Preparation of acute brain slice for ex vivo time-lapse imaging

Preparation of acute brain slices

Complete growth media composition (with 5% serum and NGF+/GDNF+):

Prepare 100ml slice culture with 5% serum 

• Mix 40 ml MEM (Minimal essential medium, INV-42360032, Invitrogen), 20 ml BME (Basal medium eagle, INV-21010046, Invitrogen), 5 ml of 5% heat inactivated horse serum (INV-26050070, Invitrogen), 0.8 ml Glutamax (Thermo Fischer Scientific, Cat# 35050061), 0.8 ml Penstrep (Thermo Fischer Scientific, Cat# 15140122), 1.1 ml Glucose (45% w/v, Sigma-Aldrich, Cat# A4403), 32.3 ml of tissue culture grade water (ultrapure). 

• Prepare desired concentrations of NGF and GDNF required for a 50 ml 5% serum slice culture media using the main stock: 50 ul (NGF-10 ug/ml main stock, INV-A42627, Invitrogen), 50 ul (GDNF-10 ug/ml main stock, INV-AF-450-44, Invitrogen), 49900 ul (CGM - Complete Growth Media with 5% horse serum).  

• For new NGF stock concentration of 50 ug/ml, the compositions are as follows: 10ul NGF (50 ug/ml main stock), 50 ul GDNF (10ug/ml main stock), 49940 ul CGM.

Equilibrating 6 well membrane inserts in MEM media 

• Before starting the experiments, ensure that the membrane inserts are calibrated by placing the inserts in a 6 well plate dispersed over MEM basal media for 1-3 hours in cell culture incubator at 37 deg C. (For long term cultures, keep the membrane inserts for equilibration in basal media overnight at 37 deg C).

Connecting wires to oxygen cylinder (carbogen)

• Use a sterile tube which fits the knob of cylinder. For successful oxygenation of buffer, use a sterile Pasteur pipette (2ml) which fits exactly the sterile tube connected from the knob of the oxygen cylinder.

Cleaning the vibratome chambers/dissection instruments for avoiding any contamination

• Spray the dissection chamber, chamber tissue holder and blade holder of the vibratome with 70% ethanol and wipe thoroughly.  Sprinkle ultrapure water to these regions and again wipe out completely to remove any excess solution. Make sure that the regions are dry and clean before euthanasia.

• Also wipe the surgical instruments with 70% ethanol and then wipe with ultra-pure water.

Oxygenation of ACSF (Artificial cerebrospinal fluid solution) prior to the initiation of mouse euthanasia

• Oxygenate 100 ml of ACSF A (LRE-S-LSG1000-1, EcocyteShop.com) in a conical flask or sterile measuring cylinder using sterile Pasteur Pipette (2 ml) connected to a cylinder for 10 minutes.

• Then pour 100 ml of ACSF B (LRE-S-LSG1000-1, EcocyteShop.com) gently through the walls of the flask and then oxygenate the combined solution for another 10 mins.

• If ACSF A and B are 10X concentrates, pipette out 135 ml ultrapure water and then add 7.5 ml ACSF A concentrate, oxygenate this solution for 10 minutes. Thereafter, add 7.5 ml of ACSF B concentrate and then oxygenate the combined solution for 10 minutes.

• Keep the whole buffer slightly oxygenated until the animal euthanasia is complete.

Mouse euthanasia 

• Place the animal in the live euthanasia chamber for CO₂ asphyxiation. As soon as the animal loses its breathing (approx. 5 to 7 mins), place the mouse in biosafety cabinet. 

Cutting of acute brain slices

• Place the animal over the working pad. Separate the head using the blade and place the head immediately into oxygenated ACSF. Using scissors and forceps, dissect out the brain from the skull and immerse it into fresh oxygenated media in a small Petri dish.

• Cut out the brain stem and glue the brain to the vibratome chamber holder, such that the region, where the brain stem was cut out, is positioned at the bottom. Before glueing make sure that the surrounding region of the vibratome chamber is slurred with ice. (Important: ice should not fall into the ACSF buffer, it might change the pH and can also lead to loss of cell viability).

• Fill the vibratome slicing chamber with oxygenated ACSF until the tissue block is fully submerged. Orient the brain so that the cortical region faces the slicing blade. Secure the blade in the holder with a slight downward tilt to ensure smooth sectioning. Begin sectioning from the outer coronal surface of the brain, cutting sequentially thick sections at 120 µm, 220 µm, and 420 µm until reaching the brain mid-height. Once approaching the ventral regions of the brain, adjust the vibratome settings to a thickness of 20 µm and cut 20 µm sections for acute slice imaging. Important: optimize the vibratome cutting speed to achieve the best quality of the resulting slices. Slice quality should be assessed using the following criteria: structural integrity, minimal tissue tearing or compression, smoothness of cut surfaces, and preservation of cortical architecture.

• Transfer the slices from the coronal cortical regions and discard the sections from other regions using a sterile transfer pipette. Transfer desired sections to 24 well plates. Make sure that you have at least three high-quality 20 um thin coronal cortical slices to be placed into membrane inserts of a 6 well plate.

• After transferring the slices to a membrane insert, wash off the old basal media and supplement with 1 ml of fresh 5% serum complete growth media. 

Calbryte 590AM staining

• For imaging calcium transients, add 0.15 ul of Calbryte 590AM (1 mg/ml, VWR-76484-390-EA, AAT Bioquest) from the main stock directly into the CGM with 5% horse serum. Shake manually the well with the slices for even distribution of the dye; then incubate at 37 deg C for 30-35 mins.

Hoechst 33342 nuclear staining

• After incubating with Calbryte 590AM for 35 mins, add 0.24 ul of Hoechst 33342 and incubate the same slices for additional 10 mins at 37 deg C in the cell culture incubator.

Washing and time-lapse imaging 

• Take out the slices by holding the membrane insert and replace the wells with 1 ml of fresh MEM basal media and incubate the slices at 37 deg C for 5 mins.

• Repeat the following steps for additional 3 rounds of washing (total of 4 washes).

• Replace the basal medium with 1 ml of complete growth slice culture medium with 5% horse serum.

• Either transfer the slices to a new coverslip petri dish/well plate or remain in the same 6 well plate and proceed for time lapse imaging in Leica MICA.

• First enter the experiment name with Date after opening the Leica MICA software.

• Switch on the live climatic conditions by turning on the temperature and CO₂ (ensure that this is done at least 15 mins before starting the experiment).

• If coverslip bottom petri dish is used for live time lapse imaging, make sure a petri dish is chosen from the drop-down options. 

• Choose 10X and then image each channel sequentially, make sure to check each channel fluorescence separately. If there is a high background, look for reducing the channel intensity by drawing the intensity arrow to the left. Also make sure that for each channel the image quality is marked to zero.

• Ensure 3D z stack is featured with the green channel, and then choose multiple regions from the same plane. Choose all the regions and then select stage focusing, by selecting the autofocus to green channel (microglia). 

• Specify the time lapse accordingly: 3 mins for time intervals, and 3 hrs for total time duration.

• In case if you are using a well plate made of plastic, instead of focusing each well by clicking on autofocus, choose the selection focus maps, then select different regions, and click on set z sequentially.