Single Cell Electroporation in vivo within the Intact Developing Brain

Single cell electroporation is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells with an intact tissue, including in vivo preparations when combined with advanced in vivo imaging techniques. Single cell electroporation of fluorescent markers permits direct visualization of cellular morphology, cell growth, and intercellular events over timescales ranging from seconds to days. In this video, we detailed the procedure for single cell electroporation of a fluorescent dye or plasma DNA into neurons within the intact brain of the XUS tadpole, and show examples of live imaging of fluorescently labeled neurons.

Oh, hi. My name is Eth Haa and I work in the laboratory of Dr.Kurt Haass in the Brain Research Center at the University of British Columbia. Today, I will demonstrate how to perform targeted single cell electroporation of neurons within the intact brain of the albino XUS Lavis tadpole.

The distinct advantage of this technique is that experimental manipulations may be performed on individual cells while leaving the surrounding tissue unaltered, thereby distinguishing cell autonomous effects from those resulting from global treatments. The first step involves fabricating micropipets with appropriate tip diameter and taper angle, followed by loading transfer material into the micro pipet. Then TA pools must be anesthetized and transferred to the ation chamber, followed by inserting the micro pipette into the brain and applying a train of voltage pulses.

Finally, ated neurons are imaged using two photon microscopy. They'll be showing you this technique in tadpoles. Single cell ation can be used in other model systems as well as in ex vivo preparations.

So with that in mind, let's get started. Single cell electroporation targets an individual cell by use of a pulled glass micro pipette electrode that restricts the electric field to a single cell in contact with the pipette tip. The physical geometry of the pipette tip is a critical feature directly affecting the success of the technique.

A solder model P 97 puller is used to generate micro pipettes from bo silicate glass with an outer diameter of 1.5 millimeters and an inner diameter of 0.75 millimeters containing an internal filament to facilitate filling of the tip without air bubbles. Pipette tips that have been found to be most effective for single cell electroporation are those with a tip diameter of 0.5 to one micron and taper angles of greater than 10 degrees. After pulling a glass micro pipette transfer material in solution can be loaded into the pipette by back filling or by using a loading syringe.

For this demonstration, we will be fluorescently labeling single neurons with a fluorescent dye seen here or with a plasmid encoding. GFP green fluorescent protein solutions should be briefly centrifuge at high speed to reduce the amount of particulate debris that enters the pipette. Since this will increase the chances of the tip clogging for new users, it is advisable to begin with fluorescent dyes since they can be readily seen under epi fluorescence.

This enables the visualization of the tip within the sample and gives direct feedback as to whether the equipment has been properly set up and whether the pipette tip is appropriate. Good job. Now that we have prepared the micro pipette, we can move on to performing the procedure for single cell electroporation.

The electrical equipment required for this technique are a stimulator and oscilloscope and the pipette holder fitted with a silver wire to be inserted into the pipette. Here we are using an axon instrument's ax separator 800 A, however common stimulators such as a grass SD nine can also be used. The ax separator is connected to the head stage, which interfaces between the ax separator and the silver wire electrode.

The tadpole external ground is simply a silver wire that remains an electronic contact with the tadpole. During electroporation, the head stage, ground lead and the tadpole external ground are both connected to the oscilloscope input. Once the electrical equipment has been set up, the pipette can be placed in the pipette holder.

The pipette holder should be mounted on a three axis micro manipulator that allows fine movement. Single cell electroporation requires a microscope with a good working distance. Since this gives more room to maneuver and allows the pipette to be brought in at an angle of about 30 to 45 degrees.

Once the equipment has been properly connected, we can begin with a single cell electroporation procedure. First tadpoles are anesthetized by immersion in 0.02%MS 2, 2 2, the tadpoles are usually fully anesthetized. After approximately five minutes, a single tadpole is transferred to the electroporation chamber.

Using a plastic transfer pipette, the chamber can be easily made in-house by carving a tadpole shaped cavity in a small silicone block and inserting a silver ground wire such that it'll be in contact with the tadpole skin when in the chamber. Next, using a paintbrush to position the tadpole will ensure that it is dorsal side up. The micro pipette is then lowered until it is in contact with the skin directly adjacent to the region of interest.

Continued lowering of the pipette will initially dimple the skin and then pass through into the underlying tissue. Here I am targeting the optic tum of the tide pole brain. I'm specifically aiming for the cell body layer region.

If you are using an xi, one can monitor the resistance of the pipette in the tissue. Generally, resistance is of 10 to about 40. Mega ohms work well.

The resistance of the pipette is a good indicator of whether the tip diameter is too small or if the tip is clogged, in which case the resistance will be high. On the other hand, if the tip diameter is too large or if the tip is broken, the pipette resistance will be low. Although the absolute resistance, the pipette cannot be measured when using a conventional stimulator.

The oscilloscope reading is an indirect measure of the pipette resistance for fluorescent dextran dies. The stimulus consists of a 10 millisecond train of positive square wave pulses applied at a frequency of 300 hertz. Individual pulses are 300 microseconds in duration and are delivered at a voltage of 50 volts.

Using these settings, the fluorescent dye is ejected from the tip as can be seen upon application of a stimulus after electro probing. One site, the pipette is retracted and the process is repeated at different sites within the tissue to improve yield. While a voltage pulse train is being delivered, one should monitor the amplitude of the resulting pulses on the oscilloscope and adjust the amplitude to between one and 1.5 microamps if necessary.

If the pulse amplitude remains low after substantially increasing the voltage or drops during the course of the procedure, this strongly suggests that the tip is clogged or that the tip diameter is too small. This can be pretty frustrating when it happens. On the other hand, if the amplitude is still high after substantially reducing the voltage applied or rises during the course of the procedure, this strongly indicates the tip is either too wide or that it is broken, or tips break.

I often become very emotional and sad. As mentioned earlier, the advantage of using fluorescent dyes is that one can immediately check whether single cells have been electroporated using the given stimulus parameters. If cells appear very dim or no cells are labeled, one would start by gradually increasing each stimulus parameter independently until the labeled cells appear to be brightly labeled like this, seeing nicely labeled cells makes my day.

On the other hand, if numerous cells appeared to be labeled, one would gradually decrease the stimulus parameters until it appeared that a single cell was brightly labeled like this one. Seeing brightly label cells is really satisfying. In the case of plasmid DNA, the pulse trains used are negative pulses are one millisecond in duration delivered at 300 hertz, and the train duration is 500 milliseconds.

After electroporation, the TA poles are placed in fresh rearing solution where they rapidly recover from anesthesia. Typically, after electro 50 to a hundred TA poles, we can proceed to screening them, which I'll show you next. Before beginning imaging experiments, it is useful to quickly screen electro tadpoles using an upright epi fluorescence microscope.

In order to determine which tadpoles contain neurons that are suitable for imaging tadpoles are anesthetized and then placed in a cigar chamber and cover slipped individual tadpole Brains are then rapidly viewed under epi fluorescent light to determine whether single fluorescently labeled neurons are present. Note that excessive exposure to the epi fluorescent light will result in phototoxicity that may kill the labeled cell. Therefore, try to minimize the amount of time that cells are exposed to the fluorescent light.

In the case of dextran dyes, screening may take place as soon as 30 minutes post electroporation. However, we find that longer intervals are associated with lower background fluorescent levels. In the case of plasmid, DNA screening is typically done at least 12 hours post electroporation.

We routinely use single cell electroporation to fluorescently label neurons within the tadpole optic tectum. Our two photon microscope is custom built, constructed from an Olympus FV 300 confocal microscope and a Chameleon XP laser light source. We typically use a wavelength of 910 nanometers to image green floor force.

Once we've found a tadpole with a single labeled AL neuron and position it on our two photon rig, we proceed to perform time lapse to photon imaging of tactile neurons over different time intervals. This allows us to observe the dynamic growth patterns of the dendritic arbors of AL cells. Here we see an example of a GFP expressing neuron that had been labeled using single cell electroporation.

This image was captured using in vivo two photon microscopy in order to quantify neuronal growth. Skeletonized three dimensional reconstructions of each neuronal image are typically generated. Time-lapse imaging with protracted intervals provides insight into the long-term consequences of an experimental manipulation.

Here we see an immature GFP expressing neuron that has been imaged daily over the course of four days, spanning the elaboration of its dendritic arbor. These images were captured using two photon microscopy and are presented as Z projections. Rapid time-lapse imaging allows us to observe the nature of more dynamic processes of neuronal growth, including the formation and retraction of dendritic filopodia.

This video shows a relatively mature GFP expressing neuron that was imaged in three dimensions every two minutes for one hour. This is an example of an immature GFP expressing neuron that was imaged every five minutes over the course of an hour. Younger neurons are highly dynamic showing extensive branch growth and branch retraction.

A zoom view of this GFP expressing neuron shows the constant addition and retraction of dendritic filopodia that is typical of developing neurons in this system. The transparency of the albino tadpole and the accessibility of its brain makes this model system ideally suited for visualizing neuronal growth and intracellular events within a live organism. Single cell electroporation allows us to visualize the growth of a single neuron and to perform cell autonomous manipulations within an otherwise unaltered brain.

So we've just shown you how to conduct single cell electroporation in zap lavis tat poles, although we are focused here on electroporation in TA poles. This technique has been used in other organisms and has also been used in hippocampal slices and dissociated cell cultures. When doing this procedure, it's important to ensure you constantly modify your pipette tips and stimulation parameters to maintain high yield.

So that's it. Thanks for watching and good luck with your experiments.