Probing Nicotinic Acetylcholine Receptor Function in Mouse Brain Slices via Laser Flash Photolysis of Photoactivatable Nicotine

Work pertaining to brain slice preparation was reviewed and approved by the Northwestern University Animal Care and Use Committee (protocol #IS00003604).

CAUTION: Lasers used for point photo-stimulation are visible Class IIIb lasers which have the potential to cause harm to the eyes. 2PLSM requires a near-infrared (NIR) Class IV laser (>500 mW), which has the potential to cause serious harm to the eyes and even burns in other tissues. Proper laser beam containment, system interlocks, plus engineered and administrative controls are required to ensure safe operation of laser-based equipment. Always seek out local laser safety personnel when working with lasers.

1. Calibration and Verification of the Uncaging Laser(s)

1. Quantify laser power to be delivered to the sample.
   1. Turn on the 405 nm laser (100 mW maximum power with 5 V control signal) and let the laser system warm up for about 10 min.
      NOTE: The laser is still shuttered (with 0 V drive) and there is no output power until the laser is sent a control voltage to modulate the output power.
   2. Place a power meter in the tissue sample plane or in place of the condenser lens. Manually center the meter relative to the optical path/objective lens.
   3. Set the meter to the correct wavelength range (400-1100 nm). Zero the meter by depressing the appropriate button.
   4. Using software controls, select 100 (out of max 1000; 1000 = 5 V) for the 405 nm laser power, which sets the laser to 10% of full power. If desired, the laser control voltage can also be fed into the PrairieView system via VoltageRecord to provide a digital record of the command signal timing and power level.
   5. Record the reading from the power meter.
   6. Select 150 (15% of max 1000) for the 405 nm laser power output and record the reading from the power meter. Repeat this for the following output powers to collect a laser power curve: 200, 250, 300, 350, 400, 450, 500, 550, 600, 650, 700, 750, 800, 850, 900, 950, and 1000.
2. Calibrate the uncaging laser galvanometers. Execute the following steps whenever there is a change to the optical components of the system, whenever there is a concern about accurate spot positioning, or regularly every month.
   1. Install the 60x water-dipping microscope objective lens that will be used in photostimulation and imaging experiments. In the acquisition/imaging software, select the 60x objective lens and an optical zoom setting of 1 (see step 3.1.4.2.).
   2. Mark a filled circle onto a clean glass microscopy slide with a red permanent marker. Place the slide on the microscope stage, with marker toward the objective.
   3. Pre-focus the microscope on the red marker region with a 4x or 10x objective. Add 1-2 mL of water to the top of the red marker dot/spot and then switch to the 60x objective and submerge the objective into the water. Focus the objective lens on the thin red marker region.
   4. Switch to 2-photon laser scanning. For most systems, move turret to position #1, move the trinocular prism out of the light path, move the scan head mirror to front, and set the laser wavelength to ~900 nm. Select the 512 x 512 pixel box option for the image acquisition parameters, which is the default pixel element for the stimulation mirror calibration routine.
   5. Start system scanning with an imaging laser power greater than minimum and fine tune the objective focus onto the thin red marker fluorescence layer. Choose a field in the fluorescence field clear of debris and evenly coated with marker.
      NOTE: The setup in this protocol uses PrairieView 5.4 acquisition/imaging software.
   6. Open the Uncaging Galvo Calibration function within the Tools – Calibration/Alignment menu of the software. Walk through the burn spots tutorial for the spatial calibration of the second galvanometer mirror pair.
      1. Within the burn spots tutorial, choose the 405 nm laser, select a laser stimulation power of 400 and a stimulation duration of 20 ms. This should yield small (~1-5 µm diameter) holes in the red marker.
         NOTE: Settings such as ~2-4 mW and 1-10 ms are typically used, but the settings are determined by the sample. PA-Nic photo-stimulation power settings are likely to be far lower than the power required during calibration to ablate a visible hole in the red marker slide. This calibration routine is useful for locating photostimulation spots but should not be used to infer absolute photostimulation volumes during physiological responses.
      2. Select Update to stimulate and refresh the image after the center spot burn and move the round red indicator to the actual spot location. Do this for the center spot, the right center spot, the lower center spot, and finally for a grid of nine spots (all corners and edges plus the center of image).
         NOTE: The center, right, and lower corrected spot locations determine the stimulation galvanometer voltages to define the true center and X and Y scaling factors to match the stimulation mirror pair to the imaging mirror pair. The software will scale, and update, all subsequent MarkPoints experiments performed at zoom settings different from the spatial calibration zoom.
   7. Test the calibration by opening the MarkPoints window and manually activating the stimulation parameters in defined spot(s) in a new area of the sample. Ensure that the correct, latest calibration file is loaded into the MarkPoints window. Activate the MarkPoints/Group or MarkPoints Series feature at a defined spot(s) or utilize the Live/Ablation feature to right/left click the mouse anywhere on the image during live scanning to apply a test pulse and to verify correct calibration.
      NOTE: The laser burn spot should now be perfectly centered on the MarkPoints indicator.
   8. Monitor and record the activation of the laser pulse power and temporal duration by tapping off the drive voltage into the VoltageRecord program (see step 1.1.4). Similarly, record the position of each stimulation spot using a scaled voltage signal from the feedback signals derived from the pair of photo-stimulation galvanometer mirrors.

2. Preparation of Photoactivatable Nicotine (PA-Nic)

1. Retrieve an aliquot of lyophilized photoactivatable drug from storage.
   NOTE: The following protocol is specific for PA-Nic; adjust as needed for other photoactivatable drugs. Although PA-Nic demonstrates exceptional stability⁵, take reasonable precautions to protect it from exposure to bright light during preparation and/or experiments. This can be accomplished by simply working in low light; restricting to red filtered light is not necessary.
2. Perform local application of PA-Nic.
   1. Pull a glass micropipette with an opening diameter of 20-40 µm with a programmable pipette puller.
   2. Filter ~1 mL of recording solution with a 0.22 µm filter. Resuspend a quantity of PA-Nic in filtered recording solution to yield a final concentration of 2 mM. For example, dissolve a 100 nmol lyophilized aliquot in 50 µL of filtered recording solution.
      NOTE: A suggested recording solution composition can be found in recent publications^5,6 employing PA-Nic photolysis.
   3. Back-fill the local application pipette with 50 µL of 2 mM PA-Nic.
   4. Secure the local application pipette into a pipette holder mounted on a micromanipulator. Connect the pipette holder via appropriate tubing to a pressure ejection system capable of sustained low-pressure application (1-2 psi).
   5. Using the micromanipulator, maneuver the local application pipette into the extracellular recording solution and position the pipette tip slightly above the mouse brain tissue located ~50 μm from the cell of interest. Consult a previous report for a detailed protocol of mouse brain slice preparation and patch clamp recordings⁸.
   6. Check the application parameters by briefly applying pressure (1-2 psi). There should be minimal to no displacement of the cell of interest. If significant movement does occur, reposition the local application pipette further away (in the lateral and/or axial direction) from the cell of interest.
   7. After achieving stable whole cell patch clamp (details for which are included in a prior publication⁸), turn on low pressure (1-2 p.s.i.) application using the appropriate manual switch on the pressure ejection device. Saturate the tissue surrounding the cell with PA-Nic for 1-2 min before proceeding to the next step.
3. Perform bath application (superfusion) of PA-Nic to the brain slice.
   1. Dissolve a quantity of PA-Nic in a volume of recording solution appropriate for continuous recirculation to yield a final concentration of 100 μM. For example, dissolve a 1 μmol aliquot into 10 mL of recording solution using a standard 15 mL tube.
   2. Begin recirculation of the PA-Nic solution at a rate of 1.5-2 mL/min by opening the appropriate flow control in the perfusion system. Recirculation occurs for the duration of the recording. To conserve valuable drug, minimize the recirculation volume by using tubing with a minimal inner diameter, and/or by shortening the overall length of tubing used in the perfusion system.
      NOTE: By taking these steps, the volume for bath recirculation can be reduced to 5 mL of PA-Nic solution. PA-Nic solutions can often be used for two consecutive recording days within the same week if stored protected from light at 4 °C.
   3. During recirculation, continuously bubble the solution with carbogen (5% O₂, 95% CO₂) and maintain the bath temperature at 32 °C.
   4. Retain the brain slice in recording solution while working with PA-Nic in low light conditions.

3. Imaging Neurons with 2-Photon Laser Scanning Microscopy

1. Perform live visualization of the cell.
   1. Identify/visualize a medial habenula (MHb) neuron using transmitted light or infra-red differential interference contrast (IR/DIC) optics and a video camera and establish a stable whole cell patch clamp recording. Refer to a previous protocol for details on patch clamp recordings from neurons in acutely-prepared mouse brain slices⁸.
   2. After establishing the high-resistance (>1 GΩ) cell-attached configuration, but before break-in, switch the set-up and software to laser scanning mode.
   3. After break-in, use laser scanning to verify that an imaging dye (diluted to a final concentration of 100-200 µM into a standard intracellular pipette solution described previously⁸) is passively (by diffusion) filling the neuron. Allow the dye (e.g., Alexa Fluor 488 in green photomultiplier tube [PMT] channel, PMT 2; or Alexa Fluor 568 or 594 in red PMT channel, PMT 1) to fill the cellular compartments for at least 20-30 min before attempting experiments that require visualization of any cellular compartments outside of the soma.
      NOTE: Distal compartments (dendritic structures, spines, axons, etc.) may require 30-40 min to fill completely²⁵.
   4. Use the software Live Scan function to visualize the neuron and subcellular compartment of interest. Choose imaging parameters that allow for accurate live visualization of neuronal features. Manipulate various settings to affect or alter the display visualization (contrast), resolution, signal-to-noise (S/N) ratio, and image frame acquisition time:
      1. Look-up-table (LUT). Open the LUT window using the appropriate icon on the side of any image window. Once open, adjust the LUT floor (min) and ceiling (max) setting of the specific image channel to enhance visualization of signal contrast shown on the screen. Lower the maximum value to ~1000 (out of 4096, 12-bit detection), which will help to pull out the dimmer signals when first searching for cells, signal, and structure.
         NOTE: These settings only affect the displayed signal, not the detected/recorded values. Human eyes can typically only make out contrast to ~50 grey levels²⁶.
      2. Optical zoom. Use software controls to select 1X optical zoom and use panning controls to locate the desired area in the tissue. This zoom setting yields the largest square, scanned field of view, and sends the largest voltages/scan angles, to the galvanometer mirrors.
         NOTE: The default configuration is for a 12 mm x 12 mm field of view inside the scan head which translates to 12 mm divided by objective magnification inside the sample. Therefore, a 60x objective lens yields scanned images of 200 µm per side at 1x optical zoom. Higher optical zoom values scan less area. 2x optical zoom is often the most useful setting for visualizing whole neurons. 4x can be useful for visualizing subcellular aspects of neurons.
      3. Number of pixels. To complement 1x optical zoom, select 1024 x 1024 pixels per line using software controls. Set the number of pixels per line in the captured and displayed image to not lose details possible from the objective lens. Use the following practical pixel values for a 60x/1.0 numerical aperture (NA) objective: 1024 x 1024 for zoom 1, 512 x 512 for zoom 2, and 256 x 256 for zoom 4. The end pixel size (~0.17 µm; 12 mm/magnification/zoom/pixels) should be half, or less, of the lateral resolution defined by the objective lens.
         NOTE: The image resolution is only defined by the laser wavelength and the objective NA (0.4 µm resolution from two-photon excited [2PE] with 920 nm and a 1.0 NA objective)²⁷. The criteria for the full excitation NA, as listed on the lens, is that the 1/e² intensity of the laser beam diameter matches (or “fills”) the entrance pupil (2 x tube lens focal length x NA / magnification) of the objective lens. The tube lens in the system described here has a focal length of 180 mm.
      4. Pixel dwell time. Use software controls to select 4 µs for the pixel dwell time, a useful default value. 
         NOTE: Changing the pixel dwell time does not change the mean signal detected; it only affects the intra-pixel averaging and these changes can be visualized through the image quality via the S/N. The image pixel dwell time is always a multiple of 0.4 µs units, and for larger dwell times the 12-bit-limited intensity value of each image pixel is the average of the 0.4 µs samples. Since the S/N ratio improves as the square root of the number of samples per pixel dwell time (4 µs equals ten samples, or 3.16-fold improvement in S/N), the improvement in image quality reaches diminishing returns for values much larger than 12 µs.
      5. Scan Rotation and Region of Interest (ROI). Set the image angle to 0° rotation using software controls (no action may be needed, as 0° rotation is the default setting for most imaging systems). If the sample is placed in an “upside-down” orientation, select 180° rotation to “flip” the image.
         NOTE: Rotating the image, at any given angle, can provide a better fit for the entire area of interest of a filled cell. Rotation can also yield a clearer basis for aligning structural changes and for performing subsequent analysis. Selecting a region of interest within the scanned image at a given zoom (step 3.1.4.2) setting retains the native pixel number (step 3.1.4.3), but the restriction in total number of area and pixels can dramatically increase the frame rate, providing improved temporal resolution of signal changes.
      6. Frame Averaging. Select a starting frame average setting of 2 frames using software controls.
         NOTE: The final image contrast (S/N) is defined by the total photons collected/detected within the signal pixels of the image. Averaging of multiple image frames can improve the S/N ratio, provided the sample does not move or is not bleached during imaging. The signal of interest remains the same value during frame averaging while the noise in the image is reduced by the square root of the number of frames averaged. Small structures in fluorescence images often require inter-pixel averaging, combining pixels within the image (often called ROIs), and/or frame averaging. Frame averaging increases scanning time by the number of images one chooses to average.
   5. Use the Pan Control, Scan Rotation, and Optical Zoom tools to orient the sample location while scanning. If motor stage manipulation is necessary to position the sample, avoid large step sizes to the X, Y, and Z axis to prevent objective/condenser collisions, vibrations, or exposure of the laser beam to reflective surfaces.
2. Collect a Z-stack. Using the Z-series tool, select a start and stop position that contains the cell of interest. Select a step size (1 µm) and then consecutively image the neuron in every Z-plane that contains the cell.
   NOTE: Z-stack acquisition settings will vary between neuron type and filling dye. Optimal parameters for Z-stack acquisition should be determined independent of parameters used for live imaging. Z-stack acquisition can be performed before and/or after optopharmacology experiments. If possible, perform Z-stack acquisition after optopharmacology to avoid any cellular damage induced from 2PLSM and to allow for optimal dye filling of small cellular compartments.

4. Laser Flash Photolysis During Electrophysiological Recordings

NOTE: Applying 405 nm or 473 nm laser powers of ≥ 1 mW produces phosphorescence inside the glass of objective and condenser lenses. This generated light is directly related to the laser illumination power; the emission is present in the green and red spectral windows and has excited-state lifetimes in the ms range. This background stimulation artifact is seen in all lenses tested and in water-dipping objective lenses from all major manufacturers of objective lenses. Condenser lenses produce much higher phosphorescence than objective lenses. This “signal” motivates the use of mechanical shuttering for protection of the sensitive gallium arsenide phosphide (GaAsP) PMT cathodes during photo-stimulation events. Using a normally closed mechanical shutter (closed when not actively scanning) represents the best solution for protection of cooled GaAsP PMTs.

1. For an hourglass-type photostimulation beam geometry, remove any focusing lenses in the light path optics that would otherwise narrow/constrict the laser beam as it enters the objective entrance pupil.
2. Using MarkPoints, select the single spot setting.
   NOTE: Other photo-stimulation settings (multiple spots, a grid of spots, spiral scanning) are possible within MarkPoints. Single spot is the simplest. Experimental goals and biological differences may necessitate a different setting.
3. Update the image using the Live Scan option to briefly image and locate the subcellular area of interest. Periodically update the image to identify any potential small drifts in focus.
4. Use software controls to increase the optical zoom (i.e., select a higher optical zoom setting than the current one), if necessary, to visualize small structures (i.e., spines or distal dendrites).
5. Place the MarkPoints single spot crosshairs immediately adjacent (~0.5 μm) to the cell membrane. Do not place the photo-stimulation spot directly over a cellular feature, as this could lead to photodamage.
6. Set the parameters for photo-stimulation using software controls in MarkPoints. Apply the starting guidelines as follows: 1-50 ms duration, 1-4 mW laser power, and ≥1 trial.
7. Select Run MarkPoints to initiate the MarkPoints protocol and observe electrophysiology data acquisition in real time.
8. Repeat steps 4.2-4.7 several times to evaluate consistency and stability, or lack thereof, of the response amplitude and kinetics.