Once this is complete, save the montage as both a dot tif and a dot jpeg file. The ID8 murine ovarian cancer cell line that is injected into the mice 10 weeks prior to imaging fluoresces in the red channel, and continues to do so throughout growth. Live imaging using a small animal optical imaging system is able to view the expanding ID8 cells in the mouse and provides a more accurate approach than simply weighing the mouse in order to asses tumor burden.

Following euthanasia and removal of the ventral skin layer, the intact organs can be viewed in more detail. However, individual organs placed on the organ scanning template display the tumor burden clearly in each organ. The tumor burden in the omentum and pancreas, ovaries and mesentery of the mouse shown on the left is much greater than those same organs in the mouse shown on the right.

The observation of differential tumor burden is confirmed quantitatively both in terms of tumor service area and tumor signal intensity. Once mastered, this technique can be used to image up to 18 living mice per hour. Dissection and ex vivo imaging of individual organs takes roughly 20 minutes per specimen.

Following this procedure, other methods like quantitative magnetic resonance can be performed in order to address hypotheses such as the impact of obesity on ovarian cancer metastasis, as shown in our 2015 cancer research article.