In this video, we're going to make a lab excursion and we're going to visit
Semihcan Sermet, who is  a Ph.D. student in my laboratory.
He's going to show you how to make whole-cell recordings
from brain slices.
Hi, my name is Semihcan Sermet.
In this video I will explain how to make whole-cell recordings
from brain slices.
I will explain the process in four parts.
First we will prepare the brain slices, then we image the neurons,
we prepare our recording electrodes, and finally we perform
whole-cell patch-clamp recordings.
Our first step is preparing the brain slices.
We first anesthetize the mouse, carefully extract the brain,
and then place it  in ice-cold slicing solution
containing, in millimolars, 87 sodium chloride,
25 sodium bicarbonate, 25 D-glucose,
2.5 potassium chloride, 1.25 sodium phosphate,
0.5 calcium chloride, 7 magnesium chloride,
and 75 sucrose which is aerated with 95% oxygen
and 5% carbon dioxide.
This solution is not a normal artificial cerebrospinal fluid.
It is modified to keep the cells healthy during the cutting process.
First, we extract the brain  from the mouse and place it
in the ice-cold sucrose-based slicing solution.
This solutions helps  to keep neurons in a good condition
during the slicing process.
The we chop off the cerebellum.
Since we will be recording [in] neocortex, we do not need [that brain] region.
Take the brain and carefully transfer it onto the platform
which will be going  into the slicing chamber.
The brain is fixed onto the platform with glue.
Then we take the platform with the brain and place it
into the chamber filled  with cold slicing solution.
The solution is bubbling with 95% oxygen and 5% carbon dioxide
to keep a balanced pH, which is 7.3.
We slice the brain with the vibrotome.
It has a sharp blade which vibrates at 85 Hertz and advances with a speed
of 0.09 millimeters per second.
Slices are 300 micrometers thick.
When the blade reaches the end of the brain tissue,
we gently push the slice towards the blade and separate it from the rest of the brain.
Then we take the brain slice with the Pasteur pipette
and transfer it to a chamber
filled with 35° Celsius warm sucrose slicing solution.
We keep the slices here about 30 minutes.
During the slicing, the vibrating blade damages cells in the superficial parts
of the slices, the parts  next to the blade.
In the warm sucrose slicing solution, the damaged cells tend to float
away from the slice, leaving healthy tissue behind.
We keep the slices incubated at 35° Celsius
for 30 minutes in slicing solution, and then we transfer them
to the room-temperature artificial cerebrospinal fluid,
containing, in millimolars, 125 sodium chloride,
25 sodium bicarbonate, 25 D-glucose,
2.5 potassium chloride, 1.25 sodium phosphate,
2 calcium chloride and one magnesium chloride,
which is aerated with 95% oxygen and 5% carbon dioxide.
Then we select the slice and place it in the microscope,
superfused with artificial cerebrospinal fluid at 35 Celsius.
We take our brain slice  from the chamber filled with ACSF,
bubbling with carbogen, and place it into the recording chamber.
The chamber is superfused with ACSF.
Then we fix the brain slice with the grid
and then start visualizing  with the microscope.
First we look at the lower magnification, 4X, to see the brain region
we would like to record from.
Then we switch to the  higher magnification,
60X, to visualize the cells.
In this case, we use a high-contrast infrared video microscopy.
It is important to find a region with healthy cells.
In order to do that, we [look at] the slice vial,
focusing up and down, until we find a good cell to record.
Now we prepare our patch-clamp recording electrodes.
We take a borosilicate glass capillary and place it into a pipette puller.
The electric [element covers] the middle of the pipette
and delivers heat to melt the glass.
The pipette is fixed at both ends and force is applied to pull the electrode.
With enough heat,  the middle of the pipette
begins to melt and the pulling force causes lengthening and thinning
of the glass capillary.
After several rounds of heating and pulling, we obtain pipettes
with a tip size of approximately  one micrometer.
Before we start the whole-cell recording, we fill our glass pipette
with the intracellular solution, which contains, in millimolars,
135 potassium gluconate,  four potassium chloride,
four magnesium ATP, ten sodium phosphocreatine,
0.3 sodium GTP and ten HEPES,
which has a pH of 7.3
and osmolarity  of 280 milliosmol per liter.
We also add Alexa-594, red fluorophore for further imaging.
In order to have a better understanding of the whole-cell recording process,
we can first look  at these schematic examples.
First, before putting our pipette into the bath
[and the tissue], we apply positive pressure in our pipette
so the intracellular solution, shown as blue, can flow out
and then the tip stays clean.
We apply a test voltage and observe the current flow
through the pipette.
Then when we touch the cell, we remove the pressure
and suck gently.
Then a membrane attaches  to the pipette tip
and then forms a tight electrical seal.
At this time, only a very small amount of current passes through the pipette tip.
This is now a so-called cell-attached patch-clamp configuration.
Then, we do further brief suction pulses
to break the membrane in order to establish
the whole-cell patch-clamp configuration.
Then we break the membrane and we see a current flow like this
charging the membrane because of the cell capacitance.
We fill our glass pipette with internal solution
and place it on the head stage,
which has the silver chloride  recording [electrode] on it.
The reason for the internal solution looking purple is that
it contains a red fluorophore called Alexa-594 to allow fluorescence imaging.
Before we move the pipette into the ACSF bath solution
we apply positive pressure on it to keep the tip clean
by always having an outward flow of solution.
First we find our pipette  under low magnification
and then we switch to the higher magnification
which we were using to visualize the cells.
It is important to move the pipette around with great care,
making sure not to damage the brain slice.
We bring the pipette down towards the brain slice,
near to the location of the neuron we would like to record.
When the pipette touches the brain slice surface,
we can see the tissue getting pushed away from the tip.
That is because of the positive pressure around 15 millibars
we apply to the inside of the pipette, causing an outward flow
from the pipette tip.
Then we reach our cell step by step.
First we focus deeper and then we go down with our pipette.
When we reach the cell, we touch the cell carefully
and move the pipette slightly towards the cell body.
Now we see a small dimple where the tip of the pipette
touches the cell body.
Then we remove  the pressure and suck gently
to establish a gigaseal.
When we have a gigaseal, we do further brief suction pulses
to break the membrane in order to establish
the whole-cell patch-clamp configuration.
Sometimes we have to continue the suction pulses
to improve the opening, reducing the [access] resistance
between the electrode and the inside of the cell.
Then we switch to the current clamp mode to measure the membrane potential.
We can then start injecting current into the cell.
Here we apply currents for 500 milliseconds.
First we apply hyperpolarizing current, then we increase the current
and depolarize the cell and see action potentials.
The more we increase the current, the more action potentials we see.
In this case, we are recording from a neocortical layer V pyramidal cell.
We can zoom in and look at the action potential waveform.
We can see that action potential [half-life] in this cell
is around two milliseconds.
Here on the left we see the two photo-microscopic image
of the cell we were recording from.
On the right, we see the membrane potential traces
of the same cell responding to the injected currents.
In the bottom trace, which is here, we see that the cell hyperpolarizes
and in the top trace,  we can see here,
the cell depolarizes  and fires action potentials.
So we've seen that the mammalian brain can be cut into thin slices
and the neurons remain viable.
We can visualize the neurons
through a high-contrast  infrared video microscopy,
allowing in vitro whole-cell patch-clamp recordings of membrane potential.