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  University of Idaho
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  Psychology Dept.
  University of Idaho
  Design - P&D  CTI

If we look in slide three, we also see there are a wide variety of non-invasive techniques.  Non-invasive techniques are primarily what we use in humans today, although we do use some others as well but for the most part, these are the things that we’re using especially in the fields of neuroscience, cognitive psychology and physiological psychology on a more global, human aspect. 

Ok, so now that we have a little bit of an overview of what the two different groups are, let’s talk about invasive techniques first.  Then in the next section we will talk about non-invasive techniques. 

The first thing to note is (as we see in slide four) that we have to need some kind of device to help us when using invasive techniques.  That device is called the stereotaxic instrument.  A picture of the stereotaxic instrument is shown in slide five.  And as we see here, there is a holder that allows you to swing the system and the instrument in all directions.  We have vertical knobs that allow you to vary the depth of the probe, we have an anterior /posterior knob which allows you to go front and back, and of course a lateral knob that allows you to go side to side.

So, once we have an animal that’s anesthetized and lying in this instrument, and we have a good idea of what we want to do, then we can use this instrument to line up the animal before we insert our piece of equipment.  Humans have very similar types of techniques and instruments as well; they’re just a lot larger.  Again, what we do is we anesthetize the animal, open the skull, and then drop in a microelectrode, a micropipette, or a micro-canula into the brain.  Then we can do some procedure.  We’ll talk about these techniques a little bit more in detail later.

For animals, especially rats, the brain is very well defined and has been examined extensively.  So we know that if we drop a micro-electrode down a millimeter and a half and we’re laterally one millimeter from midline, and we are five millimeters from some other major structure, we know that we are in x piece of tissue.  We can be highly confident of that based on the different brain atlases that we have of rats, mice, dagoos, and a variety of other different organisms.

Humans techniques, although are fairly good, are not as detailed as animal techniques. The fact is that we cannot do experimental microscopic surgery on live humans.  Ethically there’s a problem with that.  So what we have to do is use other things to help us along.  So, let’s walk through the general procedure again.  We show this in slide six.  The first thing that we do is identify the dependent measure that we’re going to measure in the organism.  Let’s say that it is freezing behavior, or moving a particular paw or whatever.  We then measure how often that measure occurs under some particular situation.

Once we’ve done that and we have a solid baseline, we anesthetize the animal. There’s a wide variety of different anesthesia types of materials we could use but a common technique and a common anesthetic is ether.  Anyway, we put the animal in a little bell shaped jar, we put some ether in and the animal goes to sleep.  Once the animal’s asleep, we open the skull with a scalpel and drill a small hole (usually with a small drill).  Of course you don’t go too deep.  So we put a hole in the skull and open the skull (usually with a little type of wire saw), and we now have a hole in the skull.  We then place the animal into the stereotaxic instrument and use the brain atlas to identify how far over, how far down, etc., where we want to go with our micro-electrode.  Then, (as we see on slide seven) we use the stereotaxic instrument to drop in a device.  Once we then have performed the technique that we want to do (e.g., destroy a piece of brain tissue or put in a canula to inject different hormones later, or whatever it is), we can then seal off the skull with a variety of different substances.  We allow the animal to recover (which usually takes several days because things have to heal), and once the animal seems to have recovered measure the dependent variable that we monitored earlier and make comparisons between the two. 

So, what are some techniques we can use?  Well as we can see in slide eight, there’s a wide variety of techniques.  There are lesions or what we call ablations where we basically destroy brain tissue.  There are cannulations, there are push/pull techniques, and electrical recordings.  We will talk about each of these techniques in detail shortly.

So let’s talk about electrical lesioning techniques first.  Lesioning in general destroys particular sets of brain tissue.  So, what you’re doing is observe the animal beforehand, you go in and destroy a piece of brain tissue, and then you see what happens to the animal after the lesion has occurred.  There is also a wide variety of different ways that you can lesion tissue. 

The first way is to use electrolytic current.  Basically what we’re using here is D.C. current.  So, (as we see here in slide 10) we observe the animal, we put in an insulated needle except at the point, then we apply the current.  In essence we burn the brain tissue that we’re trying to destroy.  The tissue ultimately dies, and after the tissue is dead we observe the animal for changes in behavior. 

The major advantage of electrolytic lesioning techniques is that it is cheap and it’s relatively simple to do.  There are some problems with the technique as well and shown in slide 11.  The first disadvantage and problem is that when you stick a piece of metal into the brain, it leaves a track.  Not only are you damaging the tissue at the end of the tip, you’re damaging tissue on the way down to that piece of tissue.  So, you’re damaging other brain tissue.  So what’s the solution to that?  Well, what we do is go in at different angle.  We use different sets of animals and different angles to destroy the particular brain tissue.  If the results are the same, we can be pretty sure that the brain tissue is involved and it’s not the track plus the brain tissue.

A second major disadvantage relates to the amount of current.  If you apply too much current, you basically deposit metal particles from the electrode.  These can irritate tissue and cause seizures which mess up your results.  So that’s the first technique, electrolytic techniques.

The second technique is what we call radio frequency techniques.  Basically radio frequency techniques coagulate tissue.  It resembles very similar stuff to a microwave oven.  So, again, what we’re going to do is anesthetize the animal, put it into the stereotaxic instrument, and to again insert our electrode.  Again it’s insulated except at the tip.  But when you apply energy in this case, it causes the water molecules inside the neurons to oscillate.  That is, they start to vibrate.  When the vibration begins to build up, just like a microwave oven, it basically heats up, and ultimately kills the cells within the particular area.

The advantage of this technique is that it avoids the metal particles, but the disadvantage is you still have the electrode track.  So, again, you have to come at different angles to try to avoid that result.

So those are two different types of electrical techniques.  Well there are also other techniques that we can use.  The third technique that I want to talk about is called chemical techniques.  Chemical techniques are the most commonly used today.  Instead of an electrode, what we do is use some kind of canula or tube.  Basically it allows you to put different types of chemicals in places where they can kill neurons or just influence neurons.  What I’ve done, (as you see in slide 15) is give you a couple of different chemicals that we can use within the brain.  For example, 6-hydroxy-dopamine kills all dopamine neurons but leaves other neurons alone.  Or Kanic acid basically kills the somas but leaves the axon tracks in place.  There are many, many other types.  In general, what we do is select the type of chemical compound that we want to use and use that within our system.

Now the next technique is a little bit opposite of the chemical neurotoxin techniques.  These are shown on slide 16.  Basically what these do are stimulate the neurons by putting in some kind of compound or neurotransmitter.  They can also be analogs or agonists so they behave like different types of neurotransmitters and particular drugs.  We can also put in some kind of compound that we have no idea what will happen.  Let’s say we want to check out and see what Coca Cola does to the brain or Pepsi.  Well, we can put that in and see what goes on in there if we have some good reason to do that.

General cannulation procedures (as we see in slide 17) are very similar to the electrical techniques that we discussed earlier.  Again you make the trephine hole, drop in the canula, you cement the canula in place, you put the material in and you observe what happens to the animal.  You can completely kill the tissue, or you can just cement the tube or a microcannula in place.  You let the animal recover, and then you inject it with a compound and observe what happens.  Either way, whatever technique that you use, the result is the same.  You’re observing the animal when some particular chemical has been added to the brain.

What’s the disadvantage of chemical techniques?  Well the major disadvantage is that over time (as you put stuff in) the area fills up with the chemical.  Just like you stick a needle into your arm and inject some medicine.  As the levels builds up it causes pressure, and it forms a little puddle of fluid in your arm.  Well the same thing occurs in the brain and that’s a major problem.  So, we’ve developed what we call push/pull techniques.  These are shown on slide 19.  Basically the major thing about push/pull techniques is that number one, the canula is a little bit wider and it has an extra tube.  An example of a push/pull tube is shown on slide 20.  So, basically what you have is an outside tube and then you have an inside tube where the material is going to be entering.  What you do is insert the solution and withdraw the excess.  

So what are some advantages of push/pull?  Well, we see these in slide 21.  Number one, it localizes better.  You can also add dyes, you can change concentrations, compounds, and on and on.  Ultimately, this technique has become a very, very important tool that we have in examining brain tissue.  The key about this technique is that we’re using miniaturization and has been helping us do a lot of different things.

Ok, so now we’ve talked about a variety of different techniques.  Let’s talk about the next technique called canulations.  Generally canulations involve placing a catheter into the circulatory system of an organism.  However, it just doesn’t have to be a circulatory system; it can be in other areas as well (spinal cord, 4th ventricle of the brain, etc.).  We’ll talk about those later.  In general, what we do is put the catheter into the circulatory system.  And as you can see in slide 22, there’s a variety of different places where we can put that catheter.  Each one of those locations has some advantages and disadvantages.  For example, putting the cannula in the arm is very simple and is not too dangerous.  Whereas putting the cannula into the jugular vein of the neck, the femoral artery of the leg, or the aorta can be significantly more dangerous. 

There’s a lot of advantages (as we see in slide 23) about using catheter techniques.  Number one, you can put it in basically anywhere.  For example, you can deliver materials into the venous system.  Some of these materials can get past the blood brain barrier.  However, sometimes we want things to just get into the vascular system and not cross the blood brain barrier.  Thus, we add different substances to make the molecules much larger.  Finally you can do the procedure in alert animals, such as humans. 

What are some disadvantages?  Now, the key for the problems with catheter techniques is shown in slide 24.  When you put something into the circulatory system, over time clotting begins to occur.  As we see in a couple of areas (the femoral artery or descending aorta), the different time periods can be significant.  On the other hand, we can put canulas almost anywhere.  Some examples (as we see in slide 25) are the mouth, stomach, liver, spinal cord and on and on. 

So, what about our next technique.  These are called osmotic pumps.  Osmotic pumps are basically used to deliver compounds by osmosis.  We can use a single compound or multiple compounds over prolonged periods.  Furthermore, these devices can be used both within animals and in humans.  An example of an osmotic pump is shown in slide 27.  Basically we have an outside chamber with some kind of tube inside with the solution. The osmotic pressure’s going to come through holes within the outside chamber and depending upon how fast you want it to come in.  The pressure basically begins to squeeze the tube and it pushes the solution into some catheter and ultimately a needle.  There’s a variety of different types shown in slide 28.  There’s a single barrel technique where you only have one compound.  You also have multi-barrel delivery techniques which have multiple solutions.  So, you have all sorts of different stuff inside one particular pump.  In the old days you had multiple cords that you tied off around different tubes.  Today we use small microcomputers to regulate the substance entering the final catheter.

The procedure is pretty straightforward.  You put the needle where you want it (If you have to, you use the stereotaxic device).  You put the pump inside the body (usually in the fat tissue of the back, and you tie off the end of the tube with some silk until you’re ready to deliver it.  Let the animal recover, you cut the silk and then you observe what happens.

Advantages, you can deliver substances for long periods.  You can also deliver constant amounts of substances instead of having spikes of a particular drug.  It’s very easy and it’s relatively safe, depending upon what you’re doing. 

Ok, now we’ve talked about catheter techniques.  Let’s talk now about a different technique called autoradiography.  Autoradiography, as we see in slide 31, uses different types of radioisotopes which expose X-RAY film.  Basically radioisotopes throw off energy and you can then use them to identify receptor sites.  As we see in slide 32, there are all sorts of atoms that we use.  Now there are two different procedures that we have for autoradiography.  These are called invivo autoradiography and invitro autoradiography.  So, let’s talk about in vivo autoradiography first. 

The first thing you do is you take some kind of isotope and you attach it to some substance that you’re trying to evaluate (e.g., a hormone).  Attaching the isotope is called a tag and the procedure is called radioactive tagging.  Steps are pretty simple, as we see in slide 34, you radiolabel the substance, you inject it in the animal, you let it circulate in the blood and it goes to the different systems within the body.  After a period of time, you kill or sacrifice the animal, you then take out the organ that you’re interested in; it could be the brain, it could be the liver, it could be all of those things.  Then you microtone or slice the tissue into little pieces.  Then (as we see in slide 35) you place the pieces on a tray, cover them with a piece of x-ray film, put them in a darkroom somewhere,  and let them sit for days, weeks, or even months.  After a period of time, you take the tissue off the pieces of the xray film and you then go and look for dark spots or white spots, depending on what you’re using on a computer screen.  If you have any spots, that’s where the receptor sites are located.  So in general, it tells you where the test receptor sites are located within the system. 

What are the advantages?  Well the major advantages (as we see in slide 36) are that it’s the first good step to find receptor sites, and it’s a good procedure if you’re not sure where the particular receptor sites are located.  It could be just in the brain, or it could be all over in different parts of the body.  It’s a good technique for new substances where you’re not sure where they’re going, and it’s also faster than in vitro techniques which we’ll talk about shortly.

What’s some disadvantages.  Well as we see in slide 37, it’s more expensive than other techniques.  Sometimes you also don’t see anything.  So, what happens if you don’t see anything, did you just waste your time?  Well, we don’t know.  It could be a procedural error, somebody screwed up and turned on the light, the assay may have decayed, we have the wrong type of isotope, or there just may not be any receptors there. 

So that’s the first technique.  So, let’s go a little beyond that and look at the next technique called invitro autoradiography.  Invivo means in life, invitro means in test tube or what we call glass.  So, how do you remember the two?  The t in vitro means test tube.  In essence we use this technique when we have ideas of where the particular receptors are located and we want finer detail than the in vivo technique.  So what’s the procedure? 

Well, as we see in slide 39, first we kill or sacrifice the animal.  Then we take out the brain or other tissue that you’re interested in, slice the tissue, and then pour the radioactive tag, hormone, or substance over the tissue.  Then you allow it to sit for a while, could be a day, could be a week, or an hour.  You then put a buffer on and wash the solution off the tissue.  Then cover the tissue with the x-ray film and allow it to incubate.  If there’s some kind of radioactive labeled material there, its bound to the receptor and again puts dots on the film.  Of course we can put it on a computer machine to count the number of different receptors that are located at that site.  So again, what we can do is count the number of particular receptor sites.  However, it is within a specific piece of tissue rather than having a general idea or area of where it was before.

Next technique is a little bit finer technique and shown on slide 41.  These are called micropunch techniques.  In micropunch, we use a microtone to slice the tissues. Then we use specialized hypodermic needle to punch out a little piece of tissue.  Just like you punch out a particular type of cookie dough for Valentine’s or St. Patrick’s Day.  However, the size is usually relatively small (10 to 15 micrograms).  What we do is take a core sample and put it into a test tube.  From there we can do all sorts of different things (as we can see in slide 42).  We can break up the tissue, we can destroy the ligens but not the cell bodies, we can look at membranes, and on and on.  Ultimately, we are trying figure out how many receptor sites you have within one small piece of tissue.  So, again, it gives you a much finer level of analysis. 

So what we started with was a broad spectrum technique using autoradiography where we were just trying to figure out where receptors were.  Then we went to invitro autoradiography where we narrowed it down, then we used is a micropunch technique to get fine levels of analysis of different receptor sites within that area.  Finally, we can do even finer levels of analysis using radio amino assays (which I’m not going to talk about). Of course, there are other techniques as well.  

So let’s kind of summarize all of this.  We see in slide 43 there are a wide variety of invasive techniques that we can use.  They’re used for lots of different reasons and we can, and it allows us to have a very fine level of analysis for what we’re trying to do. 

In general, these techniques are used very, very often within Physiological Psychology and all of neuroscience.  For that reason, they are very, very important to understand and understand well.  The only difference between using it in animals and using it in humans is the level of analysis.  Usually when you’re doing it in humans, the human has died rather than is still alive.  In the next section, we’ll talk about some non-invasive techniques.