TITLE: Ex vivo mouse brain patch clamp recordings combined with uncaging and optogenetic stimulation
AUTHORS: enrico.zampese

[DESCRIPTION]
In this protocol we detail the steps to perform ex-vivo brain slices electrophysiology and optogenetic stimulation and/or 2-photon uncaging recordings.

[BEFORE_START]
Please note that optogenetic experiments require the expression of a genetically encoded opsin in neurons of interest. Mice genetically engineered to express opsin in specific neuronal populations are available. Alternatively, opsin expression can be obtained via injections of appropriate viral vectors.
Please refer to existing protocols for brain slices preparation and/or viral injections.

[GUIDELINES]
Please adhere to institutional guidelines and regulations.

[STEPS]
SECTION: Prepare patch pipettes
1. Turn on the Sutter P-1000 puller and enter the desired pull protocol.

SECTION: Prepare patch pipettes
2. Insert a thick-walled borosilicate glass capillary and press pull.
SECTION: Prepare patch pipettes
3. Pipette resistance must be of 3 to 5 megaohms.
SECTION: Setting up patch rig and environment
4. Turn on the MultiClamp 700B Amplifier, Axon Digidata 1550B digitizer, micromanipulator, computer tower and the associated software. Note: amplifier and digitizer must be turned on prior to opening software.
SECTION: Setting up patch rig and environment
5. Turn on O2/CO2 tank and bubble aCSF solution.
SECTION: Setting up patch rig and environment
6. Take an aliquot of internal solution from the -20° fridge.

SECTION: Setting up patch rig and environment
7. Fill syringe with internal solution, place a filter on the end of the syringe, and place a MicroFil Pipette Filler on the end of the filter.
SECTION: Setting up patch rig and environment
8. Start perfusing microscope chamber with aCSF (either with a gravity system or a peristaltic pump; a gravity system is normally preferred because of the lower electrical noise).
SECTION: Setting up patch rig and environment
9. Adjust the flow rate to the desired value (recommended at least 2 mL/min).

SECTION: Setting up patch rig and environment
10. Depending on the experimental protocol, turn on stage heater and set to temperature. 

SECTION: Examine slices and patching cells
11. Secure down slice with a harp (slice anchor).
SECTION: Examine slices and patching cells
12. Locate and focus the desired brain region under the 4x objective.

SECTION: Examine slices and patching cells
13. Change the microscope lens to the 60x objective.
SECTION: Examine slices and patching cells
14. Focus on healthy neurons in slices for patching.
SECTION: Examine slices and patching cells
15. For whole cell configuration, fill a glass micropipette one-third full of internal solution. Ensure there is no residual internal solution on exterior of glass micropipette, as this may introduce salts into the micromanipulator and add additional noise to recordings. Remove any air bubbles by gently flicking the glass micropipette.
SECTION: Examine slices and patching cells
16. Gently place the glass micropipette onto the electrode holder.
SECTION: Examine slices and patching cells
17. Position the electrode using a micromanipulator.
SECTION: Examine slices and patching cells
18. Under the 60x objective, bring the tip of the glass pipette above the slice.
SECTION: Examine slices and patching cells
19. Apply a positive pressure and maintain it. Zero the voltage offset.
SECTION: Examine slices and patching cells
20. Approach the cell diagonally. The positive pressure should create a small dimple on the cell.
SECTION: Examine slices and patching cells
21. Once a dimple is formed, release the positive pressure, and apply a small amount of negative pressure. The resistance should begin to increase rapidly.
SECTION: Examine slices and patching cells
22. As the resistance increases, clamp the cell at your resting potential of interest (typically - 60 mV).
SECTION: Examine slices and patching cells
23. After a giga-ohm seal is formed, apply a few quick pulses of negative pressure to break into the cell to record in whole cell configuration.
SECTION: Examine slices and patching cells
24. Start recording.
SECTION: Recordings and uncaging/optogenetic stimulation
25. Once in whole cell configuration, allow time (15-20 mins) for the internal solution containing the Alexa Fluor 568 to fill the cell so that the dendritic tree and individual spines will become visible under 2P imaging. 
SECTION: Recordings and uncaging/optogenetic stimulation
26. Change the setup configuration to 2-photon imaging, using PrairieView. Alexa Fluor 568 is excited with a 810 nm wavelength.
SECTION: Recordings and uncaging/optogenetic stimulation
27. Experiments are performed on distal (>100 microns from soma) dendritic spines within planar sections (20 microns) of the same dendrite with zoom 4 and a field of view (POV) of 50 microns.
SECTION: Recordings and uncaging/optogenetic stimulation
28. Electrophysiological recordings, 2-photon imaging, and stimulation are acquired and synchronized via the PrairieView software.
SECTION: Recordings and uncaging/optogenetic stimulation
29. Single-photon optogenetic stimulation is obtained with a 473 nm laser, whose pulse number, intensity, and duration are controlled via PrairieView. We recommend starting with a single spot of 8 microns diameter, 3ms duration.
SECTION: Recordings and uncaging/optogenetic stimulation
30. For glutamate uncaging, DNI-glutamate (5 mM) is perfused via a microinjectior on the recorded area immediately (~5s) before initiating the stimulation protocol. 1 ms pulses are applied in proximity of single spines located in the same focal plane.  The laser power and the spot location should be calibrated to evoke a somatic excitatory postsynaptic potential of ~1-2mV.
SECTION: Recordings and uncaging/optogenetic stimulation
31. For combined blue laser stimulation and glutamate uncaging, the 3ms optogenetic stimulation precedes the uncaging protocol (1ms delay), which normally includes the stimulation of 15 spines with 1ms pulses 1ms apart.
SECTION: After the experiments
32. Gently remove and discard the recording pipette
SECTION: After the experiments
33. Stop imaging protocols and close shutters to protect the imaging system.
SECTION: After the experiments
34. Remove slice from the recording chamber. If desired, the slice can be gently transferred into a plate and bathed with a fixative solution for further processing.
SECTION: After the experiments
34.1. Note: use appropriate PPE when working with fixatives, including operating under a chemical hood.
SECTION: After the experiments
35. Discard waste according to institutional protocols.
SECTION: After the experiments
36. Rinse perfusion tubing microscope chamber with water and/or a 10% ethanol in water.
SECTION: After the experiments
37. Save and export files.