Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

The goal of this procedure is to use remote control to activate olfactory bulb neurons and observe how their activation affects behavior. This is accomplished by introducing a trends gene of channel Redin two, coupled to YFP into target neurons of adults with stereotaxic injections of a lentiviral vector. Then an LED is calibrated and prepared for implantation producing a general idea of the amount of light transmitted into the brain.

The final step of the procedure is the implantation of the LED over the olfactory bulb, ensuring its proper positioning and attachment. Ultimately, neuronal activation is confirmed through CFOs, immuno staining, and the results of behavioral testing. The main advantage of this technique over existing methods like standard electrical stimulation or pharmacological perturbations, is that optogenetic stimulation of labeled neurons is highly specific regarding both type and number of neurons activated.

The implication of this technique stand forward the therapy or diagnosis of neurodegenerative disease because of the precise temporal and spatial control offer. This method can provide insight into the role of adult bone neuron in the olfactory bone network. It can be applied to other superficial regions such as the cortical or cerebral lung neurons.

Begin by retrieving prepared vector titers from the negative 80 degrees Celsius freezer. Keep them frozen before they use and discard any leftover titers. Cut the tip of pooled boro silica glass pipettes using a pair of scissors.

The goal is to make 30 to 40 micron diameter tips. Store the prepared pipettes in a dust-free box. Next, set up a micro injector to inject 50 nanoliters back filler prepared pipette with mineral oil and insert it into the micro injector plunger, which should be attached to the stereotaxic frame.

Once the viral titer has thawed, partially empty oil from the glass pipette, and then refill it with the titer. Transfer two microliters of virus onto a sterile piece of para film. Gently lower the glass pipette into the drop and fill the pipette with one to two microliters of virus.

Check that there are no air bubbles in the pipette, and then proceed with preparing the animal after anesthetizing a mouse using ketamine and xylazine diluted in sterile saline. Make sure the mouse is non-responsive to pain stimulus, such as a toe pinch before proceeding. Now, remove the hair from the scalp of the mouse and disinfect the scalp.

Place the animal in a stereotaxic frame and use ear barss and a nose bar to secure its head with a scalpel. Cut the scalp from between the eyes to between the ears. Pull the skin aside to expose the skull and anchor the skin with a pair of clamps.

Then clean the surface of the skull using sterile saline solution. Zero the stereotaxic apparatus with the tip of the glass pipette at breg ma. Then position the tip at the injection site.

Adjust the position of the nose bar according to the calculation of the stereotaxic coordinates. Using these coordinates, the bgma and the injection site are aligned to the same height. Using a high speed surgical drill, carefully drill a small hole into the skull at each injection site.

Take care not to rup your blood vessels on the surface of the brain while drilling. Now with a bent syringe needle hooked at the tip, remove the remnants of thin bone to expose the URA mater. Check that the hole is centered at the correct position.

Lower the pipette and zero the dorso ventral height. When the tip of the pipette touches the surface of the brain, then lower the pipette to the correct dorsa ventral depth and wait 30 seconds for pressure equilibrium. Now inject 50 nanoliters of virus four times for a total of 200 nanoliters of virus waiting 30 seconds between injections.

One minute after the last injection, slowly withdraw the pipette and remove the mouse from the apparatus. Clean the incision site with Betadine solution and stitch the skin. Using surgical thread, allow the mouse to recover on a warming pad before finally returning it to its cage before implanting the miniature blue LED.

For optogenetic stimulation, it must be prepared and tested. Sold are the LED pins to a female electrical connector so that the connector is positioned on the opposite side of the LED. Check that there is no electrical short circuit, and identify the positive and negative pin on the connector.

Lastly, test the LED by connecting it to an LED controller to measure or set the absolute light power of the LED mount, the LED on a micro manipulator and prepare the light meter. Drill a three millimeter hole in an opaque PVC board and affix this hole to the power meter. Now position the LED in direct contact with the photo detector of the power meter and take measurements at the peak LED light intensity, which is usually four 70 nanometers.

Use the power meter to build a standard curve of optical power versus input current. Then use the to calculate the power per square millimeter to measure the propagation of light through the brain. Obtain vibram cut blocks of fresh brain tissue of 300, 500, 1000, one thousand five hundred and two thousand five hundred micron thicknesses using a piece of skull with a craniotomy identical to the one performed in the live mice.

Place it between the LED and the detector. Now, measure the light power without tissue. Then place each block of tissue between the LED and the photo detector of the power meter and measure the change in power.

Build a curve of tissue thicknesses versus relative transmission fraction. Transmission fraction is the ratio of the power measured through the tissue compared to without the tissue. Use this data to calibrate the LED and then proceed with the implantation surgery.

Prepare the LED and connector by disinfecting them with 70%ethanol and attaching the connector to a stereotaxic holder. After placing an anesthetized mouse in the stereotaxic frame, use a nose bar to align the surface of the skull perfectly horizontal and parallel to the surface of the miniature LED. In this case, the region of the skull corresponding with the olfactory bulb is aligned.

Now with a scalpel, cut the scalp from three millimeters anterior to the eyes to three millimeters posterior to the ears. Pull the skin aside and anchor it with a pair of clamps. Scrape away the membranes at the surface of the skull using a razor blade.

Prepare to perform a rectangular craniotomy over the off effect bulbs. First, use a high-speed drill to carefully thin the skull until only a thin layer of bone remains. Second, carefully remove the remaining bone with the help of a bent syringe needle.

Carefully avoid puncturing the dura mater. Now rinse the surgical area with sterile saline and then dry the skull. After cleaning and drying the exposed skull, apply a thin base layer of cyanoacrylate adhesive to the skull for later adhesion of dental acrylic.

The addition of two small screws may be required to fix the LED. The screws should be inserted into the skull posterior to the LED and covered with both acrylics. After attaching the LED connector to the holder, lower the miniature LED above the olfactory bulbs with its main axis parallel to the rostral cordal axis.

Then secure the LED to the skull with two layers of dental acrylic. While the acrylic is still liquid, pull back the free edges of the scalp and glue them to two layers of dental acrylic. Be careful not to apply any acrylic to the connector after the acrylic has hardened.

Remove the animal from the frame and let it wake up on a warming pad. Once returned to its cage, full recovery takes at least seven days later. A thin light electrical cable is used to tether the animal to an LED controller, allowing ample ambulatory freedom, respect the polarity of the wire as inverting.

The polarity may break the LED. After the experiment, remove the LED to verify that it was functional CH opsin. Two YFP was expressed in transduced neurons approximately 48 hours after injection and remained stable for several months.

A sagittal section of a paraldehyde fixed animal 15 days post injection showed dense labeling and the size of the injection site. In the RMS, A large number of newborn cells migrated along the RMS to colonize the different layers of the olfactory bulb. The fusion protein was expressed in all neuron compartments, including the soma dendrites and spines.

Miniature LED implantation allowed large scale and inexpensive optical stimulation of a broad region of interest by characterizing the light diffusion in the brain tissue and the threshold of channel redin two activation. Optical activation was estimated to be triggered at a depth of about two millimeters below the surface of the LED to monitor the functional activation of the channel adoption, two expressing neurons in vivo. The expression of CFOs.

A marker of neuronal activity was analyzed after repetitive light stimulation. One master. This technique can be done in one hour to inject the virus, and after at least three weeks, two more hours to implant LAD.

Calibration of LAD is important and take additional time Following this procedure. Other methods, such as in vivo electrosurgical recording can be performed to answer additional question like about the relationship between activating one type of cell and the response in another After its development. This technique paved the way for researchers in the field of systems neuroscience to explore precise correlations between newborn neuron activation and olfactory perception in the mouse olfactory system.

After watching this video, you should have a good understanding of how to inject a viral vector using STEREOTAXIC coordinates, how to prepare and calibrate an LED for implantation, how to implant the LED and how to use oxygen and stimulation to conduct behavioral experiments. Also, please remember that using viral vectors can be extremely hazardous and proper precautions, such as proper training and disposal of hazardous materials should be taken.