A Label-Free Segmentation Approach for Intravital Imaging of Mammary Tumor Microenvironment

Intravital imaging allows us to observe physiological interactions within the tumor microenvironment. And by using this method, specific dynamic behaviors that are observed can be compartmentalized using local tissue features. This protocol uses only an ubiquitous label-free signal from collagen fibers or autofluorescent metabolites to segment the TME, thereby minimizing the amount of manipulations to the mouse.

This method is broadly applicable to any intravital system where understanding dynamic interactions relative to extracellular matrix structures or vascular structures is required. Demonstrating this procedure will be Dave Inman, senior research specialist, and Erica Hoffmann, a graduate student from the laboratory. Start preparing a mammary imaging window cover glass by soaking a 1.5 12 millimeter round cover glass in 100%ethanol for 10 minutes, then dry the cover glass under a heat lamp and secure the glass to the metal mammary imaging window frame using cyanoacrylate adhesive.

Cure the adhesive overnight. The next day, use an acetone soaked swab to clean the assembled mammary imaging window of the excess adhesive before submerging the mammary imaging window in 70%ethanol for at least 10 minutes to assist with the cleansing. After drying, store the cleaned mammary imaging window in a sterile Petri dish.

Before beginning the surgery for the window implantation, autoclave surgical tools and sanitize surfaces with 70%ethanol. Prepare a sanitized tabletop for the surgery with a warming blanket covered with a sterile field. Set the warming blanket such that the temperature measured on top of the sterile field is 40 degrees Celsius.

Use auxiliary cold lighting and magnifying glasses for the surgical procedure. Wear PPE consisting of a sterile single-use lab coat, surgical sleeves, gloves, eye protection, and face mask. Next, remove the fur of the anesthetized mouse at the fourth inguinal mammary gland with a depilatory cream and rinse the surgical site with a sterile water soaked gauze.

Then prepare the depilated surgical site for surgery by sanitizing the skin surface with three alternating Betadine and ethanol scrubs. When done, gently lift the skin over mammary gland number four using forceps. Once the skin is pulled away from the body wall, remove a one millimeter section of the dermal layer at the tip of the forceps with surgical microscissors.

Without cutting the underlying gland, create a 10 millimeter incision and then release the mammary gland from the dermal layer with the gentle movement of the forceps. Add PBS to cover the exposed gland. Use a 5-0 silk braided suture to create a purse string suture along the periphery of the opening, then insert an edge of the mammary imaging window so that the dermal layer engages into the receiving notch of the mammary imaging window.

While stretching the epithelium at the opposite end of the mammary imaging window, push the metal mammary imaging window into place such that the dermal layer fully engages the receiving notch around the entire mammary imaging window circumference. Then cinch the purse string to draw the dermal layer into the notch and tie off the layer to secure the window. Apply a topical antibiotic to the dermal layer at the mammary imaging window and continuously monitor the mouse.

After regaining sufficient consciousness to maintain sternal recumbency, house the mammary imaging window implanted mouse separately on soft bedding with an igloo placed in the cage and allow the mouse to recover for 48 hours before imaging. For the imaging, use a forced air system set to 30 degrees Celsius. Use an additional objective heater to avoid drift in Z focus and allow the system to come to equilibrium at 30 degrees Celsius for at least one hour before imaging.

Then set up a heating chamber on the microscope stage. After confirming the anesthesia with the toe pinch method, add an eye ointment to the mouse. To maintain proper hydration, inject 0.5 milliliters of PBS subcutaneously every two hours for the duration of the imaging session.

Apply a water-based gel instead of water to the objective and clean the outside of the mammary imaging window glass with a cotton applicator and glass cleaner before transferring the mouse to the pre-warmed microscope stage. After laying the mouse on the microscope stage, press the collar of the mammary imaging window into a 14 millimeter receiving hole in the stage insert to stabilize the images and fit the isoflurane hose. Then bring the imaging field into focus using the microscope oculars and Brightfield illumination, observing vasculature with blood flow.

Check the stability of the field of view. If breathing movement artifacts are present, apply gentle compression to the backside of the gland with a small foam block and a cincture-like piece of adhesive tape. After compression is applied, verify that blood flow is maintained throughout the field of view.

Once the mouse is sedated and securely positioned on the inverted microscope stage, start locating regions of interest. Using a light source directed at the mammary imaging window, use the oculars of the microscope to identify potential areas for investigation. The focus should be on seeing vasculature and blood flow.

Add and save the XY positions in the software. When the appropriate power levels are set, set up the Z stack and observe the appearance of abundant collagen fibers at 20 to 50 micrometers beneath the glass surface of the mammary imaging window. Collagen will become less prevalent as the microscope sections deeper into the tumor.

The voids in the second-harmonic generation or SHG reveal the location of tumor masses. Set the top Z slice beneath the layer of solitary cells with the first collagen fibers appear at 50 to 100 microns. Set the bottom Z slice at 250 micrometers where the fibers fade out and the poor signal dominates.

Then repeat the procedure for all XY positions saved. Once the Z stack range is set, increase the dwell time to eight microseconds and optimize the power and detector settings. Optimize the power levels needed to excite the tissue for each experiment and use powers up to 90 milliwatts at 750 nanometers or 70 milliwatts at 890 nanometers at the back aperture of the objective.

Next, start with 10-minute intervals between collection points for most intravital migration movies and adjust the time intervals according to experimental goals. If signs of phototoxicity like cell blebbing or rapidly increasing autofluorescence and excessive photobleaching are observed, reduce laser power or increase timelapse intervals as the conditions indicate. For fluorescence lifetime imaging or FLIM of NADPH, start preview scanning and adjust the laser power until the readout of the constant fraction discriminator or CFD is between 10 to the fifth and 10 to the sixth.

Exceeding CFD beyond 10 to the sixth result in photon pileup and poor overall results. Once the power level is set, set the integration time between 90 to 120 seconds and start the FLIM collection to acquire photons from the field of view for the allotted time. A typical field of view within a label-free mammary tumor demonstrated abundant collagen fibers near the window and a decrease in abundance deeper into the tumor.

The use of fluorescence lifetime or NADPH aided in the identification of the vasculature. Some dark regions in the images were tumor folds or the butting up of adjacent tumor lobes. To identify vasculature, a composite image using NDPH FLIM was validated by comparing maximum intensity projections of the intravital stack before and after tail vein injection of fluorescent dextran.

The representative analysis shows the quantification of four mice and the fields of view. To delineate the boundaries of the label-free tumor masses, NADPH autofluorescence was used. The images reported the metabolic signatures of the cells and the location of blood vessels.

The AFD autofluorescence also identified the macrophages. With the segmentation scheme, the label-free tumor was segmented into compartments for the tumor nest, stroma, and vasculature using only SHG and NADPH autofluorescence. Additionally, the stroma and collagen fibers can also be classified into local regions of the aligned fibers.

The most important thing to remember for this protocol is to make sure that the mouse is properly sedated and restrained so that the field of view remains stable for imaging.