Miniscope in vivo calcium imaging empowers researchers to examine the spatially and temporally coordinated activity from hundreds of neurons in deep brain structures of freely behaving animals. Miniscope in vivo calcium imaging offers many advantages, including the ability to perform recordings that are cell type specific, large scale, and longitudinal. Our surgery protocol for viral injection and GRIN lens implantation is compatible with any commercial or custom built single photon and two photon imaging systems for the deep brain in vivo calcium imaging.
Demonstrating the procedure will be Rashmi Thapa, a graduate student from my laboratory. After disinfecting the shaved surgical area of an anesthetized mouse, use a scalpel to make a two centimeter incision through the skin along the midline to expose the lambda and bregma of the skull. After locating the 0.5 millimeter dental drill burr to a position of anterior posterior 1.94 millimeters and medial lateral 0.5 millimeters from the bregma, start drilling through the skull.
Next, load the virus into the microliter syringe using the control panel of the micro pump and withdraw 500 nanoliters of air bubble followed by 800 nanoliters of the virus at a flow rate of 50 nanoliters per second. Slowly move down the needle into the brain tissue to the targeted Z coordinate of dorsal ventral 1.75 millimeters and then slightly move it up to the Z coordinate of dorsal ventral 1.65 millimeters. On the control panel, set the micro pump to inject 500 nanoliters of the virus at the flow rate of 50 nanoliters per minute before hitting the Run button to inject the virus.
When the injection is over, line up the skin edges and carefully close the incision with a 4-0 suture. Apply antibiotic ointment on the stitched area to prevent infection. To implant gradient index or GRIN lens, excise a triangular area of the skin of 1.5 centimeter height and 1.5 centimeter base from the anterior side between the eyes to the posterior side behind the lambda using fine scissors.
After the skull is thoroughly cleaned and dried, apply cyanoacrylate to the edges of the skin and attach the skin to the skull. After five minutes when the cyanoacrylate dries, locate a dental drill burr of 1.2 millimeter diameter to a position of anterior posterior 1.94 millimeters, medial lateral 0.8 millimeters from the bregma. Drill through the skull, followed by removing the dura using a 30 gauge needle tip.
Clean all pieces of bone debris with 45 degree angled sharp forceps. Next, attach a 27 gauge manually polished blunt end needle to the needle holder coupled to a robotic arm tilted with an angle of 10 degrees and connect the other end of the needle holder to the house vacuum system. Locate the tip of the needle to just touching the bregma before clicking on Bregma button to set the Z coordinate of bregma to zero.
Set input X value to 0.8, input Y value to 1.94, input Z value to 1.0 and click on Find button to move the needle onto the top of the drilled hole on the skull. Center the tip of the needle to the drilled hole and bring it down to the Z position zero. Turn on the vacuum and start rinsing the exposed brain area with artificial cerebrospinal fluid, or ACSF, through a gravity controlled tubing system connected to a 30 gauge needle with a bent tip.
ACSF gets continuously bubbled with a gas mixture of 95%oxygen and 5%carbon dioxide while being filtered through a 0.2 micron filter. Using AutoStereota software, aspiration of the brain tissue will complete in four rounds. For the first round of aspiration, make sure the Z value displays zero.
Then in the zStep session, click and check the first and second rows. Set the values for the first row to 0.2 and 1. Set the values for the second row to 0.15 and 4.
In Mode session, set needle size 27 gauge and 1.2. Set Dims 0.9. All other values will be default.
To start aspiration, sequentially click on the NotSet, KeepZero, and Start buttons. The first round of aspiration generates a column pocket with 0.8 millimeters in depth and 1 millimeter in diameter. In the third round of aspiration, click and check the first row for zStep session and set the values for the first row to 1.8 and 1.
In Mode session, set needle size 27 gauge and 2.2. Start the aspiration by keeping other values the same as that of the previous round. After the third round of aspiration, the depth of the column pocket is 1.8 millimeters.
The last round of aspiration is to clean up blood accumulated at the bottom of the pocket. Set the values for the first row to 1.6 and 1 in zStep session and set needle size 27 gauge and 2.2 and Dims 0.6 in Mode session before starting aspiration. Once the pocket is blood free, stop the vacuum, stop irrigation of ACSF, and bring the needle up 2 millimeters in Z coordinate and 0.5 millimeters anterior to the center.
Place the sterile 1 millimeter GRIN lens into the brain tissue pocket. Next, apply the melted agarose in the gap between the GRIN lens and brain tissue with the help of a spatula. After agarose forms a gel, remove excess agarose using a micro blade.
Mix the dental cement powder and catalyst liquid in a pre-chilled mixing well and apply a layer of self-curing adhesive resin cement on the skull, starting by surrounding the GRIN lens and then covering the entire exposed skull. In a clean plastic well, mix dental cement powder and black charcoal with liquid to apply a thin layer of the mixture on top of the first layer of dental cement. Let it harden for five minutes.
Connect the Miniscope to the cable and turn on the custom developed software NuView. In NuView, click Hardware, check LED1 and click the Stream button to view the live images. To stop live streaming, click on Stop button.
Next, settle the Miniscope into a custom-built Miniscope holding arm. With the help of motorized controllers, locate the Miniscope just above the exposed GRIN lens, making it parallel to the surface of the lens. Slowly bring down the Miniscope towards the GRIN lens and adjust its Z position until the best plane of focus is found.
Apply the first layer of dental cement around the Miniscope base without altering the position of the Miniscope. After the cement is hardened, gently remove the holding arm such that the Miniscope can stand on its own on the mouse head. Apply a second layer of dental cement around the base to fill all the gaps, ensuring there is no LED light leakage from the gaps.
Let the dental cement harden. After briefly anesthetizing the mouse with isoflurane, loosen the locking screw in the base with a small screwdriver before removing the protective cap and clean the surface of the GRIN lens with an acetone-soaked cotton swab. Connect the Miniscope to the cable and start the NuView software to begin identifying the best focal plane with adjusting the position of the Miniscope relative to the base by slightly tightening or loosening with the help of blunt forceps.
Once the best focal plane is determined, tighten the locking screw before disconnecting the cable and placing the mouse back in its home cage. Turn on the behavior camera software to view the mouse behavior arena through a livestream function. Manually adjust the focus of the top camera.
Select Trigger Strobe and check enable/disable trigger, followed by clicking Record button. Then use Browse to select a location where behavior recordings will be saved. Select the desired image format.
Bring the mouse close to the arena and connect the Miniscope to the cable linked to the data acquisition system. Then place the mouse in the center of the arena. In the behavior camera software, click Start Recording.
In NuView, check LED1, click on Capture and then trigger buttons to start recording. This allows simultaneous recordings of calcium imaging and mouse behavior. Hit Stop after 3, 000 frames and save the files.
When the recording is done, briefly anesthetize the mouse with isoflurane, detach the Miniscope from the base and put the protective cap over the base before placing the mouse back in its home cage. The cell map from suboptimal in vivo calcium imaging contained some active neurons, while that from successful imaging included several hundreds of active neurons in the focused area. In the typical example of an unsuccessful in vivo calcium imaging, the region was observed to be dark and contained less than five active neurons, or the region was bright but had no active neurons.
In the representative analysis, the maximum projection fluorescence cell map and calcium transients from a successive in vivo calcium imaging recording are shown. The postmortem assessment for GCaMP6f expression and GRIN lens implantation in the medial PFC of an experimental mouse indicated that GCaMP6f was expressed and the GRIN lens was implanted precisely in the desired brain region. To obtain a good calcium imaging, it is necessary to ensure that the bleeding has completely stopped and the field is free of blood clots before implanting the GRIN lens.
This technique can be utilized to study neural circuit changes in mouse model of different human brain disorders and test if a drug candidate can normalize the altered circuitry.
