Longitudinal Two-Photon Imaging of Dorsal Hippocampal CA1 in Live Mice

This method describes a chronic preparation that allows optical access to the hippocampus of living mice. This preparation can be used to perform longitudinal optical imaging of neuronal structural plasticity and activity-evoked cellular plasticity over a period of several weeks.

This procedure provides repeated optical access to the dorsal hippocampus of live mice to study the mechanisms of memory formation and recall while they perform learning tasks. This technique allows the use of light microscopy to study the dorsal hippocampus of live mice at micrometer level spatial resolution longitudinally over several weeks. Memory loss is a general hallmark of some brain diseases such as Alzheimer's disease.

Understanding the mechanism of memory formation and recall could ultimately help patients suffering from these diseases. This method could be applied to other animal systems with a skull such as small mammals. Monitoring vital signs during a survival surgery can be both distracting and stressful.

It might be useful to first perform the procedure on a dead animal in order to focus on the more crucial aspects of the procedure. To prepare the imaging cannula, first cut a three millimeter diameter stainless steel tube into a 1.6 millimeter long metal ring. If the edges of the ring are not blunt after cutting, file out the irregularities.

Next, use a pair of forceps to dip one side of the metal ring into a UV curing optical adhesive. Then position the metal ring at the center of a glass coverslip with the side of the ring covered by adhesive touching the coverslip. Turn on the UV curing LED driver unit and shine 365 nanometer wavelength light onto the adhesive for one minute to cure the adhesive.

Make sure all sides of the ring are equally illuminated by changing the direction of the light source. After the adhesive has hardened for at least two hours, firmly hold the cannula from the open end of the metal ring by means of a hemostat. Using a dental drill fitted with a rotating file, file off the excess glass coverslip until it is flushed with the sides of the ring.

Prepare the required instruments and anesthetize the mouse as described in the accompanying text protocol. Then remove the hair and disinfect the skin over the mouse head. Next, use scissors and forceps to make a small cut in the scalp in the position close to lambda.

Move laterally in the direction of the ears then rostrally in the direction of the eye orbits to form a triangle on the sagittal axis approximately four millimeters rostral to bregma. Apply a single drop of lidocaine to the skull. After two minutes, clear the periosteum and dry the skull using a cotton swab.

Then position the ear bars to fix the mouse's head. Using a micro drill with a 0.5 millimeter width burr, make a small hole in the frontal bone opposite the imaged hippocampus approximately 1.5 millimeters from the sagittal suture and two millimeters distal from the coronal suture. Then screw a 0.86 millimeter width stainless steel bone screw into the skull hole.

Secure it in place using adhesive cement. Let the cement dry for 30 seconds to a minute. Next, use a three millimeter diameter trephine drill to make a small hole in the parietal bone.

Position the hole approximately 1.5 millimeters distal from the sagittal suture and two millimeters distal from the lambdoid suture. Carefully remove the bone flap. Then remove the meninges using Dumont forceps.

Ablate cortical matter to reach the external capsule. Use a 19 gauge blunt needle connected to a vacuum pump and irrigate with saline to avoid dehydration of the exposed tissue and wash away any residual blood after the bleeding is resolved. Suck cortical tissue slowly about 50 to 100 microns at a time until the cortex detaches from the capsule exposing the fibers of the cingulum or the corpus callosum.

Then carefully peel the dorsal fibers until the deepest fibers, the alveus of the hippocampus are exposed. When finished, rinse the tissue with saline. Next, use thin forceps to dip the bottom cannula into saline and position it over the skull hole.

Then push the cannula into the skull until the glass coverslip is in contact with the fibers. Dry the skull and apply a quick adhesive cement over the skull to hold the cannula in place. Be sure to also apply adhesive on the rim of the cannula but be careful not to let the adhesive run into the cannula.

Once dry, use a stereotaxic arm to position a head holder plate over the cannula in contact with the skull. Prepare a mixture of dental acrylic and then apply it across the entire cranium. Cover the entire exposed skull, the screw and the open skin with dental acrylic to stabilize the preparation.

After 15 minutes, apply a removable adhesive film onto the head holder plate to prevent debris from entering the cannula. When ready to image the brain, again follow along in the accompanying text protocol for details on anesthetization. Once sufficiently sedated, use forceps to carefully remove the adhesive film from the head holder plate.

Remove the film gently to avoid damaging the preparation. Next, position the mouse under the microscope over the heating carpet and secure the head plate to the holder. Apply eye ointment to the animal's eyes.

Then clean the imaging cannula by using a syringe and a thin needle to drop deionized water into the cannula followed by removal with a vacuum pump. Use low magnification long working distance objectives to visually check the cannula for residual water, dirt, integrity and the presence of fluorescence. Then align the cannula to the optic axis by adjusting the angles of the head holder arms.

Switch to a 25X objective with a 1.0 numerical aperture and a four millimeter working distance. Then add enough deionized water to the cannula to fill it and maintain excess water on top of the cannula. Finally, use two photon excitation and image the fluorescence signals.

Shown here is a 2P image stack from a thy1-GFP transgenic mouse brain. Thy1-GFP mice express cytoplasmic enhanced GFP under the control of the Thy1 promoter in a sparse random population of pyramidal neurons. Generally, the major axis of pyramidal neurons in the dorsal hippocampal CA1 is roughly perpendicular to the XY imaging plane.

These regions are manually mapped to a low magnification three-dimensional stack showing the pattern of GFP expression in the volume below the imaging cannula. For longitudinal tracking, several brain regions within the field of view of the cannula are defined. Each region corresponds to an area of approximately 240 by 240 micrometers and contains between one and seven dendritic segments.

The imaged series here shows one micrometer z-step image stacks of CA1 pyramidal neurons and basal dendrites acquired at different time intervals for 14 days. However, longer imaging durations and intervals are possible. Although most images are of dendritic spines in the stratum oriens, it is also possible to image dendritic spines in the oblique dendrites of the stratum radiatum.

Another extension of this technique is to image activity-evoked plasticity in CA1 pyramidal neurons by injecting the dorsal CA1 hippocampal area with a viral vector. This allows for the plasticity of hundreds of CA1 pyramidal neurons in each animal to be measured shown here as a 2P image stack. It is crucial to prevent damage to the hippocampus when removing the overlying tissue.

Additionally, one should place the cannula correctly to ensure a stable preparation. The described preparation allows for the use of different kinds of optical imaging such as miniature microscopes that can be implanted on the animal's head. This allows the researcher to follow neuronal activity in freely moving animals.

Originally, the technique was used to study structural plasticity in hippocampal neurons. Today, it is implemented in studying neuronal activity also in other brain areas.