The overall goal of the described experiments is the monitoring of membrane potential signal integration and information processing by an individual nerve cell optically with sub micrometer and sub millisecond resolution from multiple sites on a neuron. This is achieved by first building an imaging setup with appropriate sensitivity to allow the recording of small fractional changes in light intensity corresponding to the membrane potential transient. Next, select the neurons with appropriate anatomy for effective imaging neurons with long intact processes in a single focal plane close to the surface of the slice.
Then load the preselected neurons with voltage sensitive dyes to allow optical tracking of the membrane potential changes. Results show the unique capability of this approach to obtaining information on electrical signal integration by individual nerve cells by monitoring membrane potential signals from axons, dendrites, and dendritic spines. In cellular neurophysiology.
New tools are being developed because of the two longstanding limitations of conventional electrophysiology, low spatial resolution of electrode measurements, and inability to record from small structures such as axon collaterals or dendritic spines. To overcome these drawbacks, patch electrode approach can be combined with optical techniques that permit permit parallel recordings from all parts of a neuro. Here we describe such a technique, multiple side optical recording of membrane potential transients with single spine resolution.
In this experiment, an upright microscope and three cameras are used to adjust for uniform illumination. First, insert an appropriate neutral density filter in the laser beam path. Next, focus the objective on the surface of the standard fluorescent slide.
While observing the CCD image of the field of view on the monitor screen, adjust the position of the receiving end of the quartz light guide and the position of the output end of the light guide using appropriate actuators on the epi fluorescence condenser in order to achieve centered and uniform illumination of the field of view. To determine the vibrational noise, use a standard fluorescent slide with a non fluorescent mark on the surface. Record the light intensity with the neuro CCD in continuous mode.
Focus the objective on a dark edge of the black mark a record the light intensity for about 100 milliseconds at two kilohertz frame rate. Then display the spatial average of the fractional light intensity traces from approximately 20 pixels receiving light from the uniformly illuminated area and from approximately 20 pixels along the edge of the black mark. The excess noise in the spatial average from the pixels with high contrast edges reflects the vibrational noise in the system.
It is essential to have the mechanical shutter mounted off the vibration isolation table so that shutter opening does not introduce vibrational noise in the optical recording. This is a critical step. Mount the shutter using plastic screws and vibration isolation material to reduce the acoustic vibrational noise that can be picked up by the optical recording system.
To isolate the vibration, adjust the vibration isolation table until the vibration noise of the pixels covering the sharp edge of the image is smaller than the shot noise. The equipment wires on the table must be loose in order to avoid transmitting the mechanical vibrations to the microscope scope. In this procedure, prepare the brain slices of 300 to 400 micron thickness according to standard procedures with a mouse line expressing EGFP in individual nerve cells of interest with a spinning disc confocal system.
Visualize the EGFP labeled neurons in the slice. Next, select the neurons with intact dendritic axonal trees, and with processes running parallel and close to the surface of the slice for membrane potential imaging. Then fill a glass patch pipette from the tip with dye-free intracellular solution up to two thirds of the electrode taper by applying negative pressure for about 15 seconds.
After that, backfill the electrode with the solution containing the membrane IMP permeate voltage sensitive dye dissolved in the intracellular solution. Now, patch a previously selected neuron within one minute of loading the dye into the electrode. Apply low positive pressure about 10 millimeters of mercury before the pipette is lowered into the bath, and while positioning the electrode in the slice, this ensures that no dye leaks out of the electrode before the seal is established.
Increase to high pressure around 100 millimeters of mercury just before the electrode contacts the cell surface. Establishing the seal in the whole cell configuration, allow the free diffusion of the dye from a somatic patch pipette to the soma for 20 to 45 minutes. During the dyed diffusion, monitor the physiological state of the neuron by recording the evoked action potentials in the current clamp mode.
Additionally, monitor the amount of staining by recording the resting light intensity from the cell soma at a frame rate of two kilohertz and at a fraction of full light intensity adjusted with neutral density filters. Continue the downloading process until the action potential starts to broaden. At the end of staining, carefully pull the patch pipette away from the soma in voltage clamp configuration to ensure that the transition from whole cell to outside out patch configuration is attained in the process.
S then incubate the slice for an additional 1.5 to two hours at room temperature to allow the voltage sensitive dye to spread into the neuronal processes. After a significant amount of dye has diffused away from the soma into the dendritic and axonal processes, the wave form of the action potential should be completely restored. In the next step, locate the soma of the stain neuron under the low intensity fluorescence.
Reduce the light intensity with neutral density filters to the minimum required to visualize the object of interest using an XY stage Scan the area of the slice containing the fluorescent neuron until the fluorescent soma is identified. Position the soma of the stained cell in the center of the microscope field rep atch the neuron with a standard patch electrode with no dye under the DIC. Then visualize the neuronal processes under the low intensity fluorescence at a frame rate of 10 to 40 hertz in the continuous recording mode of the CCD for voltage imaging.
Using an XY stage position, the neuronal process of interest in the middle of the imaging area. In addition, protect the SOMA from the excitation light using the partially closed field stop iris of the microscope to reduce the photodynamic damage. Next, record the optical signals related to back propagating action potentials in the individual dendritic branches.
Then record the optical signals related to action potentials in the axon. After that, record the optical signals related to back propagating action potentials in the dendritic spines. Use the appropriate frame rate for accurate reconstruction of the signal wave form.
At the same time, keep the recording periods and the exposure to high intensity excitation light as short as possible in order to minimize dye bleaching and photodynamic damage. The recording of optical signals is shown here.Here. The upper panel shows the high resolution confocal image of a stained neuron with its axon in the recording position.
While the middle panel shows the low spatial resolution fluorescence image of the axon obtained by CCD used for membrane potential imaging, the lower panel presents the electrode recordings from the soma and optical recordings from the A IS and the node of rvi. The raw data and processed signals are shown. This figure shows the action potential signals from the axon of an L five cortical neuron loaded with the voltage sensitive dye action.
Potential related signals were recorded at a frame rate of 10 kilohertz. These traces indicate the action potential transient from three locations. One, the electrode recording from the soma.
Two, the optical recordings from the axon hillock, and three, the optical recording from the first note of vie, and here are the superimposed signals from the same three locations. This figure shows a burst of four back propagating action potential signals from multiple locations on the dendritic arbor of an L five cortical neuron initiated by short depolarizing current pulses at 100 hertz. Shown here are the action potential signals from an individual dendritic spine.
Here is the anatomical reconstruction obtained from a stack of spinning disc confocal images, and here is the fluorescence image of the same region obtained with the CCD camera. For membrane potential imaging, the fluorescence intensity traces corresponding to back propagating action Potentials from locations one through three as outlined on the CCD image are shown in the right panel. The bottom trace indicates the electrode recording from the soma.
The prerecording preparation takes about three hours, including the waiting period for the dice spreading to distal processes. The recording time is limited by normal rundown of the preparation and by PO photodynamic damage. The amount of photodynamic damage is proportional to the excitation light intensity, and to duration and number of individual recording trials.
In some experiments, it is possible to carry out more than 50 recording trials over one or two hours. Other measurements require high excitation, light intensities, and longer recordings, and are limited by photodynamic damage to only several recording trials. After watching this video, you should have a good understanding of how to carry out voltage imaging from the entire axonal and endr structure of individual neurons in brain slices.
