In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons. In vivo. Calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single cell resolution.

The method consists of implanting a cranial imaging window injecting a fluorescent calcium dye that can be taken up by a large number of neurons, and finally recording the activity of neurons with time-lapse calcium imaging. Hi, I'm Payman go from the laboratory of Carlos Porter Cayo in the Department of Neurology at the David Geffen School of Medicine at UCLA. In previous videos, our lab demonstrated a procedure for performing craniotomy and to photon imaging of blood flow in mice.

Today we will show you a procedure for performing in vivo to photon calcium imaging in mice. So let's get to it Before beginning. The dye injection solution must be prepared.

First, prepare the fluorescent calcium indicator solution. We use a ethyl ester dyes or am dyes for short. These are dyes that can be taken up by cells when injected extracellularly, we typically use organ green BAFTA 1:00 AM or flu oh 4:00 AM at a concentration of one millimolar.

We first prepare a 20%solution of onic F1 27. In DMSO, we warm the solution prior to use every day until it's clear. We dissolve the calcium indicator in 20%onic F1 27 DMSO for 15 to 20 minutes to a concentration of 10 millimolar.

We then dilute the solution to one millimolar in the solution containing in millimolar one 50 and ACL 2.5 KCL 10 heaps, pH 7.4 and vortex for an additional 10 to 20 minutes. We add 100 micromolar sulfur rumine 1 0 1 to selectively label astrocytes and to visualize the pipette during the injection of the dye, we next implant a cranial window over the area to be imaged. The general approach to this surgery is described by us in a previously released job video article, but the following modifications are important for calcium dye injection.

A smaller craniotomy of two millimeters in diameter is made over the cortical area of interest, taking extreme care not to damage the underlying dura. A cover slip is secured in place over the craniotomy, but only partially covering the craniotomy, leaving a small gap between the edge of the craniotomy and the glass cover slip to allow room for a pipette to enter the cortex obliquely prior to injection. A shallow well with dental cement is made around the craniotomy.

The well for calcium imaging experiments should extend as far in the rostral direction as possible and be shallow enough to allow for the introduction of the injection Micro pipette, we filter the solution with a 0.45 micron centrifugal filter immediately before injection. The mouse is then transferred to the stage of the microscope and the head immobilized as shown in our video on blood flow imaging. Next, we pull glass micro electrode pipettes to a tip diameter yielding two to four mega ohms in resistance.

Using a pipette puller, the calcium indicator dye mix is loaded onto the glass pipette. Using a micro syringe using a micro manipulator, the micro pipette is gently lowered onto the surface of the brain using a four x objective and then more finely positioned. Under a 40 x objective, a suitable location for injection is chosen such that there are few or no overlying blood vessels.

To obscure the imaging using two photon microscopy, the tip of the micro pipette is visualized and advanced slowly into the cortex to a depth of 200 micrometers below the dura. To perform this accurately as possible, the Z focus reading of the objective is set to zero. Once the dura is visualized, then the micro pipette is visualized and the Z focus reading is taken as the micro pipette is lowered into the brain to 200 micrometers below the dura.

The D mixture is then pressure injected at 10 PSI for one minute. Using a pico spritzer, we usually deliver two to four injections of the AM calcium dye mix separated in space by approximately 200 to 300 microns. This bulk loading approach of slightly overlapping injections ensures that an area as large as 600 by 600 microns is adequately stained for calcium imaging.

Imaging begins one hour after the injection of the am dye mixture to allow for the dye to diffuse and be taken up by neurons. In vivo two photon Imaging is performed with a custom-made two photon microscope using a chameleon ty sapphire laser tuned to 800 to 880 nanometers and Cambridge technology. Galvanometer mirrors images are acquired using scan image software developed in Carl's Vida's.

Laboratory laser power is typically well below 70 milliwatts at the sample. Whole field images are acquired using a 20 x 0.95 NA Olympus objective at 1.95 to 15.63 frames per second, five 12 by 2 56 pixels to 2 56 by 64 pixels. Line scans are acquired at 500 hertz during imaging.

Animals are kept under light isof fluorine, anesthesia 0.6 to 0.9%and are also kept at 37 degrees Celsius using a Harvard apparatus. Temperature control device care is taken to keep the respiratory rate of the animals as close to 100 breaths per minute as possible. This level of anesthesia use is the lowest level that immobilizes the animal, but still permits spontaneous activity to be recorded.

So now let's have a look at a typical calcium imaging Experiment. There are two three neurons Of somatosensory barrel cortex of a postnatal to eight 10 mouse. Were stained with organ green BTA am and imaged with a 20 x 0.95 NA objective.

Images were taken every 0.512 seconds. This three minute movie shows co activation of large proportions of this stain neurons occurring one to two times Per minute. We've just shown you how to Perform in vivo calcium imaging of network activity in layer two three of the cerebral cortex.

The advantage of this method over electrophysiological recordings is that it is less invasive and allows the recording of activity in dense neuronal networks. When doing this procedure. It's important to remember to take extreme care when performing the craniotomy not to damage the dura.

Even a small amount of subdural bleeding will greatly obscure the imaging. So that's it. Thanks for watching and good luck with your Experiments.