炎症真皮における撮像CD4 T細胞間質への移行

The overall goal of this imaging protocol is to visualize CD4 positive T Cell dynamics in the inflamed skin. This method can help answer key questions in the field of immunology, such as, what are the mechanisms that underlie immune cell motility, and function in inflamed tissues. The main advantage of this technique, is that it provides a minimally invasive way to study CD4 T cell dynamic behavior In Situ.

Visual demonstration of this method is critical, as the steps to prepare the ear for imaging are difficult to learn, and proper positioning of the ear is necessary to capture high quality images. I'm Alison Gaylo, a graduate student, and I will be doing the demonstration of this protocol. Begin by aspirating 300 microliters of fluorescently labeled effector T cell suspension per mouse into a syringe equipped with a 27 and a half gauge needle.

Then turn the syringe needle side up, and gently flick the shaft of the syringe, moving the plunger up and down as necessary to remove any bubbles. When the cells are ready, transfer the mice into a clean cage, and place them under a heat lamp, until the tail veins are vasodilated. Then transfer the first mouse into the restrainer, and wipe the tail with an alcohol swab.

After identifying a lateral tail vein, slowly inject 200 microliters of cells into the vein. When all of the cells have been transferred, draw 50 microliters of complete fluorescens emulsion into a 28 and a half gauge insulin syringe and press down firmly on the plunger to remove any large air bubbles. Then place a thimble on the left index finger, and carefully grasp the mouse's ear between the left thumb and thimble, with the ventral ear facing up.

Then slide the needle, bevel side up, into the dermis in the outer third of the pinna, and slightly off center, and slowly inject ten microliters of emulsion into the tissue. After injecting all of the animals, monitor the mice until they are fully recovered. Three days following the immunization, vasodilate the tail veins of the injected animals as just demonstrated.

After anesthetizing, immobilize the tail of the first mouse with a pair of forceps. Wipe the tail with an alcohol swab, then carefully slide a 30 and a half gauge needle catheter into a lateral tail vein, checking for patency by gentle pressure to the syringe plunger. When the catheter is in place, apply one to two drops of cyanoacrylate tissue adhesive to the injection site, and allow the adhesive to dry for approximately 30 seconds.

Then use scissors to carefully trim the whiskers and the hair from the back and sides of the ear. Next, use a cotton swab to moisten the inner surface of the ear with PBS. Then rotate the mouse, and flatten the injected ear onto a 24 by 50 millimeter number one point five glass cover slip, until it is flush with the glass.

Blot away the excess PBS. Then use two pairs of curved forceps to grasp an approximately 20 millimeter long piece of fabric tape lengthwise at the top corners, and place the bottom of the tape onto the cover slip at the top of the mouse's ear. Roll the tape over the rest of the ear, pushing the excess hair out of the way with the forceps as necessary, and use a dry cotton swab to gently press the tape around the ear to ensure a tight seal, taking care not to press on the ear itself.

It is essential that the ear is taped properly to the cover slip with a good seal on both sides and with minimal air bubbles, as this will affect the quality of the images that can be obtained. Now tape a fluorescent microscope imaging platform to a 37 degree Celsius heating block, and apply vacuum grease to both sides of the ear area of the platform. Rotate the mouse to place the cover slip onto the platform, taking care to align the ear in the center of the felt.

When the ear is in place, snap the isoflurane nose cone into the holder on the platform, ensuring that the cone is secure, and completely covering the mouse's nose. Then use a cotton swab to firmly, but carefully, press the cover slip onto the imaging platform, spreading the vacuum grease under the cover slip. Wrap two 20 millimeter pieces and two longer pieces of tape around the upper portion of the platform to affix the cover slip to the platform.

Next, move the mouse to the microscope stage. Secure the platform with more tape, and pipe a double layer of vacuum grease onto the cover slip around the ear. Then wrap a water filled heating blanket around the mouse, and fill the vacuum grease reservoir with 37 degrees Celsius distilled water.

For In Vivo time lapse imaging, begin by positioning the objective over the center of the ear, and lower the objective until it just contacts the surface of the water in the reservoir. Next, using an external light source, slowly lower the objective until the surface of the ear comes into focus. Then, lower the inner and outer curtains around the microscope stage, and adjust the laser wave length to 900 nanometers in the MP laser controller window, the laser power in the acquisition setting laser window, and the PMT voltages in the image acquisition control window.

In the acquisition setting size and mode windows, set the microscope to five twelve by five twelve pixel resolution, with a two microsecond per pixel dwell time, and select XY repeat to activate a live imaging mode for scanning through the tissue. Once an appropriate imaging field has been located, locate the highest cell in the dermis, and click the set zero button to set the Z position to zero. Scroll down in the Z direction to measure the extent of the cell depth.

Then set the starting and end positions in the acquisition setting microscope window, and adjust the instrument PMT voltages and laser power in the bright Z window to optimize the visualization of the cells throughout the depth of the imaging field. Check the depth and time buttons, and set a common filter to scan the image three times per line in the image acquisition control filter mode window. Adjust the Z slice depth in the image acquisition microscope window so that it takes approximately one minute to capture a complete stack as noted in the time view window.

Then, to assess the stability of the tissue, set the number of repeats to five in the acquisition setting time scan window, and click the scan button to capture a five minute time lapse image of the area. If the tissue is stable after five minutes, set the number of repeats to between 30 and 45 in the acquisition setting time scan window, and collect a 30 to 45 minute time lapse image, monitoring for any minor tissue drift as the image is collected. Save the pre-antibody image in the appropriate format, and then draw the antibody mixture into a one milliliter tuberculin syringe, taking care to remove all of the air bubbles.

Gently press the plunger so that the antibody solution forms a droplet at the end of the needle, and lift the curtains around the stage to locate the catheter. Replace the PBS syringe with the syringe of antibody, and then slowly inject the antibodies into the catheter. When the antibodies have been successfully injected, re-lower the curtains and note the time of injection.

Then immediately start collecting a new 20 to 40 minute imaging sequence, in the same location, using the same instrument settings as the pre-antibody image. Finally, save the post-antibody image file in the appropriate format, and carefully remove the mouse from the microscope stage, monitoring the animal until it is fully recovered. Imaging of the intact ear dermis by this protocol facilitates the acquisition of high resolution and time lapse images of effector T cell dynamics in the inflamed dermis.

In this movie, the green adoptively transferred effector T cells can be observed moving throughout the dermal tissue. After the administration of antibeta one and antibeta three entogrin blocking antibodies through the catheter, however, the previously motile cells arrest within the dermis. The cells exhibit a decreased average velocity after antibody administration as well as a significant decrease in their meandering index.

That is, the ration of the total displacement to the total track length. Autofluorescents from hair follicles, shadowing and autofluorescents from overlying hair, and air bubbles trapped between the ear surface and the cover slip, can all lead to imaging artifacts, which obscure the visualization of the T cells, and interfere with the automated image analysis software. In addition, using multiple sources of heat can cause tissue stability issues, leading to large oscillations during imaging that make interpreting the results difficult.

Once mastered, imaging the CD4 T cells in the dermis can be completed in two to three hours, if the procedure has been performed correctly. While attempting this procedure, it's important to remember to follow the appropriate animal care and use guidelines for working with mice. This procedure can be adapted to visualize other T cell subsets or types of immune cells in the ear, using different models of infect, or different methods of inducing inflammation.

After its development, intravital imaging paved the way for researchers in the field of immunology, to explore the dynamics of ongoing immune responses in living animals. Don't forget that working with a multi-phosphor laser can be extremely hazardous and that precautions, such as proper shielding, should always be taken while performing this procedure. After watching this video, you should have a good understanding of how to induce inflammation in a mouse ear dermis, how to use intravital imaging to capture the time lapse images of CD4 T cell activity before and after antibody administration.