Electrophysiology
As the Italian physician, Luigi Galvani, discovered in 1790 while he was working on frogs, the nervous
system uses electrical activity to perform its functions. This biological activity is studied and recorded by
the science of electrophysiology. Electrophysiology uses a wide range of techniques that can be grouped
under two headings: invasive and non-invasive.
In this topic, I will concentrate on the invasive techniques we use, while the non-invasive techniques will be
covered by Dr Giampietro elsewhere on this module. This topic is divided into four parts: in the first part, I
will briefly introduce the types of biological electrical activity; in the second part, extracellular recording;
third part, intracellular recording; and finally, single channel recordings.
The biological electrical activity that we investigate in electrophysiology comes in a number of forms. There
are large voltages that are generated by animals, such as electric eels or electric rays, which is used to stun
prey, which can reach up to 700 volts. Then there is the negative resting membrane potential of about -70
millivolts, comparing the outside of both cells to the inside, and this is the membrane potential that most
neurons have. There are also post-synaptic potentials, which are small variable changes in the membrane
potential and that are in the range of one to 40 millivolts. And finally, there are the action potentials, which
reach 100 millivolts, and they are large, fast and occur in an all-or-none fashion. Over the next few screens,
I'll look at each of these in turn.
So first, let's look how electric fish, such as electric eels (Electrophorus) and rays (Torpedo) produce
electricity. These fish possess specialist cells called electroplaques, which have membrane sodiumpotassium exchanges. At rest, sodium-potassium exchanges use ATP to alter the ions distribution across
the membrane. This generates a potential difference across the electroplaque. When the fish wants to give
an electric shock, the electroplaques are activated by a nerve. The nerve releases acetylcholine onto
nicotinic-type acetylcholine receptors that are ligand-gated ion channels. These channels allow the flow of
sodium ions into the electroplaque, depolarising the electroplaque and generating a brief potential change
of around 0.12 volts, or 120 millivolts.
Electroplaques are stacked in their thousands in electric organs and the sum of these can generate
hundreds of volts. Small voltages are used to detect prey, while larger ones - up to 700 volts, as I said
earlier - can stun. Similar to electroplaques, neurons also have sodium-potassium exchanges in their
plasma membrane. These, as before, generate high cytosolic potassium and low cytosolic sodium, using
ATP as their energy source. In neurons, chloride ions are co-transported out of the cell using the potassium
gradient as energy. There are many immobile negative ions, such as phosphate groups, found on proteins
inside neurons. Overall, these all generate a resting membrane potential of between -60 and -80 millivolts
inside the neuron.
Now let's take a closer look at the resting membrane potential, or Vm rest. The presence of the potential
and concentration differences across the neuron's plasma membrane mean that there's a range of
directions of movement for the different ions through their respective channels. For example, sodium ions
move into the neuron, as both their concentration gradient and electrical gradients direct them inwards.
Conversely, the concentration for gradient for potassium ions makes them move out of the neuron,
despite the inward electrical gradient of about -70 millivolts. Chloride ions move into the neuron down
their concentration gradient, despite the electrical gradient of -70 millivolts, which would tend to keep
them out. And finally, calcium ions move into the neuron down what is the highest concentration gradient
of up to 10,000 to one, and also down the electrical gradient, which is into the neuron.
Next, let's look at post-synaptic potentials. The ion gradients in a neuron allow both depolarising and
hyperpolarising potentials. Excitatory post-synaptic potentials, or EPSPs, are generated by the activation of
ion channels that depolarise neurons, usually by transmitters that activate channels permeant to positive
ions: sodium, potassium, and calcium. For example, when glutamate acts on ionotropic receptors, this
activates sodium and calcium to flow into the cell, as shown in the picture on the left.
On the other hand, inhibitory post-synaptic potentials, or IPSPs, are generated by the activation of ion
channels that hyperpolarise neurons, usually by neurotransmitters that activate channels permeant to
negative ions, such as chloride - for example, when GABA acts on GABA A receptors and allows chloride
into the neuron, as shown on the picture on the right.
Both EPSPs and IPSPs are graded in amplitude due to the concentration of the neurotransmitter, as well as
time - the length of time the neurotransmitter is in the synaptic cleft. And both EPSPs and IPSPs are
additive, but decay in amplitude as they move around the neuron.
Now, let's look at the fourth item in the list I gave you earlier, and this is the action potential. Action
potentials require activation of voltage-gated sodium and voltage-gated potassium channels. At resting
membrane potential, both sodium and potassium channels are closed in their resting state, as shown at
point A. A depolarisation leads to the opening of the voltage-gated sodium channels, which lead to further
depolarisation, and further voltage- gated sodium channels opening in an all-or-none way at B. At the peak
of the action potential, C on the diagram, sodium channels close and go into their inactivated state, and
the potassium channels start to open. Next, potassium ions move out and hyperpolarise the cell back
around the resting membrane potential, and this is shown in D. At the resting membrane potential, both
sodium and potassium channels return to the closed resting state at E, where we started.
Action potentials are all-or-none in terms of amplitude, around 100 millivolts. They are brief, lasting only
between 0.5 and two milliseconds, and can travel fast at long distances in excitable tissues without
changing amplitude. Here is a summary table of the biological activity covering the main points.
In this part of the topic, I will cover the electrophysiological techniques we use to record electrical activity
of nervous tissue. So, I'll talk about extracellular, intracellular and single channel recording in that order.
Extracellular recording, which we abbreviate to ER includes five recording types: field potentials, whole
nerve activity, multi-unit activity, single unit activity, and finally, the use of multi-electrode arrays, or MEAs.
Intracellular recording, or IR, is used to record activity within single cells and involves the use of either
sharp electrodes or patch suction electrodes. Single channel recording, or SCR, is used for recording the
activity of single ion channels and uses patch clamp-type electrodes. We will look at each of these in turn,
starting with extracellular recordings.
So, let's look in more detail at extracellular recording. As I indicated earlier, there are five versions of
extracellular recordings that we're going to show you: firstly, field potentials; then, whole nerve
recordings; multicellular recordings, or multi-unit recordings; single unit recordings; and MEAs. In each
case, the electrode is outside but very close to the neuron. Under these conditions, electrodes pick up only
field potentials and low frequency filtered action potentials. You cannot record membrane potential, or
VM rest, or post-synaptic potentials using these techniques. Here, I'm going to illustrate field potentials
recorded from a rat's brain slice.
This is a slice of the rat's brain. It is in fact a slice of the hippocampus area, and we have placed three
electrodes into the tissue but not into the cells. One is a stimulating electrode in red, and the two
recording electrodes are shown in blue. When we stimulate the stimulating electrode, this activates the
Schaffer collaterals and causes neurotransmitter release onto the Purkinje neurons in the area called CA1,
which is where the two electrodes in blue are positioned. The signal from the blue recording electrode on
the left show something called an FEPSP, and this is a field excitatory post-synaptic potential, and this is
the effect of glutamate receptors opening and causing a depolarisation of the neuron at this point. The
other electrode, which is on the right, this is recording the action potential known as the somatic
population spike, so it's in fact the sum of many action potentials of CA1 cells in this area.
So here, we can record both the incoming EPSP and the resultant action potential generated by it in the
dendrites of the neuron. In a similar way to the previous example, the compound action potential of a
whole nerve can be recorded extracellularly. In the example shown here, the action potential of the sciatic
nerve of the frog is being recorded. Here you can see that the nerve has been isolated and placed on
platinum wires at one end for stimulating and the other for recording. The outer chambers are sealed with
silicone grease - SG in the figure - and filled with olive oil to electrically isolate the nerves. Only the action
potential traveling down the individual axons will be recorded.
You can see that increasing the intensity of the stimulation from 1.2 volts to five volts in the graph
increases the compound action potential size; because the stronger the stimulation, the more axons are
recruited to fire action potentials, meaning that the compound action potential gets larger and eventually
all the axons will be recruited and the compound action potential will be maximal.
I have recorded two videos for you: a recreation of Galvani's experiment on frogs, where l stimulate the
nerve to cause the frog leg to twitch; the isolation of the whole nerve from a frog and recording of the
action potential, as I highlighted to in the previous slide. Similar compound action potentials can be
recorded from rats- in this case, the isolated vagus nerve of one.
Here, you can see the different types of axons from A-beta, A-delta and C fibres. These different axons can
be separated by the intensity of the stimulus and their conduction velocity - AB fibres being the fastest and
C fibres the slowest.
Now, I'll move onto multi-unit extracellular recordings, and these can be done in vivo. Here, an electrode is
placed in an anaesthetised rat brain; the recording area is called the lateral geniculate nucleus or LGN,
which is part of the visual pathway. With the electrode in place, a flash of light is used to activate the
neurons. The largest action potentials you see here are from one neuron very close to the electrode, but
the smaller action potentials here are from a cell that is further away from the electrode. Therefore, you
are simultaneously recording from two different cells.
Here is an example of a single unit recordings, this time done in a human brain, and at the medial temporal
lobe. This patient gave consent for this and was having a deep brain stimulation electrode implanted, so
the recording wasn't done without good reason. As we see at stimulus one, we have a picture of Halle
Berry and we can see the neuron that was being recorded from the subject responded with action
potentials. Stimulus 2 is a different picture of Halle Berry and the neuron still responds. Stimulus 3 is a very
different picture of Halle Berry, but the neuron still responds. They even put a picture of Halle Berry's
name up and it responded, however, it did not respond to a picture of Michelle Pfeiffer. Therefore, this
neuron was called a Halle Berry neuron, and it is an example of association neurons that may exist in the
human brain.
And this is my final example of extracellular recording where we can use multi-electrode arrays, or MEAs.
These electrodes, rather than being sharp pointed things, are actually electrodes that are bedded in the
bottom of this dish shown in the picture. There are, in fact, 64 of them in the bottom of the dish shown on
the first square here, and if we look at those individually, they are inert to the cells - so don't affect the
cells - but they can pick up extracellular electrical activity neurons.
What we can do is actually grow the neurons inside these dishes in an incubator and we can monitor, then,
the electrical activity in a very non-invasive manner. In this slide, I show a picture of an MEA with cells that
are growing on it. These cells are called NG108-15 cells and you can see the cells are growing on the multi-
electrode array and the black circles are where the electrodes are. So, you can see the cells are very happy
in this dish and they're happily growing on the electrodes, so we can record their electrical activity. Week 2
In this slide, I can show you examples of the action potential activity from the MEAs that you've just seen.
On the top left, you can see there is no action potential activity, but just some background noise. In the
middle left, where it says '12 millimolar potassium chloride', this is what's been applied to those cells and
made them depolarised and start to fire action potentials. As I said in the middle trace, you can see two
action potentials, and after a period of time, the action potential activity increases on the bottom trace in
the left. On the right, we've applied a more natural neurotransmitter, ATP. The right top trace just shows
the background activity in control situations. Then, I apply one millimolar ATP in the middle trace and the
action potentials start to fire, and finally, is a trace a few seconds later where there's high levels of action
potential activity. And this shows we can record, in a non- invasive manner, action potential activity from
multi-electrode arrays.
In this slide, I have given you all the main points about the extracellular recordings, particularly comparing
their advantages of the different technique to their disadvantages.
Now we come on to intracellular recordings.
Here I’m going to compare two things: firstly, current-clamp with voltage-clamp, which is the nature of the
recording that is occurring; secondly, I’ll be talking about sharp electrodes versus patch-clamp electrodes
as the nature of the recording electrode used in the different techniques. First of all, I’m going to talk
about using sharp electrodes. Here, a biological preparation is a creature called Ciona intestinalis, which is
a sea squirt, and we isolate the oocytes from it, which we can then put an electrode into, which is actually
penetrating the plasma membrane, so is intracellular recording using a sharp electrode.
So, here is a comparison of sharp electrodes in the two different recording setups - either in current-clamp
on the left or voltage-clamp on the right. Top left, A, shows a single action potential. So, what we've done
here is stimulated the oocyte with a small brief pulse that has caused an action potential to occur. As you
can see, this is in the order of 100 mV. Below that, I have done some different intensities of stimulation.
You can see the red line, the intensity was not big enough to stimulate an action potential, so it's not
present. The green one shows a large action potential and the blue one shows one of the same size, but of
slightly longer duration.
These are recordings in current-clamp, where we're injecting currents and recording the voltage. On the
righthand side, we have the same cell and the same electrode. And in C, we are in voltage-clamp mode.
Here, you can see a single voltage step and the current that results from it, and this voltage step induces
the current that we see when it generates the action potential, but instead of looking the voltage, we're
now looking at the current flow through the membrane that generates that voltage pulse: the action
potential. Similarly, we can apply different voltages as shown in D, each of different colours. If the voltage
is too low, as we saw in B, the green one shows no current is evoked because the voltage was not strong
enough. However, as you can see in the dark blue one, a large current is generated when we use a larger
voltage. As you can see, the current-clamp and voltage-clamp are related to each other. One in currentclamp mode, we are recording the voltage but changing the current, and in voltage-clamp mode, we are
stimulating with voltage and recording the current. Now, we use patch-clamp electrodes. The patch-clamp
electrode does not go into the cell, it goes to the cell's surface and interacts with the membrane and forms
what we call a gigaseal and you can see that occurring in A.
This is known as an on-cell patch, where the glass electrode has made a gigaseal with the cell membrane. If
we're very lucky, we may find a few channels in that action membrane that we've sealed to, and we would
be able to record single channel activity, known as cell-attached patch recording. If we pull the electrode
away at this point after we make the gigaseal, this pulls the patch of membrane off the cell and we can
then have access to the intracellular side of the patch. We can apply drugs there and still record the single
channel activity in B. Another alternative shown in C is where once we've got cell-attached patch, we
actually break through the membrane with the electrode still attached to the whole cell and this goes to a
version called whole-cell clamp, where we have access to the inside of the cell which enables us to record
from all the ion channels in the membrane of that particular cell - this is called whole-cell recording. If we
pull the electrode away at this point, we would then develop an outside-out patch, where we'd pull off a
small patch of membrane - if we're lucky, with one or two channel in it - and there, the outside of the
membrane would be exposed to the outside environment, as would be normal for the cell. One final
variation of the patch-clap technique is a little bit more sophisticated and this involves filling the electrode
with a solution that contains a pore-forming antibiotic. As before, we make a gigaseal as in A, but instead
of moving the electrode or breaking the membrane we just wait until the antibiotic diffuses to the
membrane and forms pores in it by itself. This is called perforated patch. This has a big advantage over
whole cell voltage clamp under some conditions in that the contents of the cell are not dialysed by the
electrode solution and you don't lose things from inside the cell because the holes formed by the
antibiotics are small enough to prevent this happening- but we still get electrical activity recordings. In this
slide, we see using whole-cell or patch-clamp recording electrodes, on the left, a current clamp recording
where action potentials generated in rat retinal ganglion cells are recorded. As before, we're using current
ejection to record the voltage. On the right, we have the voltage-clamp situation where I'm using voltages
to generate the currents that evoke voltage change. This time, I'm using a stimulus that is acid or pH
change, where we're evoking acid-sensing ion channels – ASICs - to generate a current when they open. At
the top, on the right (number A), you can see that as we change the membrane potential, the current size
changes and at the bottom (B1 and 2), as we increase the acidity of the application of the compounds, that
increases the size of the current. So, as with the oocytes, we can use whole-cell or patch-clamp recording
electrodes to record in current-clamp and voltage-clamp mode. Here again is whole-cell patch clamp in
current clamp mode. Here's an example showing that you can use electrophysiology with another
biological technique. In this case, simultaneously, we are recording the intracellular calcium levels in this
neuron. So, we have, at the top of each section, a trace that records the concentration of intracellular
calcium, and at the bottom, the action potentials that that particular neuron is generating. So, if we take
the top left section, you can see that the action potentials evoked number 17 and that produced a
relatively small rise in intracellular calcium inside the same cell. Top right, we've increased the number of
action potentials to 28, the calcium concentration increases because more calcium channels are opened
each time an action potential occurs. And then, bottom left and bottom right, we are sequentially
increasing to 36 and 48 - the number of action potential generated - and this correlates with the increase
in calcium rise that we get, showing that calcium entry during an action potential is related to the number
of action potentials that are opened or activated in that cell. And the final technique I'm going to talk
about, is single channel recording. In this example, this is a cell-attached patch where we have two single
channels in the patch.
The line on the bottom of each of these traces, C, is the line at which both channels are closed. O1 is the
level at which one channel is open, and O2 is the level at which both channels are open. As you see, it looks
very step-like. So, no channels can be open at C, one channel can be opened at O1 and two channels can
be opened at O2. It's probably easier if I start with a lower trace first. You'll notice on the left I have a sign
saying 'Vp- 10', and this means we've just depolarised the cell by a small amount, 10 millivolts. As you can
see, the single channel conductance is quite low and the time the channel is open is also low. Indeed,
towards the right of that trace, the channel almost closed most of the time. If we move up to the middle
trace, Vp -20, you can see now, most of the time, one or other or both of the channels are open, so the
activity of the channel has increased but so has the single-channel current- which has got larger. The top
trace, Vp -30, is even more depolarised. Here you can see, the channel is open even more often, either one
or both of them, and the size of the current has got bigger. This means you can record two important
things about the channel: you can calculate its single channel conductance and you can measure its voltage
dependence due to the change in voltage that the channels are recorded at. Now, we're going into a
situation where we can use patch-clamp in vivo. This is a very recent technique or application of the
technique. So here, we have an animal that is held in a stereotactic and anaesthetised to keep it still.
Again, we put the electrode into the brain, but this time, as it shows in A in search mode, we are applying a
positive pressure to the back of electrode- we're actually forcing air into the back of the electrode. This
pushes out the fluid out the front and this means it keeps the path clear so the electrode can be moved
through the tissue by a manipulator. Once the electrode hits a cell, we see a change in the pulse and it
becomes irregular due to the heartbeat frequencies that you pick up when you touch the cell, and we turn
the pressure off so it goes to neutral. We then allow the cell to go to cell attach mode, as before, and the
electrode seal, giga-ohm seal, is formed. After a small period, in D, we then apply a very short sharp
suction - this time, a negative one - so we're sucking the electrode solution up which destroys the patch of
membrane that's made under the seal, and this allows us to go into the whole-cell mode as before. So,
we're basically recreating that previous slide but in the brain of a living animal. This enables us to record
quite sophisticated responses shown in the section on the right, where the whiskers of the mouse have
been stimulated by moving them, and you can see it evokes action potentials and EPSPs in the cells in the
brain that are associated with that whisker response. Here, I've given you a table on the applications of the
different techniques. I've separated them into the columns: which one can be used in humans in vivo,
which ones can be used in non-humans in vivo, and finally, which techniques can be used in vitro. The only
technique that's used in humans in vivo is extracellular recording, but only part of an existing treatment.
For example, if somebody was going in for deep brain stimulation electrode implantation, you could then with the patient’s permission - do some recordings as the electrodes were being implanted. In nonhumans, as are shown, both extracellular recordings can be used, either implanted in live awake animals or
in anaesthetised animals. Intracellular recordings can only really be done with anaesthetised animals
because they need to be kept still. And finally, single channel recording can again only be done in
anaesthetised animals. In vitro, all three techniques can be used, and sometimes, in some cases, human
tissue can be used under these conditions as well, but obviously with the correct permissions from the
patients involved.
Here, we have a summary slide for all the intracellular recording techniques that I've described, giving you
both the advantages and some of the disadvantages of each type. So hopefully, what I've shown you is that
electrophysiology can record the electrical activity of whole brain tissue, a single neuron or even a single
ion channel in the brain. Electrophysiology is a dynamic, functional, SI unit-based, real-time, hi-fidelity and
high temporal resolution approach. Many electrophysiological approaches can be used vivo.
Electrophysiology can be used simultaneously or in conjunction with optical, molecular, biochemical and
pharmacological techniques. I think we're not going to understand the nervous system without it.
Immunohistochemistry
Why examine tissues and cells? Well, by doing this, we can study tissue anatomy, and cytoarchitecture. We
can study the distribution of proteins within the tissues and, most importantly, we can study and
understand pathological changes associated with disease. So histological study of tissue is essential for
clinical diagnostic neuropathology and also basic and translational neuroscience research. Let us next think
about tissue sources. For this, we can use animal models, post-mortem donor tissue, pathology samples
and or even surplus surgery material.
Let us first think about the use of animal models and their advantages and disadvantages. One advantage
of animal models is that they enable us to examine tissue throughout the course of disease from early to
end stages. Although some animal models occur naturally, most are generated based on specific genetic
defects that have been identified in humans, so we can use animal models to assess the effect of these
specific mutations. We can also use these models to analyse whether therapeutic strategies have any
effect. However, we must also remember that there are limitations of using animal models for
translational research. This is because animal models often do not always fully recapitulate the human
disease. Therefore, we must always interpret the results of studies that use models organisms carefully.
There is also the need to carefully consider the welfare of animals that are used in research, so there are
always ethical concerns to be taken into account when undertaking any type of animal research. So, what
about using human tissue? Some of the most valuable research comes from the study of donated human
tissues. Most donations are done post-mortem with full consent given by the donor or members of the
donor’s family. In some cases, if consent has been given, pathology samples or surplus surgery material can
be used for research.
What are the advantage of post mortem donor tissue? Well, using human tissue can reduce the need for
animal research. It is arguably a better material to study human diseases or diseases for which no animal
model exists. However, the use of human tissue is carefully regulated. Also, as you may imagine, most
post-mortem tissue that is available generally comes from the end stage of the disease and there is little
availability of such tissue from early disease stages. Now that we’ve obtained tissue for our research
project we have to prepare our sample so that it can be used effectively. The first thing to remember is
that once tissue is removed from a living organism or a deceased donor, the irreversible processes of
autolysis and necrosis will begin, leading to cellular damage. Also, tissues from post-mortem donations are
more likely to have signs of cellular damage than samples obtained from living donors. So, when we
prepare tissue for histology our aim is to preserve this tissue in as life-like a manner as possible and to
prevent irreversible cell or tissue destruction, which can be caused by the presence of pathogens. There
are two common methods for tissue preservation: chemical fixation and cryopreservation, and we shall
look at these in turn.
There are many chemicals that act as fixatives. The choice depends upon what is required in experimental
design, as no fixative is ideal for all applications. Fixatives that are best at preserving cellular ultrastructure,
such as glutaraldehyde, can interfere with the subsequent staining processes, while other chemical
fixatives, such as acetic acid and methanol, can produce some good staining results but give inferior
cellular and tissue morphology. Most applications use formaldehyde, which gives best balance between
optimal morphology and stain quality. So what do chemical fixatives do? Well they stabilise proteins and
other macromolecules which are enmeshed amongst these proteins, such as carbohydrates. Cross-linking
fixatives - for example, formaldehyde and glutaraldehyde - create covalent bonds between proteins in the
tissue. They lead to good preservation of morphology as the process anchors proteins relative to each
other inside the cells and between the cells. Protein tertiary structure is maintained, but the fixation
process tends to be slow. Precipitating fixatives, such as ethanol and methanol, disrupt hydrophobic bonds
between proteins causing them to irreversibly precipitate. While this can be useful for some techniques,
this type of fixation is much less suitable for antibody-based techniques – what we call immunostaining.
Now let’s consider how to fix tissue. We can fix the first tissue by simply immersing it in a chemical fixative
as a liquid. The fixative, in solution, will diffuse through the tissue over time. While this is more applicable
for small, already dissected samples, larger samples can easily be placed into the same tissue but fixed with
agitation – this means stirring. Alternatively, we can inject fixative as a fluid directly into the circulatory
system. This is called perfusion. Perfusion requires an intact circulatory system but can deliver the fixative
very quickly throughout the organ or entire experimental animal via the blood vessels. This method is
always going to be preferred, if possible, because it gives superior preservation. Several factors will affect
the quality of tissue fixation.
Changes in pH, for example, will affect the reactivity of the fixative. The rate of the fixative diffusion will
determine the length of incubation in the fixative. Formalin, for example, will penetrate tissue at
approximately one millimetre per hour. The size of the specimen is also very important to consider. So
where possible, samples are dissected to no larger than five millimetre cubes as this facilitates optimal
fixation.
Temperature also plays a role. Fixation at four degrees centigrade will retard degenerative changes but will
also reduce the penetration rate to the fixative. In contrast, room temperature fixation will accelerate
fixative penetration, but also potential degenerative changes. We will now move on to consider
cryopreservation. Cryopreservation is the preservation of tissue structure and components by freezing
them rapidly without fixation, although, in many cases you can fix the tissue and then freeze it.
Using cryopreservation, we can stop tissue degradation by rapidly cooling sample with dry ice or liquid
nitrogen – what we call ‘snap-freezing’. This method is very rapid, and unlike chemical fixatives, does not
modify protein structure. However, the morphology is relatively poorly preserved, as ice crystals form
during the process can damage cellular structure. This is particularly true if the snap-freezing is not carried
out correctly.
However, if the sample is rapidly cooled to below -70 using liquid nitrogen, the liquid water is converted to
vitreous water without going through the crystalline phase, and this will minimise damage to the tissue, or
what we call the ice crystal artefact. Cryopreservation does not permanently fix the tissue, thus, over time
degradation will occur and will progress rapidly if the sample is not stored at the appropriate temperature most often, in -80 freezers. The next step in tissue processing for histology is embedding.
Chemical fixation will not harden tissues sufficiently to allow thin sections to be cut - for example, at six
microns thin. So the sample is embedded in a solid medium that will give extra support for tissue structure
and also give the sample sufficient rigidity to enable cutting of thin sections. There are many embedding
mediums available, and the choice depends upon experimental requirements. For example, if we wish to
study neural architecture, we would use harder embedding media, such as plastic resins, since we can cut
much thinner – for example, one to two micron sections – using this method.
However, paraffin wax is the most common embedding media in histology laboratories. Essentially, the
sample is impregnated with liquid paraffin wax which hardens and forms a matrix in and around the
sample, preventing tissue distortion. It is a very versatile embedding medium and allows for sections of
different thicknesses to be cut. Softer embedding media, such as agarose and gelatine, are suitable for
production of much thicker sections.
Cryopreserved tissue is already hard enough to allow sectioning, but small or irregularly shaped samples
can be further supported with media that freezes at the same density as most soft tissues. Commercial,
water soluble, embedding media – such as OCT - are viscous at room temperature, but upon freezing,
solidify and act as a support medium for the tissue, and also allows for the tissue to be handled for a short
time without risk of thawing.
We are now going to discuss the processing of paraffin wax embedding and the stages involved. Paraffin
wax, with a 56 degrees to 60 degrees melting point, is used. It's a widely used embedding medium. It is
sufficiently hard to support tissues, yet soft enough to allow sections to be cut using a microtome blade.
However, it is not soluble with water, thus tissues need to be processed before it can be embedded in
paraffin wax.
There are many variations in times and chemicals used, but the sequence of four events and the rationale
are the same. Initially, you have to dehydrate your tissue – that is, remove all of the water from it –
because water is not miscible with molten paraffin wax. So it must be completely removed by sequential
immersions in increasing concentrations of alcohol, commonly 70 per cent, 90 per cent and 100 per cent
alcohol.
The number of changes in each of those is up to each laboratory. Once the tissue has been dehydrated, it
needs to be cleared of alcohol because the alcohol is not miscible with molten paraffin wax either. We
have to remove it completely by a solvent that is miscible in both alcohol and paraffin wax: for example,
xylene or Histo-Clear.
Once the tissue has been cleared, we need to replace the xylene with molten wax. This is called infiltration.
Finally, once the tissue has been infiltrated with molten wax, we have to embed the tissue in a block of
paraffin wax. The tissues are oriented in metal moulds containing fresh molten paraffin wax, and then they
are allowed to cool. Upon cooling to four degrees C, the wax blocks are easily removed from the metal
mould, ready for sectioning or storage. In the next part of this lecture, Carl Hobbs, the Histology and
Imaging Manager at the Wolfson Centre for Age-Related Diseases at KCL, will tell you about the machinery
we use in the laboratory to process and cut tissue sections.
We can process the tissues by hand, but more commonly - and while you have many, many tissues to
process - you will use a tissue processor. There are several types of them. The two common ones are a dip
and dunk machine, for example, the one shown here, which has 12 stations in it. The specimens are placed
in a basket and loaded onto the processor. After predetermined times in each station with agitation the
basket is automatically transferred from graded alcohols to either xylene or Histo-Clear, and then to
melted paraffin wax before being removed for embedding in fresh molten paraffin wax. This is also called a
dip-and-dunk processor. This is an example of an enclosed tissue processor.
Tissues are loaded into a chamber and the processing reagents are sequentially pumped in and out under
vacuum to increase processing efficiency. This can be used as a high throughput processor. Once tissues
have been infiltrated with paraffin wax, we then need to embed them. Commonly, you will use a paraffin
wax embedding station, as shown here. So once they've been processed on the machine, for example, or
by hand, the specimens are transferred to a molten wax tray on the embedding station. Tissues are then
placed in metal moulds filled with molten wax and oriented optimally before placing on the cold plate to
set the wax. This image shows an operator embedding a piece of tissue in molten wax which is placed in a
metal mould. The in-set shows a high power of the set wax block being taken out of the metal mould. In
this processing system, plastic cassettes are used to enclose each piece of tissue while processing. When
embedding, the lids are removed and discarded. The tissues are then placed in the metal moulds, as
mentioned earlier, and the plastic base of the cassette is placed on top. This plastic cassette will also have
a unique number written on it for that specimen. Once set, the block plus plastic base is removed. This
plastic base is rigid and can be fixed into the vice of a microtome so that the block is firmly held in place
while sectioning the block of tissue.
Once we have mounted our block of wax embedded tissue onto the microtome we need to section it.
Sectioning is the process of cutting thin slices from the sample, which are required for microscopic
examination. There are a number of devices available for sectioning and we will go through them. Most
common is a benchtop rotary microtome. This rotary microtome is used to cut sections from paraffin
embedded tissues, and the thickness of the section conventionally ranges between three and 10 microns.
A sledge microtome can alternatively be used. Unlike any other type of sectioning, paraffin wax sections
can be cut as ribbons, as is seen on the image.
Once sections are cut, they are floated onto water in a bath, maintained at about approximately 40
degrees centigrade. This softens the wax surrounding the tissue sections so it expands allowing the tissue
sections themselves to become flat. Sections may then be separated to be mounted onto slides;
individually or mounted as ribbons.
This slide shows a ribbon of sections floating out on warm water in a water bath. The ribbon of sections
can be seen from which two sections have been detached by hand, by the operator. One of them is being
mounted onto a microscope slide. Another type of microtome with which to cut sections is the vibratome.
This is used for samples embedded in softer media not paraffin wax, such as agarose or gelatine, and 50 to
500 micron sections can be cut from these. The sections are cut by a blade vibrating at high lateral speed
but advancing slowly through the sample. Sections are collected and are stained as free floating sections or
can be mounted onto slides. This slide shows images of sections that have been cut by the vibratome, so
they're vibratome sections.
On the left hand side, the sections have been cut but, they have not yet been stained. The sections in the
dish on the right-hand side are sagittal sections of rat brain, stained using cresyl fast violet, and are ready
to mount onto slides and cover slip for microscopical examination. The image here is of a cryostat. A
cryostat is used for sectioning of snap-frozen tissues. Sections are cut using a refrigerated cabinet
maintained at approximately -20 degrees centigrade, and it contains a rotary microtome. Cryostat cut
sections are usually in the range of eight to 50 microns.
Cut sections are usually mounted onto slides and can be immediately fixed if the tissue has been frozen as
fresh tissue, or of the sections can be stored for later use. Thus, you will have to store them at either -20,
for short term, or -80, for long term storage. Because the tissue is snap-frozen, the vitreous water is hard
enough to act as the embedding medium for cutting sections. However, one may use an embedding
medium, if required. This is particularly useful for delicate or very small samples. One uses a specialised
embedding medium, such as the Optimal Cutting Compound - OCT - which freezes at the same density as
most soft tissues. This is an example of another type of microtome. This is a sliding microtome and it's
commonly used to section frozen samples without the need for a relatively more expensive cryostat. This
machine is fitted to the bench top and the tissue is kept frozen by blasting it with CO2 gas, for example, or
cardice - that is, solid CO2. The sliding microtome can produce 15 to 200 micron sections which are stained
as free floating sections.
As you have already learned, microscopic examination of tissues requires a cutting of thin sections from
the tissue so that light can pass through the section. However, excepting where sections are cut from
tissues that have been impregnated using the Golgi method, for example, as will be described shortly, the
components of the tissue are mostly invisible, so many techniques have been devised that use dyes, heavy
metals, or fluorochromes to induce colour or contrast in these components.
Additionally, optical contrast of sections can be enhanced using specialist optical techniques, such as phase
contrast and differential interference contrast microscopy. But, these are most useful for studying living
cells in tissue culture and will not be considered further here. Antibodies raised against specific
components of tissues or cells can be used to specifically identify and locate such components, for example
proteins, in sections, and these can then be visualised by immunostaining techniques which will be
discussed in part 4. However, here in part 3, I will discuss dye staining and metal impregnation techniques.
Dye stains are some of the oldest histological techniques available to scientists. Long ago researchers
discovered that with specific formulations, different dyes have strong affinities to particular tissue or
cellular components. It is important to remember that these dye staining techniques, unlike the
immunotechniques that will be discussed later, are selective but not specific. For example, stains such as
crystal violet can be used to visualise the presence of Nissl substance that is found in neurons.
Therefore, this stain will react with all neurons in the brain section making it an excellent choice for
studying the cellular patterns within the specific brain area. However, this technique is not a good choice
for studying the detailed morphology of a neuron, or specific types of neurons. Different histological
techniques can be used to investigate specific features of neural structure and function.
Shown, here, is a typical neuron, with an axon that is myelinated. The axon is a long process that you can
see extending from the neuronal body towards the bottom of the slide and it is surrounded by a substance
that insulates and protects it. This substance is called myelin and it is made by cells called
oligodendrocytes, a type of glial cells in the central nervous system. This myelin sheath allows electric
impulses to travel rapidly along the axon. Let us explore how histology can be used to visualise cells and
their processes in the nervous system using a neuron as an example.
Nissl staining is an example of dye staining achieved mainly by ionic interaction between the dye and the
tissue component. The Nissl staining method was developed by the German pathologist Franz Nissl at the
end of the 19th century and is still widely used today to identify neurons. Neuronal function requires the
continuous synthesis of large quantities of proteins such as neurotransmitters. These are the chemical
messengers that enables signals to be transmitted from one neuron to another neuron across the synapse,
the small gap between the neurons.
Neuronal cell bodies house the machinery for protein production called the rough endoplasmic reticulum,
or RER. Attached to the RER are ribosomes, the structures where RNA is bound and used as template to
assemble many proteins needed by the cell. The RER forms large granular bodies and RNA in these
granules can be visualised with basic aniline dyes for example crystal violet. Aniline dyes are synthetic dyes
originally made from the analyne that can be obtained from cold tar. Basic or cationic dyes in aqueous
solution can ionize to form net positively charged dye molecules or cations. That will combine with net
negatively charged components in the cell. That is acidic components or anions. In particular, DNA and
RNA.
In neurons, because they are actively synthesizing many proteins, aniline dyes but also stain Nissl bodies
that are rich with RNA molecules. These will stain more intensely the other cationic components that have
lower densities of charge. In the image, note that the nuclei of the two neurons are very pale staining. This
is because the DNA is enwound to allow RNA to be transcribed or made. On the other hand, the cytoplasm
of the neuron is intensely stained, because this is where the RER is found. The other less intensely stained
components are nuclei of other cells that are not so actively synthesizing proteins.
In addition, axons and fine structures such as small dendrites will not be visible with this stain. As an aside,
the terms acidic and basic are histological terms that refer to the negative or positive charge of a dye in a
clear solution. Anionic or cationic will perhaps be better terms to use. Let me give you two examples. You
don't have to know but either of these studies in any detail. They are just being used to give you a flavour
of how Nissl staining can be used for study of neural lost following injury or disease. Let's first look at the
set images on the left-hand side.
The images in the bottom panels are from an ischemic gerbil brain, where you can see a much lighter
purple stain, indicated by black arrows, when compared to the images in the panels above, which are from
a healthy brain. This indicates the loss of hippocampal CA1 neurons, following ischemic damage. Meaning
damage caused by the restriction in blood supply to tissue.
The images on the righthand side show you how abnormal growth and development of the cerebral cortex
is easily detected with Nissl stain in a mouse model of lissencephaly, a disease caused by defective
neuronal migration, resulting in lack of development of the brains grooves, and folds, or sulci, and gyri. We
can also identify cell types other than neurons in brain sections using dyes. Luxol Fast Blue is an acidic dye
in solution that is used to visualise central nervous system myelin sheaths in paraffin wax sections. With
this technique myelin is stained a deep blue.
Remember myelin sheaths as I mentioned earlier are formed by oligodendrocytes in the central nervous
system. Composed of cholesterol and glycoproteins, myelin serves to insulate the axons of neurons, which
significantly increases the conduction velocity of action potential. This Luxol Fast Blue technique, may be
used to study loss of myelin in neurodegenerative diseases such as multiple sclerosis.
Two examples of Luxol Fast Blue staining in the central nervous system are shown on this slide. Both are
counterstained with cresyl violet. A counterstain is so called because it either produces a contrasting
background colour to the main stain or provides complementary information. In this instance the cresyl
violet provides additional information by identify Nissl granules. The upper image shows a ventral hole of a
spinal cord where the white matter marinated myelinated tracks run on the periphery in transverse
section. Note the purple staining of the Nissl in multi neurons due to the cresyl violet counter stain.
The lower image shows intense blue staining of the myelin in the white mater in cerebellum in
longtitudinal section. The nuclear cells of the granule cell are pale blue due to cresyl violet counter stain.
So the Luxol Fast Blue stain is useful in a study of myelinated nerve tracts in the central nervous system,
and changes that may occur to myelination. Here are two examples of studies where the staining has been
used.
Again you do not have to know details of the studies. On the left-hand side are images from a study by
Lovato et al, Luxol Fast Blue and cresyl violet stains, we use to effectively demonstrate complete loss of
myelinated axons of a corpus callosum shown by the black arrow. And malformation of the hippocampus,
shown in the box inset, in mice lacking the tumor suppressor neurofibromatosis two gene, which has a role
in brain development. By combining the two stains we can visualise both the neuron of cells bodies and
their myelinated axons, visualising the structure of the entire brain.
The images on the right hand side demonstrate how Luxol fast blue may be used to detect loss of myelin
due to injury. Here we can see the complete loss of myelin in the white matter of the mouse spinal cord
after contusion. Contusion occurs due to traumatic injury and often causes inflammation and bleeding
from blood vessels near the injury site. There are also staining methods that allow visualisation of fine
structures such as dendrites and even dendritic spines.
Remember that neurons receive information from other neurons via the dendritic spines that protrude
from the dendrites and pass on this information via their axon. The Golgi stain, not really a stain, but a
metal precipitation, was developed by Camillo Golgi, and first published in 1783. It was modified by
Santiago Ramon y Cajal to draw neurons and neuronal circuits for the first time, founding the study of
neuroscience.
Basically a small piece of formalin fixed tissue is immersed in potassium chromate, then silver nitrate. The
Golgi methods stained the small, random subsets of neurons in exquisite detail. Also the technique
produces little background staining and the neurons are stained in dark black or brown producing a high
contrast of fine structures such as dendritic spines. This colour is produced by silver salt deposits in the
neuron. However, exactly why only some neurons react is unknown. To illustrate this technique, in the top
image you see a drawing of rodent hippocampus made by Ramon y Cajal, using the Golgi method to
illustrate neuronal network.
The lower image is of a Golgi stained cerebellum section. Part of the Purkinje cell is shown and one can
easily identify primary and secondary processes and also the dendritic spines. Now, some examples. The
Golgi stain is often used to study neuronal morphology. The method can be used on very thick sections of
the brain, up to 500 micrometers, or even whole brains. This means that for some neuron types, we can
visualise the entire neuron and most of their processes in the same section. We can reconstruct the
neuron and see what affects, for example, a neurodegenerative disease may have on its morphology. This
image on the left-hand side is from a paper by Milatovic et al and it shows how this method can be used to
stain sections of the mouse striatum from which tracings of medium spiny neurons can be generated. The
Golgi stain is also often used to quantify the number of dendritic spines that a neuron has. For example,
the image on the right-hand side is from a study by Glantz and Lewis, who use the Golgi method to show a
loss of dendritic spines on the prefrontal pyramidal neurons with patients with schizophrenia.
In this section, we will discuss immunotechniques. Immunostaining, immunodetection, immunolabeling,
are all terms used for a technique that visualises specific molecular targets, mostly proteins in tissues. We
call these targets antigens and visualise them by the use of antibodies raised or made against the antigens.
For example, antibodies can be raised against a protein, the antigen, in a rabbit. Let us assume that this
protein is called NueN. This is a nuclear protein expressed by neurons. The serum from the rabbit will
contain antibodies against NueN, what we call anti-NueN antibodies, and these can be harvested, and
applied to a tissue section. If the NeuN protein is present in a section, then the anti-NueN antibodies in the
serum would bind to it and generate a NueN/anti-NueN complex. I will explain this process in a little more
detail shortly. Once a complex is formed, it may be visualised by subsequently cutting out an enzymatic
reaction on a section. This is called immunohistochemistry.
Alternatively, it may be visualised by using fluorescent dyes. This is called immunofluorescence. The term
immunocytochemistry that you may come across, refers to the process when an enzymatic reaction is used
to visualise antibody antigen complexes on cells grown in a laboratory in a tissue culture dish. You will hear
more about cell culture later in this module.
I should mention that the term immunoflorescence is also used to describe the immunolabeling of cultured
cells by florescence dyes. Immune detection methods take advantage of the interaction between
antibodies and the molecular targeting tissues called antigens. Before we go any further, let's look at some
terminology.
So antigens are large macro molecules usually a protein or a molecule with a protein component, like a
glycoprotein or a lipoprotein. An epitope is a small section of the antigen. Normally at eight to 15 amino
acid sequence that is recognised by the antibody. So, each antigen will have many potential epitope sites.
As I'm sure you know, different cell types express specific groups of proteins, as well as many proteins that
are common to all cells. Also, many proteins are compartmentalised within a cell. For example, they exist
within the nucleus, or the mitochondria, or the rough endoplasmic reticulum, or even in the membrane.
With immunodetection techniques, it is possible to not only distinguish between cell types.
For instance, neurons versus oligodendrocytes, because a specific cell type will express a specific protein or
groups of proteins. But also to visualise a location of proteins within the cell, for example, in the cell
cytoplasm or on the cell surface. This is important because a normal location of a protein maybe altered in
disease. There are two main types of antibodies made for research: monoclonal antibodies and polyclonal
antibodies. I will first introduce monoclonal antibodies.
A monoclonal antibody preparation contains only a single antibody type that's specifically recognises a
single epitope on the antigen molecule. Monoclonal antibodies are produced by immunising or injecting an
animal, for example, a mouse, with an antigen. In this case, the protein that we wish to generate an
antibody against. This can be a human protein. B cells are then extracted from the spleen of that animal.
These B cells are then fused with myeloma cells.
These are immortal cells and so can be grown indefinitely in a laboratory. By fusing the B cells with the
myeloma cells, we generate a cell called a hybridoma cell and this cell can also be grown indefinitely in a
laboratory and can secrete antibodies. So now, we have a population of hybridoma cells that can secrete
antibodies. However, not all hybridoma cells in this mixture will generate the antibodies that we want, but
some will. And each of these will generate an antibody that interacts with the specific epitope on our
protein of interest. We are able to isolate individual hybridoma cells from this mixture and then expand
them to generate a population of hybridoma cells. All of which generate a specific antibody that recognises
a specific epitope on the antigen used for immunisation.
Polyclonal antibodies are also produced by injecting an antigen of interest into an animal, usually a rabbit.
The animal's B cells will produce antibodies against the antigen, and these antibodies can be found in the
serum of the animal.
As mentioned earlier, a protein has many epitopes and different B cells will make antibodies against a
different epitope of the protein. So the serum will contain a collection of antibodies that recognise
different epitopes of the same protein. Hence the term, polyclonal.
Polyclonal antibodies may vary between production batches. And they also cross react with other proteins.
This is because the more epitopes that are recognised by an antibody mixture, the more opportunity there
is that any one particular epitope, that is amino acid sequence, may be found on an entirely different
protein.
Here is a very brief introduction to antibody structure. Antibodies belong to the immunoglobulin protein
family. They are Y-shaped as you can see in the cartoon and are composed of four polypeptide chains. Two
identical copies of a heavy chain and two identical copies of a light chain, so called because of their relative
molecular weights.
The arms of the Y contain the antigen binding site. Antibodies can be divided to five different types: IgG,
IgM, IgA, IgD and IgE. However, most antibody reagents are IgG, or occasionally, IgM.
Having briefly discussed antibodies, let us now look at how we can actually use them to detect proteins in
our sections. Let us think about how we can visualise a protein P, in the membrane of cell within the
section. We can do this using a direct method or an indirect method. In both cases, antibodies are applied
to the tissue section. In a direct method, the primary antibody, seen in green, so called because it binds
directly to the antigen, is directly linked to a report molecule shown in red.
These report molecules, and we will talk about these on the next slide, enable us to visualise the bound
antibody. This direct technique is suitable only for highly expressed proteins. Because if there are very few
epitopes available, the reporter signal may be too weak for us to see. The indirect method is a method that
allows us to significantly amplify the reporter signal strength. See the righthand side image.
Again, the primary antibody binds to the antigen. But in this case, this antibody has no reporter linked to it.
The section is then incubated with the second antibody, termed a secondary antibody that binds
specifically to the primary antibody. It is a second antibody that has a reporter linked to it. The secondary
antibodies are polyclonal antibodies, so will react with epitopes all over the primary antibody. The figure
on the right illustrates this, but only shows two of the bound secondary antibodies. Thus, multiple
secondary antibodies will bind to the primary antibody. and as all of the secondary antibodies carry a
reporter, the signal is amplified. The reporter molecules may be an enzyme or a fluorochrome. We will look
at these different types of reporters next.
In the fluorescence method, the antibody, primary or secondary, is linked to a fluorochrome. The
fluorochrome is a reporter molecule that is not itself coloured but will emit coloured light at a specific
wavelength in the visible spectrum when illuminated by UV light.
We have specialised microscopes, called fluorescence microscopes, that enable us to view such
fluorochromes. This method can be used to detect the presence of more than one protein at the same
time by using two or more different antibodies, each linked to different fluorochromes that emit light at
different wavelengths, as shown in the cartoon on the left-hand side. In the case of the enzymatic
detection method, the antibody, again, primary or secondary, is linked to an enzyme, most often an
enzyme called horseradish peroxidase, or HRP for short.
This is found in the root of horseradish. A substrate is then added to the tissue. And the enzyme and
substrate interact to generate an insoluble coloured product at the site of the antigen-antibody complex
that can be visualised under a light microscope. Such a substrate is called a chromogen.
The direct and indirect methods of automatic detection are shown on the right-hand side of this slide
where the yellow depicts a coloured precipitate being formed. However, there are some other factors that
we need to consider when carrying out immunohistochemical experiments. These are: the incorporation of
positive and negative controls, antigen retrieval, and blocking of non-specific binding. I will talk about each
of these next.
It is very important to include positive controls to assess fidelity of the technique and specificity of primary
antibody. I will explain with an example. BrdU, or bromodeoxyuridine, is a synthetic analogue of the
nucleoside thymidine, one of the four bases in our DNA, and is used to identify dividing cells. When
injected into the brain of a mouse, for example, BrdU can be taken up by cells and incorporated into their
DNA in place of thymidine when they divide, as shown in the cartoon. The presence of BrdU in a cell can
then be detected using an anti-BrdU antibody. Such a method is often used to assess changes in adult
neural stem cell division in the hippocampus. Let me explain. In the adult brain, the dentate gyrus, a region
within the hippocampus, is one site where new neurons are generated throughout our lives.
This process of generating new neurons is called neurogenesis. Neurogenesis within the hippocampus is
important for memory and also hippocampal neurogenesis has been associated with changes to our mood.
For example, in animal models of depression, antipsychotics have been shown to increase adult
hippocampal neurogenesis.
An example of BrdU staining in the hippocampus can be seen on the slide. So, in our hypothetical
experiment, let's assume that mice have been injected with BrdU to assess changes in stem cell
proliferation in the hippocampus after drug treatment. So we are looking for BrdU staining in the
hippocampus as a means to assess neural stem cell proliferation. What if we see no antibody staining
within the hippocampus? Does this mean that there was no BrdU incorporation and hence no division? Or
did the antiBrdU antibody not work? We can confirm that our anti-BrdU antibody and staining protocol
worked by including a positive control.
For example, we could use a piece of small intestine isolated from the same animal, and look for BrdU
incorporation in its tissue. This is because we know we can always find dividing cells here in the crypts of
Lieberkühn at the base of the villi. If we see staining here, this would confirm that BrdU was incorporated
and the staining procedure worked, even if was not seen in a treated animal hippocampus.
So this is example of the benefits, the necessity, of using a positive control. For a negative control, it is
mostly sufficient to make the primary antibody. Any positivity seen is then assumed to be caused by nonspecific binding of the visualisation reagents. One then compares results of the negative control and the
positive control against the test results before drawing a conclusion.
Let's now think about antigen unmasking, or antigen retrieval. We heard in the first part of this lecture how
fixation is important for preserving tissue morphology. But fixation techniques can also affect your ability
to detect antigens using immunohistochemistry. In essence, fixation procedures can mask or alter
epitopes, so that they can no longer bind to the primary antibody. Antigen unmasking or retrieval refers to
any technique where the masking of an epitope is reversed so that the antibody can again bind to it. In this
slide, two such techniques are highlighted, heat-induced epitope retrieval, or HIER, and protease-induced
epitope retrieval.
The mechanism by which HIER is achieved is not really known. But HIER can be performed using microwave
ovens, pressure cookers, steamers, or water baths. However, it is recommended to use commercial
systems specifically designed for HIER. HIER is normally carried out with sections immersed in a buffer
solution. Such a solution resists change to pH. That is, the acidity or alkalinity of a solution. A list of buffers
can be seen on the slide. Each will be tested to see which works best with the antibody being used. In
protease-induced epitope retrieval, sections are preincubated in enzymes. And Proteinase K, trypsin, and
pepsin have all been shown to have some effect.
Which enzyme to use and for how long is decided by trial and error. An over digestion will destroy both the
antigen and the tissue section so this method has inherent difficulties. On top of that it has been shown to
only work for a small proportion of antigens. Unfortunately, the antibodies we use can sometimes bind to
nonspecific components in cells and tissues with low affinity, by that I mean low strength. And can result in
a false positive signal. This is more often true for polyclonal antibodies.
To prevent this happening excess protein is added that will compete for and block binding to these
components, as shown in the cartoon. Serum contains a range of proteins that can compete out the
antibody from binding to sites other than its target epitope. Normally researches block with the serum of
the animal species that was used to raise a secondary antibody. For example, if a secondary antibody is
obtained by inoculating a goat with a mouse IgG antibody the serum used to block nonspecific binding
would be goat serum. Alternatively, we can use an excess of proteins like bovine serum albumin to block
nonspecific binding and this is very effective.
So when we carry out an immunostaining experiment we always need to consider the following: • what
positive and negative controls to include • do we need to use antigen retrieval • and what should we use
to block non-specific binding. Let me finish by showing you a couple of examples. On the slide is an image
of a paraffin wax section from a mouse model of Alzheimer's disease showing astrocytes surrounding an
amyloid plaque. These are protein aggregates that build up in the Alzheimer's diseased brain.
The method used to stain this section was a double indirect immunofluorescence staining. To do this the
section was incubated simultaneously with two primary antibodies that each recognise a different protein:
glial fibrillary acidic protein, or GFAP, and beta-amyloid. GFAP is a protein expressed by astrocytes in the
brain and beta-amyloid is a protein fragment that makes up the amyloid plaques. The anti-GFAP primary
antibody was then visualised with a secondary antibody linked to a green fluorochrome. And the anti-betaamyloid primary antibody was visualised with a secondary antibody linked to a red fluorochrome. The blue
that you can see in the image are due to the nuclei of cells being stained with a fluorescent blue stain
called DAPI. For my final example I'm showing you a paraffin wax section from a normal mouse brain
showing an area of the hippocampus. This section, again, stained with an anti-GFAP antibody to visualise
astrocytes in the section. But this time the binding of this antibody is detected by indirect
immunoperoxidase staining using DAB. The section is counter stained with hematoxylin and a blue dye to
show all nuclei. To summarise, from this presentation I hope you can see that histotechniques are
invaluable tools for neuroscience and have contributed significantly in the elucidation of many normal and
disease processes. Today we have talked about the importance of tissue preservation by fixation or
freezing and about the processing and embedding techniques used to support the tissue for sectioning. We
have discussed some of the staining techniques that can be used to identify the cell types and proteins
within a section and provided some examples about such techniques can help us identify changes that can
occur in the diseased or damaged tissue. In topic two of this week Carl Hobbs will demonstrate how we
actually carry out these techniques in the laboratory.