Functional Calcium Imaging in Developing Cortical Networks

The overall goal of this movie is to demonstrate how to measure network activity from developing cortical brain slices using calcium sensitive fluorescent indicator. DY brain slices are made from the developing enteral cortex of a young mouse. Both neurons and astrocytes are loaded with calcium dependent markers.

Changes in the fluorescence of calcium dependent dyes reflect changes in cellular activity. Non neuronal cells, including astrocytes, microglia, and endothelial cells can be stained using specific marker dyes. Cortical network activity is recorded using time-lapse Multiphoton imaging cells are detected within the network by making a 3D stack of images through the region of interest.

Changes in cellular fluorescence across multiple cells can then be analyzed to read out the activity of the developing cortical network. A hallmark pattern of activity in the developing nervous system is spontaneous synchronized network activity. Synchronized activity has been observed in many different developing neuronal regions, including intact spinal cord cortex and dissociated neuronal culture preparations during periods of spontaneous activity neurons, depolarized to fire, single spikes or bursts of action potentials activating many iron channels, including voltage gated calcium channels leading to calcium influx.

Highly synchronized activity has been measured from local networks using field electrodes or multi electrode arrays. These enable high temporal sampling rates, but give lower spatial resolution due to the integrated readout of cell activity. Single cell resolution of neural activity is possible using patch clamp electrophysiology of single neurons to measure firing rates.

However, the ability to measure from a network is limited to the number of neurons patched simultaneously, and typically is only one or two neurons. The use of calcium dependent fluorescent indicator dyes has enabled the measurement of synchronized activity across a network of cells. This technique gives both high spatial resolution and sufficient temporal sampling to record spontaneous activity of the developing network.

Fluorescent calcium sensitive indicators such as URA 2:00 AM ester contain carboxylic acid groups which are able to bind calcium mines. These fluorescent dyes are activated by specific light wavelengths either using one photon or two photon microscopy. The number of photons emitted from the dye is transiently altered upon binding of calcium.

This change in photon number or fluorescence is reported as the delta FF signal and corresponds to a change in the calcium level within the neuron. The measurement of changes in calcium sensitive indicators is a useful readout measure of network activity patterns in developing cells. Hello, I'm Rianna Morty, And my name is Julie Abbots.

We are from the Department of Interpretive neurophysiology and the university in Amsterdam. We have been using calcium imaging to study functional activity in developing tissue in the mouse cortex using calcium sensitive indicators at multifocal imaging. The aim of this short movie is to demonstrate how these methods can be used to measure both spontaneous and evoke activity patterns in developing networks in the nervous System.

Remove the brain Rapidly from the skull of a young mouse to reduce damage to the neural circuitry. Dissect out the brain in ice cold slice solution. The slice solution contains coline chloride instead of the sodium commonly used in A TSF for neuronal recordings.

In these experiments, we are making brain slices from enteral cortex of the developing mouse brain. Place a piece of filter paper on top of a petro dish and wet with a few milliliters of A TSF. Transfer the brain onto the piece of filter paper.

Bisect the hemispheres using a single edge razor blade. Separate the two hemispheres. Place one back into the icy slice solution with the remaining hemisphere.

Remove the cerebellum with the razor blade. Flip one hemisphere onto its midline, and make a cut of approximately one millimeter from the dorsal surface with a slight angle of the blade towards the rostral end. Turn the brain over onto.

Its recently cut surface using a cotton bird and metal spatula. Transfer the brain to the metal cutting block. Gently push the brain off the spatula with a cotton bird onto the glued surface.

300 Micrometer slices of brain are cut for young tissue cutting speeds and blade frequency are typically slower than those used for more mature tissue. Once cut, transfer each slice using a glass pipette slices are transferred into the holding chamber, which contains oxygenated A TSF at room temperature. The A CSF contains a higher ratio of magnesium to calcium than the A CSF solution.

Used for recording slices are left for one hour to recover. To load the cells with a calcium dependent indicator or cell specific marker slices need to be transferred to a chamber for the staining procedure. Although commercial chambers are available, one can easily be assembled from standard lab equipment for very little cost.

To make a staining chamber, you'll need the following basic lab equipment, two plastic petri dishes, one 10 milliliter syringe, a section of silica tubing, a cell culture insert with a semipermeable membrane super glue, three small tubing connectors, and a fine borne needle. Take the large Petri dish and make a small hole with a heated rod through the sidewall. Take the small Petri dish and make a hole of similar sized diameter large enough for the silicon tubing to pass through.

Pass a section of silicon tubing through the hole and form a loop inside the small dish. Use super glue to seal the open end of the tubing. Then stick the remainder of the tubing to the inner wall of the small petri dish.

Place the small petri dish into the center of the large one and glue in place. Take the remaining end of the silicon tubing and pass it through the small hole in the sidewall of the Petri dish. Take a fine needle and make small holes in the silicon tubing inside the petri dish.

Make the holes at evenly spaced intervals for an even gasper fusion. Take a cell culture well with a semipermeable membrane using a heated scalpel. Cut away the top one centimeters of the well to leave a shallow dish to hold the slices during incubation.

Place the shallow dish in the center of the small dish surrounded by the porous silicon tubing. Make a hole approximately half to one centimeter diameter in the lid of the large dish. Take a 10 milliliter plastic syringe using a heated scalpel.

Make an angled cut to remove the end of the syringe. Apply super glue to the cut surface of the syringe tube. Stick this directly over the hole on the large Petri dish lid.

Attach a tubing connector to the end of the syringe tip. Place the shallow dish in the center of the small Petri dish ensuring that the porous gas tubing is lying around the dish. This tube is then connected to a carbogen gas supply to oxygenate the slices.

During incubation, replace the lid of the large dish, which is also connected directly to the gas supply via the syringe tip. This will bring a supply of car and gas over the slices during the incubation to avoid bleaching of the dye. All procedures are carried out with as little light as possible, and both the dye and slices are kept in the dark.

Fill the staining chamber with approximately one and a half milliliters of A TSF from the slice holder. Place the interface chamber inside and fill with another milliliter of A TSF. It's important to maintain a good oxygen supply to keep the tissue healthy and for good loading of the slice.

Staining occurs at around 35 degrees C to facilitate uptake of the dye into the neurons while the staining chamber is heating up. Prepare the indicator dye for fira 2:00 AM Ester. Add nine microliters of DMSO with one microliter of onic acid to a 50 microgram vial.

DMSO and onic acid. Act as perme agents to enable the dye to be taken up through the lipid membrane. Vortex the vial for 15 minutes to ensure that the dye is completely dissolved.

For staining, transfer each slice into the interface. Insert in the staining chamber. Pet the dye directly onto the region of interest above the slices, close the lid of the staining chamber.

Leave the slices incubated in the dark for 20 to 40 minutes depending on the age of the maus used. After incubation, transfer the slices back into the slice holder to wash off any remaining excess dye. The staining protocols for calcium imaging can also be adapted for older tissue.

This requires an additional step of pre incubation. Transfer the old slices into a shallow pre incubation dish filled with three milliliters of a CSF and eight microliters of crema four solution at 0.5%Heat to 35 degrees C for three minutes. Then transfer the slices into the interface, insert in the staining chamber, and follow the normal staining procedures.

It is also possible to stain these slices for non neuronal cells, namely astrocytes, microglia, and endothelial cells. Sulfur rumine 1 0 1 can be used to stain astrocytes for sulfur domine. Make a solution of 10 micromolar pipet the purple dye over the region of interest in the slices.

Leave to incubate for a period of 15 minutes. Transfer the slices back to the holding chamber to remove any excess dye. The FSE conjugate of tomato lectin can stain for microglia and endothelial cells.

For tomato lectin, make a solution of 20 micrograms per milliliter. Pet the dye over the region of interest in the slices. Leave the slices to incubate for a period of 45 minutes.

Then transfer the slices back once more to the holding chamber. During imaging slices need to be stable under the microscope. Usually a metal harp is placed to hold down the tissue, however it can unevenly distort the surface of the slice giving only a limited field of view in focus for imaging to avoid.

This slices are stuck to the recording chamber using POLYETHALINE or PEI solution. PEI is a polymer used to enhance the attachment of cells to a surface at least one hour. Before placing the slices in the recording chambers, fill these chambers with a few milliliters of PEI solution to cover the floor of the chamber.

Firstly, rinse away PEI from the chamber with distilled water, followed by a CSF solution trans. Transfer a slice into the recording chamber and remove excess A CSF solution. Position the slice in the middle of the chamber using a fine paintbrush.

Remove all further A CSF using small pieces of absorbent filter paper. Take care to ensure that the slice has no more A CSF around its edges. Finally, pet half a milliliter of A CSF gently onto the slice without loosening it from the chamber.

Place the chambers into a large oxygenated interface container and leave for one more hour in the dark. Changes in calcium sensitive indicator dyes can be recorded with either one or two photon microscopy. We use a titanium sapphire laser coupled to an Olympus microscope to enable two photon imaging in our neuronal networks.

In addition, we make use of a L Vision biotech trim scope system with an EM CCD hammer MATSU camera for increased frame scanning rates. Position the slice chamber under the microscope with a stable flow of oxygenated A-T-S-F-A-T-S-F containing 1.6 millimolar calcium and 1.5 millimolar. Magnesium is heated to approximately 30 degrees C in the setup using white light.

Locate the region of interest in the slice and focus on the surface. Turn off the lights and close the recording cabinet door. To reduce background light levels.

Many different commercial or open software packages are available to record and analyze images In our lab, we use inspector software for acquisition from Lavis Biotech. To begin imaging, select the CCD camera mode for detection. Set the wavelength relevant for your indicator.

For fira 2:00 AM Ester, we set the excitation to 820 nanometers in the trim scope software. Select the 64 B scanning mode. Choose the optimal size field of view and pixel resolution for your region of interest.

Select an appropriate frame rate and pixel spinning if required for your image sampling. Open the laser shutter ready for continuous imaging. Using continuous imaging mode.

Adjust the gain and laser intensity and focus on the neuronal network you wish to image. Stop the scanning. Select the time-lapse settings for frame rate and sequence duration.

Following time-lapse imaging, make a Z stack of images marking 20 microns above and 20 microns below in one micron steps. This Z stack is used for cell detection during analysis. Export the recorded files as a stack of TIFF images.

To conclude, we have shown the use of calcium indicators to measure synchronous spontaneous network activity in the developing cortex. More details on the methodology and reagents can be found in the data sheets that accompany this film to enable you to set up the technique in your own lab.