Quantifying Synapses: an Immunocytochemistry-based Assay to Quantify Synapse Number

One of the most important goals in neuroscience is to understand the molecular cues that instruct early stages of synapse formation. As such, it has become imperative to develop objective approaches to quantify changes in synaptic connectivity. To quantify synapse number in a variety of experimental preparations, cultured neurons or cryo section brain tissue preparations are stained with selective pre and post synaptic markers to label points of synaptic contact between neurons.

The cells or tissue are then imaged on a fluorescent microscope and eight bit images required for each of the channels corresponding to the two immunohistochemical markers. The images are analyzed using image J 1.29 with puncta analyzer. To determine the extent of colocalization between pre and posts synaptic markers, points of colocalization can then be used to determine the number of synapses present in the preparation.

The main advantage of the technique described here over existing methods like electron microscopy, is that the synapse assay allows for a much higher throughput analysis and quantification of synapse number. The main implications of this technique extend towards the diagnosis and therapy of clinical disorders such as Alzheimer's disease, epilepsy, or autism in which abnormalities and synaptic connectivity have been implicated as a basis for the underlying pathology. This technique can be utilized to identify forms of intervention that can reduce or ameliorate the deficiencies we see in affected neuron's ability to form synapsis with each other.

Though this method is useful for studying brain development, it can also be applied to other model organisms or organ systems in which you're interested in studying or quantifying the number of cell to cell contacts for which you have the ability to selectively label each opposing face of every contact site. Prepare neuronal cultures by plating rat retinal ganglion cells purified from rat retina, harvested from P five to seven animals on glass cover slips coated with poly de lycine. In a 24 well plate, the cells are allowed to grow at 37 degrees Celsius for three to five days prior to transfection.

Following transfection incubate the cells for an additional seven to eight days to allow synapses to develop when the plated cells have sufficiently matured. Remove the culture media from the wells and add 500 microliters of prewarm, 4%para formaldehyde to each well fixed for seven minutes. At room temperature following fixation, rinse the cells three times with phosphate buffered saline.

It is important to ensure that the cells are never allowed to dry in the wells. Once a buffer is removed from a well, it should promptly be replaced by the next buffer. Prepare blocking buffer containing 50%normal goat serum and 0.2%Triton X 100.

After removing PBS from each, well add 200 microliters of the blocking buffer to each well and block for 30 minutes. At room temperature, remove the blocking buffer and rinse three times. With PBS, prepare a primary antibody dilution in 90%antibody buffer, 10%NGS solution containing the pre and post synaptic antibody pair of choice.

Here a rabbit antis synapsin, a pre-synaptic marker and a mouse anti homer. A post-synaptic marker are used Centrifuge, the primary antibody dilution for five minutes at maximum speed in a benchtop centrifuge to remove any precipitated antibody, then add 200 microliters of primary antibody solution to each. Well place the plate in a humidified container to prevent drying of the primary solution.

Incubate overnight at four degrees Celsius following the incubation with primary antibody. Rinse each well three times. With PBS, add 200 microliters of the appropriate secondary antibodies, diluted one to 1000 in antibody buffer containing 10%NGS prepared.

As before, place the dish in the humidified chamber and incubate for two hours at room temperature in the dark following the incubation with secondaries. Rinse each well three to four times with PBS. Place a drop of vector shield mounting medium with DPI on a glass slide.

Then invert the cover glass with the cells onto the drop to mount. Finally, apply clear nail polish around the edges of the cover slip. Take care to avoid nudging or moving the cover slips during application.

Since this will share the cells, allow the slides to dry for at least 30 minutes in a dry, dark place. Here, imaging is performed using a zes axio imager fluorescence microscope equipped with appropriate filter sets and a 63 times oil immersion objective. Place a slide containing non transfected cells on the microscope stage.

Using the DPI filter set to visualize cell nuclei, select cells that are at least two cell diameters away from their nearest neighbors. Use of DPI for this step will aid in avoiding bias and ensure randomness of cell selection. For each selected cell, obtain eight bit images using the appropriate filter sets in this case, GFP and Texas Red.

The superimposed pseudo colored image should show pre and post-synaptic markers in red and green respectively. Next place a slide containing transfected cells on the microscope stage. Here the cells have been transfected with a TD tomato expression vector.

Use this marker to select cells as before. For each selected cell, obtain eight pit images using the appropriate filter sets in this case, GFP, Texas Red and Sci five. The superimposed pseudo colored image should have the post-synaptic markers in red, green, and blue.

Here, the puncture analyzer program is used for quantification of co localized synaptic puncture. The Puncta analyzer plugin was written by Barry Walk and is available upon request in image J 1.26. Open one of the collected images.

Use the circular selection tool to select a region approximately one cell diameter radially around the SOR of interest. With the region of interest selected, go to the plugins menu and select puncture analyzer in the analysis options window that appears Select Red Channel Green Channel. The first subtract background and set results file.

Click okay. Then define a location in which to save the results. These results can be exported to Excel for further analysis in the window that appears.

Next, make sure a rolling ball radius of 50 is selected and uncheck the white background option. This modification is not required but is often preferred by users of the application. For ease of visualization, click okay.

A new window will appear alongside a mask corresponding with the red channel image. Adjust the threshold until the red mask corresponds as well as possible with as many discreet individual puncture. Without introducing too much noise, click done.

Set the minimum punctu size to four pixels. Do not modify anything else. Click okay.

Repeat the previous step this time for the green channel. After repeating this with the green channel, the plugin will provide quantification corresponding to puncta in each channel separately and a co localized puncta between the two channels. The synapse assay shown here can be applied to cryo sections from brain and to any other nervous system tissue such as spinal cord or retina.

Provided that there is a suitable pre and post-synaptic marker pair. Begin by harvesting brain tissue from a mouse that has been euthanized by exsanguination and profusion with PBS. Place the whole brain in 4%paraldehyde in PBS at four degrees Celsius overnight the next day.

Rinse the brain three times with PBS. Cryo protect the brain by placing it into 30%sucrose. In PBS.

The tissue will initially float keep at four degrees Celsius until the tissue sinks to the bottom. At that stage, the cryo protection is complete. Next in a two to one solution of 20%sucrose to OCT and PBS embed brainin at the desired orientation for sectioning.

For example, sagittal or coronal. Next place a flat metal surface on top of a bucket of dry ice. Place the embedded brain on it to freeze once frozen, transfer the brain to a freezer bag and place them at minus 80 degrees Celsius for up to a year before sectioning.

Using a cryostat cryo section the tissue into 12 to 16 micron sections and mount on glass slides. Next, dry the slides by placing them into a 37 degrees Celsius oven. Then rinse them three times with PBS to remove residual OCT.

A paraffin pen is used to create a hydrophobic barrier around the slide and blocking buffer is added to the slide prevented from flowing off the slide by the paraffin border. Place the slides in 20%normal goat serum in PBS for one hour at room temperature to block next place. The sections in primary antibodies diluted in PBS with 0.3%triton and 10%normal goat serum.

Here, rabbit antit, PSD 95 is used to label glutamatergic, post-synaptic compartments and Guinea pig anti v glu two is used to label glutamatergic pre-synaptic terminals. Place the slides at four degrees Celsius and incubate for 36 to 60 hours following the incubation. Wash the primary antibody off by immersing the slides three times in PBS for 15 minutes each.

After this step, make sure to protect the slides from direct light. Apply secondary antibodies diluted one to 200 in PBS with 0.3%triton and 10%normal goat serum. Here, goat anti Guinea pig.

Alexa 4 8 8 and goat anti rabbit. Alexa 5 9 4 are used. Incubate for two hours at room temperature in the dark.

Wash the slides by immersing them four times in PBS for 15 minutes each to mount. Add small drops of vector shield mounting medium with dappy and cover the slides with cover slips. Apply nail polish to inhibit movements of the cover slips.

Imaging of tissue section should be performed using a confocal microscope with appropriate lasers using a 63 times oil immersion objective for each section image, serial optical sections at 0.33 micron intervals over a total depth of five microns for a total of 15 optical sections. At least three sections from each animal must be imaged and at least three animals must be included from a given experimental condition. Generate maximum intensity projections from groups of three consecutive sections, yielding five projections representing one micron of depth each.

These are quantified using Image J as described for RG Cs with a modified ROI to determine the number of synapses formed in the absence of astrocytes. Immunochemistry was performed on dissociated rgc treated with basal growth media using antibodies against bassoon shown in red and homer shown in green as expected. Very few points of synaptic colocalization are observed to determine the effect of astrocyte secreted molecules on the number of synapses formed in vitro dissociated RG Cs were treated with mouse astrocyte conditioned media every three days for a total of 12 days following the staining.

Numerous synapses were evident in this neuron, but soon is shown in red. Homer is shown in green dissociated RDCs transfected with cytoplasmic Lee expressed TD tomato were treated with only basal growth. Medium synapses are not seen.

TD tomato is seen in blue and synapse in is seen in red. There is very little signal seen for the post-synaptic marker homer shown in green. When the same cells are chronically treated with rat astrocyte conditioned medium, many synapses are seen.

TD tomato is in blue, synapsin in red, and homer in green here. A purified RGC is shown that has been treated with astrocyte conditioned medium and has been stained for pre synapse in red and post-synaptic homer green markers threshold masks generated using puncture analyzer correspond to the two channels present. Upon identifying points of colocalization Puncta analyzer outputs a graphical representation of these points.

A numerical output is provided by puncta analyzer that indicates the number of discrete puncta identified in both channels and for points of colocalization. The data produced by analysis of this neuron illustrates the ability of astrocytes secreted molecules to promote the formation of synapses shown here as a synapse assay in which colocalization of pre and post-synaptic immunohistochemical labeling was quantified to demonstrate the potentiation of astrocyte induced synapse formation by overexpression of the neuronal receptor for the important astrocyte secreted synap genic molecule thrombo spon. Shown in this figure is the average number of synapses per neuron chronically treated with basal growth media, gm, or with astrocyte conditioned media.

A-C-M-A-C-M more strongly promotes synapse formation in retinal ganglion cells over expressing the thrombo spon receptor compared to cells over expressing an empty vector. To quantify the synaptic density in mouse superior colliculus pre and post-synaptic compartments were separately labeled using compartment specific immunohistochemical markers. As shown here, there was an increase in synapse number from postnatal day seven to P 14.

This illustrates the increase in synapse number in the superior ulus over this developmental time window. Once mastered, this protocol can be completed in about two days for dissociated neuronal cultures and at about five days for brain sections from harvesting from the animal following this procedure. Other methods like electrophysiology and electron microscopy are utilized to verify the changes in synaptic number reflect changes in both functional synapses and ultra structurally normal synapses respectively.

After watching this video, you should have a good understanding of how to apply immunohistochemistry to your experimental preparation to quantify synapse number. This protocol details how to use compartment specific antibodies to selectively label pre and post-synaptic compartments. In this context, we define synapses as points of colocalization between the signals obtained from each of these markers.