Chronische Imaging of Mouse Visual Cortex Mit einem ausgedünnten Schädel Vorbereitung

Dendritic spines are membrane protrusions from a neuron’s dendrite that receive synaptic input. Lengthening, shrinking appearance or disappearance of spines occurs in response to changes in neural network function that result from specific patterns of neural activity. This phenomenon is known as cortical plasticity to observe morphological changes in dendritic spines.

In live animals, an imaging plate is affixed to a mouse’s skull. An area of the skull is surgically thinned and images are required at low and high magnification. The areas are then re-image at later time points.

The results obtained show that dendritic spine morphology can be clearly visualized using a thin skull preparation on multiple time points. And that spine morphology is highly plastic, showing changes in structure and also the appearance and disappearance of dendritic spines. Hi, I’m Emily Kelly in the laboratory of Anaya Maka in the Department of Neurobiology and Anatomy at the University of Rochester.

Today we’ll show you procedure for chronic imaging of the mouse vial cortex following a fins called preparation. We use this procedure in our laboratory to study dendritic spine turnover. So let’s get started.

Begin the procedure by sterilizing the tools for aseptic surgery. Then sterilize the workspace using 70%ethanol and cover the operating surface with clean dressing. Monitor the depth of anesthesia in a fentanyl cocktail anesthetized mouse by testing its response to a toe pinch the mouse shown here is A-G-F-P-M transgenic mouse in which GFP is expressed.

In layer five parametal cells that project dendritic processes into superficial layers. To maintain a body temperature of 37.5 degrees Celsius, place the animal on a heating blanket and monitor it with an attached rectal thermometer. Then apply ophthalmic ointment to the eyes to prevent them from drying out using scissors.

Remove the hair from the back of the mouse’s head from behind the ears to the eyes. Then clean the top of the animal’s head with an application of 70%ethanol, followed by Betadine scrub and Betadine solution. Make an LCU incision behind the ears and up along either the right or left side of the midline, depending on which hemisphere will be imaged to the eyes, fold the skin over and away from the surgery site.

Do not remove the skin since it will be sutured later using fine forceps. Gently pull the skin up and away from the area of interest. Using clean forceps, gently scrape away the periosteum from the exposed area of the skull.

Apply a small amount of 10%fair chloride to the skull to completely dry the periosteum membrane and ensure that it has been removed completely. It is important that the skull is completely dry. To ensure proper adhering of the glue, gently scrape away dried periosteum membrane with a microsurgical blade.

Next, apply a thin layer of Sano acrylic glue around the imaging window of the stainless steel head plate and affix the plate to the skull. With the wooden end of a cotton swab stick, apply glue to the inside seams of the imaging window, making sure to fill all the gaps between the imaging plate and the curvature of the skull. Place a drop of the glue accelerator into the window.

Then quickly wipe away the excess. Allow the glue to dry once the glue has completely dried. Using a dental drill affixed with a 0.7 millimeter stainless steel burr, begin to thin the skull in the imaging window.

Alternate drilling with occasional application of 0.9%sterile saline. To avoid generating too much heat on the skull. Thin a four by four millimeter window on the skull using small strokes across the skull surface.

With the dental drill. Test the thinness of the skull with blunt end forceps. The skull surface will indent slightly with gentle pressure.

The optimal skull thickness for imaging is between 10 and 30 microns. Once the skull is thinned, clean the window of all debris and place saline on the skull. Photograph the imaging window with a digital camera mounted to a dissecting scope.

The brain vasculature in this photograph will aid in locating the position to place the imaging plate for subsequent imaging sessions. Transport the animal to the two photon rig. The animal should remain on the heating blanket and body temperature should be monitored continuously throughout the imaging session.

A two photon microscope with a mi tai laser. A 10 watt solid state pump for maximum power and a modified Olympus flow view confocal unit are used. The system is optimized for deep tissue imaging and external detectors are installed as close to the objective as possible to allow for maximum detection of fluorescence from the brain, add a drop of saline to the imaging window using low digital zoom.

Identify an area containing brightly labeled neurons using a digital camera fixed to the microscope. Ips, take a photograph of the imaging area. This is necessary to return to the same imaging location at a subsequent time point.

Obtain a low magnification stack through the entire visible extent of the brain showing the extent of dendritic ramification. This will be used to locate the imaging area at subsequent time points as both blood vessels and dendrites. Maintain their structure in young and adult animals using imaging software.

Increase the digital magnification eightfold, adjust the XY coordinates if necessary, and collect a high magnification stack, which will show detailed dendritic morphology including the locations and structure of dendritic spines. Take detailed notes regarding each stack collection, including pan coordinates, the range of collection, the Z step size, and the number of steps in the stack. These notes will be used when returning to this dendri or axon at a subsequent time point.

Following imaging, remove the head plate by gently separating it from the skull surface. Using forceps. Remove any residual glue and wipe the skull with 0.9%sterile saline.

Suture the skin over the skull together using number six suture thread. Wipe the area with 70%ethanol following the imaging procedure. Place the animal in a clean cage under a heat lamp until it wakes up and is mobile.

Then return the animal to its original cage to image the animal again at a later time point anywhere from two days to many months later, remove the stitches and reopen the skin on the head. Using aseptic methods, use the digital image of the brain vasculature taken during the first surgery to locate the original imaging location. Then re-fix the head plate as before.

Use a microsurgical blade to gently thin away any bone that might have regrown between imaging time points if necessary. Thin with a dental drill, clear the imaging window of debris and apply a drop of 0.9%sterile saline. Carry the animal to a two photon microscope and locate the original imaging area using the digital fluorescent image taken during the previous session.

Start the flow view program. Open the original low magnification image, affix a transparency sheet to the computer screen and sketch the major blood vessel pattern using a transparency pen. Next, open the live image and realign the blood vasculature to the sketched pattern on the transparency film.

Remove the transparency to to re-image the eight x zoomed in structures. Open up the desired collected stack from the previous imaging session. Sketch the structure onto a transparency film affixed to the computer screen.

Zoom in eight x from the live image and align the image with the transparency. Depending on the accuracy of aligning the low magnification location, it may be necessary to also adjust the angle of the image slightly set the pan coordinates to the first day’s Imaging parameters, including step size and number of images collect the Zack re-imaging can be performed twice. The number of times the scalp is opened should be limited to avoid disrupting the integrity of the skull, which will result in poor imaging resolution in GFPM expressing mice.

A subset of layer five parametal cells and corresponding dendritic processes were visualized using in vivo two photon laser scanning microscopy here, GFP labeled visual cortex dendrites are shown. This image is a projection of a single image obtained every five microns from the PIA to a final depth of 175 microns. The boxed region is shown at higher magnification here on day zero, and again on day four here, higher magnification of a dendritic branch image from day zero and day four compared.

These images demonstrate stable spines indicated by the white arrow heads lost retracting spines, which are marked by red arrowhead and new spines indicated by the green arrowhead. We’ve just shown you how to perform chronic two photon imaging in mouse visual cortex using a thin skull preparation. When doing this procedure, it’s important to remember to take special care during the thinning process to avoid damage to the skull and underlying dura as this will result in poor imaging resolution and take detailed notes.

So that’s it. Thanks for watching and good luck with your experiments.