#07 Biochemistry Protein Characterization Lecture for Kevin Ahern’s BB 450/550


Another Monday beckons,
another week beckons. One day closer to an exam. Student: Whoo! Kevin Ahern: Yay, huh? One day closer to
your opportunity to show me how much you know. That’s good. I hope you had a good weekend. Student: Fantastic. Kevin Ahern: Fantastic? Student: We won. Kevin Ahern: Are we
talking about football here? Student: Yeah. Kevin Ahern: Okay. So the football team won. So last time,
I threw out the topic to you of the 2D
gel electrophoresis, and I think that’s a really
phenomenal technology. I think it allows not “I think”, I know it allows us to do amazingly
complex analyses of cells. And if we have cells that
have different experiences one being a tumor cell,
one not being a tumor cell, one being treated with a drug, one not being treated with
a drug, one being starved, the other not being
starved, et cetera, et cetera we we can use this
technology to see very clearly at the protein level how these changes occur
inside of the cells. Several students after the
class asked me if there were libraries of gels that were
out there that are cells of known treatments. The answer is, there are. But many laboratories
will actually do their own side-by-side comparison
because one of the things that you see is the reproducibility
is not 100% the same, so if you’ve done
both of them in your laboratory at the same time, you’re a little bit more
able to compare them. So that’s something that happens. But, yes, there are libraries
of such things out there. And I just realized,
I haven’t checked the camera to make sure it’s
properly on the screen. So give me just a
second to check that. Doo-do-doo-doo. And the answer
is, it was perfect. Alright. There’s nothing worse than
looking at your video afterwards and you see you had it
about halfway on screen and about halfway off the screen. And you guys like that
about as much as I do, so, yeah, maybe less than I do. One of the things I skipped
over in getting to tell you about 2D gel electrophoresis
was to tell you about gel electrophoresis itself. So that’s how I’m going
to start the lecture today, telling you how gel
electrophoresis works and I’m going to talk about two different types
of gel electrophoresis. The first type I will talk about is actually the
simpler of the two, and it is what we
refer to as DNA, separating DNA by agarose
gel electrophoresis. Agarose is, and there’s
the word right there agarose gel electrophoresis, I keep popping out here agarose gel electrophoresis
is a technique. I don’t have
a figure for it anymore. Your book used to have
a figure and then they took that away from me, so I don’t
have the figure out for it. But I can tell you
it’s, in principle, very much the same as
polyacrylamide gel electrophoresis. So let me just show you
what that looks like. Agarose gel electrophoresis
is what we use to separate fragments of DNA. We can also separate
fragments of RNA with it. We do not use agarose gel
electrophoresis to separate proteins, and you’ll see why that’s
the case in just a little bit. The first reason, though, that
we don’t use it to separate proteins is that nucleic acids
are way bigger than proteins. The biggest molecules in the
cell are DNA molecules, by far. Proteins don’t even come
close in terms of size. What the agarose provides, in
the case of DNA separations, or what the
polyacrylamide provides, in the case of protein
separations, are a matrix. And we can think of this matrix sort of like it’s
schematically shown here. The matrix is a series
of strands or connected things that provide a support. The support is to support
the liquid of the buffer. So just like we could
take a mix of Jello and put it into
water and boil it, when it cools down, it
forms a solid support based on what was in there, so,
too, can we do with materials for the gel, the difference
being, in the case of a gel, that these strands that
provide the support will provide little channels or
little holes through which the
macromolecules can elute. And I’ll show you how
that happens, okay? Agarose has bigger holes
than polyacrylamide does. So we need those bigger holes
to separate DNA molecules. So how do I separate using
gel electrophoresis for DNA? Well, first of all,
I take my DNA molecules that would be a mixture
of different sizes. And I would apply them
to the top of my gel, as you can see here. So I make these
little indentations that are what are called “wells.” And into these wells, we pour
our mixture of DNA fragments. DNA fragments are
negatively charged. They’re polyanionic, meaning that they have
many, many negative charges. For every base that we add,
we get another negative charge. So the charge is
proportional to the length, and the length is
proportional to the length. Now, you’ll see why that sort
of makes sense, in a second. The charge is
proportional to the length, and the length is
proportional to the length. And what we do in
separating these guys is we use an electric field. The electric field we use places
a negative charge at the top. You can see that little
negative ion right there. And it places a positive
charge at the bottom. The DNA molecules,
being negatively charged, are repelled by the
negative at the top and attracted toward the
positive at the bottom. Well since the ratio of the
charge to size is constant, that is the longer
molecules have more charge, but they also have more size the separation that happens
between these molecules is solely on the
basis of their size… solely on the
basis of their size. The smallest guys can
move the fastest through these channels and they
go racing through the gel. The largest molecules don’t
have that same mobility and it takes them longer
to get through the gel. So at the end of a stint
of gel electrophoresis, what we see is the gel products. So this is a protein gel, but a DNA gel would
look very much like this, where we have fragments that
have been separated by size. So this would be the
largest molecules up here. These would be the smallest
molecules down here. And these are specific
fragments, in this case, that have been
purified of a protein that have a given
size that’s there. So, in principle, DNA
electrophoresis and protein electrophoresis are the
same after we have to do some manipulations to proteins
to make that happen, and I’ll show you
how that occurs. So DNA electrophoresis
makes sense? Yes, sir? Student: So if the
charge on the bottom isn’t great enough that it’s,
it’s not just going to tear through the gel? Kevin Ahern: So his question
is the charge on the bottom great enough that it’s just
going to not tear through the gel? In fact the molecules will,
if you leave it long enough, go all the way through the gel. Yes, they will. So they will go
all the way through, this is cutting out. They will go all the
way through the gel. So there’s several
variables that we have. We don’t need to
consider them really here, but I will tell you we can
change the percentage of agarose, which will actually change
the size of those holes that the DNA molecules
are passing through. So we can optimize
that for different things that we’re
trying to separate. And I’m getting some noise. Maybe that took care of it. So that’s DNA electrophoresis. It’s pretty straightforward. With protein electrophoresis, we’ve got a different
consideration. And the reason we’ve got
a different consideration is, first of all, proteins are globs. And second of all, proteins don’t have a
uniform mass-to-charge ratio. Some proteins are going
to be positively charged. Some are going to be
negatively charged. Some are going to be neutral. And that charge is really unrelated
to the size of the protein. So if we try to separate
proteins without some other things to give an artificial
size-to-charge ratio that’s constant, then we’re going to have trouble. Because if I take my
mixture of proteins and I’ve got some
positive ones on top and some negative ones in there, the positive ones aren’t
even going to enter the gel. They’re not even going to go in. Boy, this is really
misbehaving today. Alright. So I have to do something,
then, to make the, I have to do something
to make the proteins have a reasonably constant charge,
or size-to-charge ratio. So the trick that’s
used is a very clever one and it works very, very well. It may seem a little odd, at
first, but it’s actually a very, very good way to give
proteins an artificial size-to-charge ratio
that’s constant. What we do is take
the mixture of proteins that we want to separate, and we add excess
detergent, called SDS. That stands for “sodium
dodecyl sulfate.” So it’s a long carbon
chain molecule that has at one end a sulfate. Now, that sulfate is
negatively charged. When these proteins
encounter the SDS, if you recall when I talked about what detergents
can do to protein, what did I say would happen? They denature, they unfold. So this protein that
starts out as a glob, first of all, elongates
out into a nice long chain. So, visually, we could imagine
this guy is going to look something like a straight
DNA molecule, not as big, but a straight
DNA molecule. The second thing
that happens is these sodium dodecyl sulfates
completely envelope the chain. Alright? They just completely go all
the way around the thing, making like a Twinkie
or something, okay? A Twinkie’s got the little chewy center, right? The chewy center
being the protein, and it’s got this coat of
stuff all the way around it. Well, that coat, of course, is proportional to the length
of the polypeptide chain. Longer polypeptide chains
will have more of those sodium dodecyl sulfates
than smaller ones will. So the size-to-charge ratio
is relatively constant. It’s not absolutely constant, but it’s relatively constant. And, in fact, for most purposes, it’s constant enough
that we can get very, very good separations
based on size. So once we’ve done that, we take our mixture of proteins, that are now all
coated with this SDS, and we separate them
on a polyacrylamide gel. And as I said earlier, the only difference
between agarose and polyacrylamide
is that polyacrylamide simply makes smaller pores, smaller holes, for those
proteins to go through. We apply an electrical current, just as we did before, negative at the top, positive at the bottom, and we separate solely
on the basis of size, how fast they can move
through that chamber. Now, and so when we do that, we actually end up getting a gel. This actually is a protein gel. We can see these are marker
proteins that have different sizes, the largest ones being up here, the smallest ones
being down here. And if we know the sizes of
these known proteins over here, we can actually determine
the size of an unknown protein by seeing where
does it line up with. Is it 50,000 in molecular weight? My protein must be about
50,000 in molecular weight. So this technique has an acronym. First of all, polyacrylamide gel electrophoresis
has the acronym PAGE, P-A-G-E. When I use SDS, which I almost always
do with proteins, we call it SDS-PAGE. SDS-PAGE. So SDS-PAGE allows me
to separate proteins on the basis of their size, very much like
I separate DNA molecules on the basis of their size. So when I’m doing that 2D
gel electrophoresis that I talked about on Friday,
that second dimension, or the first dimension, we had isoelectric focusing, and I said we cut it open, and we laid it on
top of this gel? Well, this gel is
a polyacrylamide gel. We have to make sure
we get some SDS in there so the proteins all
make the Twinkie shape, right? And then we run
them through in that second dimension as SDS-PAGE. So the first dimension of
a 2D gel is isoelectric focusing. The second dimension
is SDS-PAGE. And thanks to that, we can actually separate
these molecules and determine, literally, the amount and
presence or absence of virtually every protein that’s
made in a given cell. Student: So that SDS part, you just call that
isoelectric focusing? Kevin Ahern: What’s that? Student: Is that SDS part, you just call that
isoelectric focusing? Kevin Ahern: No. Isoelectric focusing is
a different technique. That’s the one where we
used the charge to separate the molecules in the tube? Student: For DNA. Kevin Ahern: No. That’s for protein. That’s what I talked
about last time. Isoelectric focusing, I put the stuff in the tube, and I had the things that had
minus 50 all the over to plus 50? Right? So that first dimension, I separate on the basis of
what was essentially the pI. Okay? And the second dimension
I separate on the basis of size. That second dimension
is known as SDS-PAGE. Yes, sir? Student: So the SDS coat
doesn’t affect so much with the isoelectric focusing? Kevin Ahern: Okay. That’s a good question, a common question. So people will frequently say, “Does it screw up the
isoelectric focusing?” Well, no, because they’ve
already been separated by the isoelectric focusing. And so we’re just covering
what’s already been separated on the basis of pI with SDS. It doesn’t screw
anything up, at all. If it did, we would
have a problem. Student: Okay. So they’re not actually
run at the same time? Kevin Ahern: They’re
not run at the same time, because we have to separate
on the basis of pI first. That would be
a good exam question if we tried to put the SDS in
with the isoelectric focusing, what would happen is, everything
would be negatively charged. It would all go to one end. Good question. Yes, Shannon? Student: So how is it physically
transferred to the PAGE? Kevin Ahern: It’s just
laid on top of the gel. It’s just, so instead of
having individual wells, I would just have a long thing. I’d lay my little
tiny tube up there. And I would just lay it on there and run electrical
current through it. Yeah. The people who run
2D gels, it’s an art. Believe me, it’s an art. So getting that
little tube on there and not breaking it and
fracturing it and everything, so that it lays evenly
across the gel surface, is very important, and
there really is an art to it. Yes, sir? Student: I’ve seen agarose gels
for DNA that are relatively small. Is there a standardized
size for these? Or is there a variation in size? Kevin Ahern: His question is, do we have variations
in polyacrylamide that we use for protein gels, because we do see variations
that we use for agarose. And the answer is, yes, we do. So we can run polyacrylamide
gels varying the percentage of polyacrylamide that’s there
and also varying the number of links between the
individual strands. And that is based on the
chemistry that’s involved in making a polyacrylamide gel. And all we’re doing,
in either of those, is really determining
the size of those holes that the proteins
are moving through. So we can adjust that. If we have a bunch
of small proteins, we would run a different
kind of a percentage of a gel than we would if we
had very large proteins. Student: Do they
vary the actual size of the plate itself, though? Kevin Ahern: Do they vary the
actual size of the plate itself? Yeah, you can do that. If you’re looking at something
that is quick and dirty, you can run a very
tiny little gel. If you want to go and do
2D gel electrophoresis, you would typically
run a fairly large one because you want to get as much separation as possible for those. So, yeah, they do
vary the size, as well. Okay. So, good questions. That was pretty much
what I wanted to say. I didn’t show this last time, but that was from
a 2D gel electrophoresis, the difference between a normal, proteins from
a normal human colon cell and those from a
colorectal tumor. And you could find many
differences between these, and those are, as
you might imagine, of a considerable
amount of interest. Well, one of the
things that we do, we’re doing all
these techniques for, is so that we can
purify it, that is, so that we can make plenty of a pure amount of
compound of proteins. And, as I mentioned earlier, one of the things we see
in the purification process is that that purification, we never really know what’s
going to work before we do it. So we have to really be
very careful to monitor during the purification
process where is my protein. Is it in the pellet? Is it in the liquid? Is it in the first part that
comes off the anion exchange or the last part that comes
off of the anion exchange? Because the worst thing
that you can do is assume it’s in one fraction and
throw that fraction away. And I can tell you
hundreds of stories of people who’ve
done exactly that. So people get very careful. If you’ve been working on this, this is your PhD
project, or whatever, you’re going to be
checking every component of that purification
process to make sure that you’re not throwing the
baby out with the bath water, as it were. Yes, sir? Student: So I assume some
of your stories are like, okay, the protein
should be here… Kevin Ahern: Should
be here, yeah. Student:…but it’s
actually in this part. Kevin Ahern: Right. Right. So that can be a real
problem if that protein is life and death for you, in terms of
a thesis or something. So what you see
here on the screen is a sort of a depiction
of protein separation of a purification process. Here is the unpurified. Here was the stuff after
we did one fractionation. I haven’t talked about
salt fractionation, but one way of separating things. Another separation. Another separation. At each step, it’s
getting purer and purer, until we finally see at the end, hopefully, something that
is almost absolutely pure for us to work with. Well, there are some
considerations for that, relevant to the numbers, and I need to sort of
step you through this and tell you
a little bit about this. So one of the things we want to
know in doing a purification is, first of all, where
is our protein? But, second, how efficiently
am I purifying this protein? Because I’m going to be
publishing this result, and I want to report to others, “Hey, this method really is good. “It works very well,” et cetera, so that others
will have an idea about how much material they’re
going to get out of it. And so what you see on the screen is a table following the
purification of a protein. And, yes, I think that you
should be able to do calculations like I’m going to describe
to you in a second here. So what we see are the several
steps in this purification that you saw in the last figure. You had a homogenization
just to bust open the cells. You fractionated it. You did ion exchange
chromatography. You did gel filtration. And finally, you did
affinity chromatography. And so what this table
is showing you is, really, how much of the
protein that you’re getting, apart from everything
else in the process. So, in this case, we
started with a protein and we had 15,000
milligrams of protein. And it’s very easy to determine the protein concentration
of a sample. I bust open a bunch of cells. Maybe I drew up a liter of cells. I bust open the cells, and I get 15,000 milligrams, which is quite a bit of protein, out of my cells, that’s here. I haven’t done any purification. All I’ve done is just
bust open the cells. So I need to know how much
material I have to start with. Well, that’s a
good starting point, but of more importance to me is, well, how much of my
protein is in there? And I don’t know
in terms of weight but I can measure the
activity of the protein. Let’s imagine my protein converts
one molecule into another. I can take that crude mix and
take a very tiny aliquot of it and treat it with this compound
that it normally acts on. And I can ask the question, well, how many molecules
of this compound that it works on got converted? So I have a definition there
of what’s called a “unit.” Let’s say a unit might be
a conversion of one nanomole of this molecule
into something else. So I would say, well, I
measure how many units I have in my total mix that’s there. In this case, I had 150,000
units of my desired protein. The specific activity
of that is the total number of units that I have, divided by the total amount
of protein that I had. So specific activity would
give me units per milligram. This would be 150,000
divided by 15,000, or 10. The yield is 100%. And the yield is 100% because
this is my starting material. I haven’t done anything to it. I haven’t lost anything. I haven’t gained anything. And my purification
level is 1 because, again, I haven’t purified, I haven’t done anything with it. This is just where I started at. After the salt fractionation, I go back and I look and say, “Whoa! I lost a lot of protein here.” I’ve only got 4,600
milligrams of protein, but謡hoa!悠 still have
most of my protein activity. That’s really good. I’ve retained a good
deal of my original. I only lost 12,000 units, but I got rid of about 2/3
or 3/4 of the total protein. So this tells me that I have purified my protein
to some extent, because now my specific
activity has changed from 10 units per milligram
to 30 units per milligram, meaning I got rid of
a bunch of junk I didn’t want. I threw out a little bit of
the baby with the bath water, maybe the leg or
something like that. [scattered laughter] But… that’s bad. [scattered laughter] Bad professor, okay? But I have 30
units per milligram. My yield is the number
of units that I have now, compared to how many
units I started with. So this, divided by
this, times 100 gives me, I’ve got 92% yield. That’s pretty darn good. Ninety-two percent of
my protein is there, and three-quarters of the
junk I don’t want is gone. The purification level is 3
because the specific activity improved by a factor of 3, 30 divided by 10 gave me 3. Well, I can continue
this process, and you can go
through the numbers and see each of these
as you go through. And the bottom line
will be that as I get further and further
to the bottom, I keep losing more
and more of my protein. There’s no method that’s
going to be absolute in terms of keeping
all my protein. But what we see is that the
specific activity goes up enormously, which means that this stuff right
here is really relatively full of my protein and there’s very
little other stuff that’s there. So this has a total
activity, 52,500 units, and it only has 1.75
milligrams of protein. That’s 30,000
units per milligram. I’ve got a yield of 35%, meaning I did throw away
a lot of the protein. But, by golly, I purified that
sucker by a factor of 3,000. Alright? So I got rid of an awful lot of
junk in the methods that I used to purify my protein. You might say, “Well,
how do you know when you get it absolutely pure?” And the answer is,
you never really know. But you can analyze it
on a gel and you can see, are there a bunch of
other bands on this gel? Or is it just my protein
that I’m seeing on that gel? And that can be a
really useful thing. If we look at the gel
that I just showed you, Oh, wrong one, okay, we can see this
has basically gone, they don’t really show other
bands here unfortunately, but you might imagine
that if you had a protein that wasn’t very pure you would
see some other molecular sizes that would be in
this particular lane. And you would see those
disappear the further I get along with my purification. Questions on what
I’ve just told you? Yes, sir? Student: So the purification
level was the current specific activity level divided
by the previous? Kevin Ahern: The purification
level is the current specific activity level
divided by the starting level. So the starting had 10, and I’m looking at how much
I’ve purified it compared to what it was I started with. So in this case, I had 10
units per milligram to start. Down here, I had 30,000
units per milligram. So my purification level is
30,000 divided by 10, or 3,000. Make sense? Yes? Student: So the gel
filtration is the SDS-PAGE? Kevin Ahern: No. Gel filtration is the method
I talked about the other day where you separate
on the basis of size. Student: I thought
SDS-PAGE separated by size. Kevin Ahern: It does. But there are other things
that separate by size. So gel electro-, that’s
where you had the beads with the little holes in them? Yeah. That’s gel filtration. Yes, sir? Student: If you
continued purifying this, would you start seeing
diminishing returns? Kevin Ahern: If you
kept purifying this, would you see
diminishing returns? In fact, at every
step of purification you will always see
diminishing returns, yes. Yeah. Alright. So that’s purification. What I want to do is
spend a little bit of time talking about
characterization of proteins, and we’re going to talk about
some techniques of spectroscopy and also some just simple
tools to work with proteins. I’m going to skip over
amino acid analysis and I’m not going to hold
you responsible for it. There are, suffice it to
say that there are a variety of chemical and chromatographic
tools for analyzing the amino acids in proteins, but the reality is that people don’t do that very much anymore because it’s much easier to determine the
amino acid composition of a protein if you
have the DNA sequence, and it’s much easier
to sequence the DNA. So I’m not going to talk
about amino acid analysis, and as I said, you won’t
be responsible for that. One of the things that we have
to do in working with proteins let’s say we get
our protein very pure and we want to start
to characterize it, start to understand
better what all is in it, what it’s comprised of, one of the things
that we have to do is we actually
have to take and cut the protein into smaller pieces. Some proteins can be quite large, many have molecular
weights of 200,000 or more, and those are really very
difficult for us to work with some of the methods I’m
going to be showing you about. So it’s desirable, then,
to be able to cut proteins into smaller pieces. And to do this, we use a series
of chemical reagents or enzymes, depending on what
we’re trying to do, that will specifically
break peptide bonds in a protein at specific places. The first one I’ll tell you about
is actually a chemical reagent. It’s called cyanogen bromide. Now, cyanogen bromide has a very
interesting and useful property. When you take and you treat a
protein with cyanogen bromide, what happens is every place
that there’s a methionine residue the peptide bond will be broken. Every place there’s a methionine, the peptide bond will be broken. Well, since methionines
occur in any given protein at specific places, we get
a specific set of fragments that arise from treatment of a
protein with cyanogen bromide. You can see other
reagents that are here. Blah, blah, blah. The only chemical
reagent that I expect that you will know about
is cyanogen bromide. It’s the most commonly used one, and it is very simple, in terms of what it does. In addition to chemical
reagents that we use to chop proteins
into smaller pieces, we commonly use enzymes. And enzymes come from
our digestive system, for the most part, okay? Our body has enzymes
that break down proteins in our digestive system
so that when we eat food and there are proteins in there, we can break those proteins
down into amino acids that we can use for
our own purposes. So some of these that
we use are really useful. One is trypsin. Trypsin is a very
simple one to understand. It cuts on the carboxyl side, and I’ll show you
a figure for this in a second but it cuts on the carboxyl side of lysine and arginine residues. That’s really useful. So again, lysine and arginines
are at specific places in a given protein. By treating
a protein with trypsin, I get a specific set of fragments
arising from that cleavage. We’ll talk more about
trypsin later in the term. Thrombin cleaves on the
carboxyl side of arginine. That’s a very useful tool. So if I compared the
pattern that arose from cutting
a protein with thrombin compared to the
cutting with trypsin, I would guess I would probably
get more fragments with trypsin because trypsin cuts near
two different amino acids whereas thrombin only cuts
near arginine right here. Well, later in the term
I’ll talk about chymotrypsin. I’ll just point out that
it cuts near all of these. But you’ll notice a pattern. This is a benzene ring, a
benzene ring, a benzene ring. So these guys all have
aromatic amino acids that they cleave
next to, right here. In addition, to some extent it
will cleave other amino acids. And the one characteristic you
could notice of all of these, at least of four of the five, is that they are hydrophobic. Tyrosine’s the only one that’s
not very hydrophobic there. Well, again, I do think that you
should know where thrombin cuts. I think you should
know where trypsin cuts. And I think you should know
where cyanogen bromide cuts. The other ones are just sort of, I don’t think it’s really
necessary for our purposes. But you should know that
enzymes are very useful. They’re called
proteolytic enzymes, or they’re called proteases. Proteases cleave peptide
bonds in other proteins. And if the question is, will they
cleave bonds within themselves, the answer is, one protease can
cleave another protease, you bet. And so, if you leave a protease
in a tube over a period of time, it will eventually
lose all of its activity because it cuts itself to pieces. Student: Well, how
do you store it, then? Kevin Ahern: How do you store it? You store it frozen. Student: Oh. Kevin Ahern: Yep. So you isolate it under conditions
where it’s not very active, and then you ship it
and keep it’s frozen so that it’s not able to act. Shannon? Student: Is it true that you can
store it refrigerated if it’s dry? Kevin Ahern: Can you
refrigerate it if it’s dry? It depends on the
protease, but yeah, some of them are
actually shipped dry, but they’re often kept frozen for
that very same reason, as well. Let’s see. I talked very briefly before
about reducing disulfide bonds. I’ll just very briefly
show you here, again. I mentioned that, when
I talked about mercaptoethanol, I said mercaptoethanol
would take a disulfide bond and convert it back
to sulfhydryl groups. I said, at the time, there’s a molecule called
dithiothreitol that will do the thing, and now you see dithiothreitol
doing the same thing. What’s happening is that
this guy is donating electrons to this disulfide bond. So the electrons go here. And when this guy
loses its electrons, it becomes a disulfide bond. The same thing happens
with mercaptoethanol, actually, as it’s reducing
disulfides in a protein. This can be important because, again, if we want to
get the pieces apart, we may want to break the
disulfide bonds of a protein. And, let’s see. No surprise. You’ve had in basic biology, the relationship
of the genetic code, which shows how the sequence of
DNA ultimately can be converted into the sequence of
amino acids in a protein. And, as I mentioned earlier, it’s actually much
easier to determine the sequence of a protein
by sequencing its DNA than to try to determine the
individual sequence of amino acids solely starting from
the protein alone. So DNA sequence is
a much easier way to determine your
protein sequence. Well, I want to spend
some time talking about an immunological technique
that allows us to identify specific proteins
from an SDS gel. So let’s imagine I’ve
got that SDS get that I separated those
proteins on before. And you saw there were
a series of bands that were there, on the side of the gel. One of the questions
you might ask is, “Well, I’m really interested in
a particular protein of my own. How could I tell which
one of those bands is the one I’m interested in?” So one way of doing that is
this technique that’s called “western blotting.” And let me take a few minutes and describe western
blotting to you. To do that, I need
to, first of all, tell you a little
bit about antibodies. Antibodies are proteins
of the immune system that provide protection for
us against outside invaders. And they work by binding
to specific structures. So this is a schematic
diagram of an antibody. It has one end that binds. It actually has two ends that
bind to specific structures. And I’ll describe those
structures in a second. And the immune system, this allows the immune
system to fight off invaders. We’ll talk about the immune
system in 451 next term. But this allows the immune
system to fight off invaders. The reason that we use antibodies in this technique is because
of their ability to bind to only very specific structures. So let’s imagine that I’m
interested in studying a protein that’s a protein from HIV. And I’ve got this mixture of
proteins on the side of my gel, and I really wanted
to know which protein there was the one that was mine. To do this, I would have
had to made an antibody against my protein of interest. So let’s say I’ve got
my purified protein. I’m interested in studying it, using a western blot technique. I would actually inject this
protein into, say, a bunny rabbit or something like that. And the bunny rabbit’s
immune system would see this protein coming in
as a foreign invader. It doesn’t hurt the
bunny rabbit in any way. The bunny rabbit’s immune system
makes antibodies against that. And then I can collect some
blood from the bunny rabbit and isolate those antibodies
that bind to my protein. And that can take
a few weeks to get set up. Just a second, Shannon. I’ve got my antibody
that’s there. And it’s binding
specifically to my protein. Did you have a question? Student: Oh, yeah. Is it possible to
carry this out in vitro? Kevin Ahern: To carry out the
antibody generation in vitro? Student: Yeah. Kevin Ahern: No. The immune system
has to recognize it and then the antibody
has to be synthesized. So it can’t be made in vitro, no. Alright. Now, so I’ve got an antibody
that binds to my protein, alright? That’s the most
important component of what I’m going
to be showing you. And the beauty of an
antibody is it’s specific. It will bind to my
protein but it won’t bind to all the other proteins
I might find in blood, or all the other proteins
I might find in a plant cell or whatever it is that
I’m putting on that gel. It will only bind to the protein
that I’m interested in, ideally. So, I’ve got an antibody
that’s specific for my, it’s called the “antigen.” That’s the thing it binds to. So there’s the antibody. There’s the antigen. There’s the binding. And the binding
can be quite tight, and is necessary for
us to do our analysis. Now, this antibody is going to
be used in this technique called western blotting that
I’m going to show you. So I’ve got my
mixture of proteins. I take my mixture of proteins and I apply them to the
top of an agarose gel, I’m sorry, a polyacrylamide gel. And I separate them. So I’ve got an SDS-PAGE. I’ve separated my proteins. In this case, they’ve actually
cut out the specific band, but you could do the whole
gel, if you wanted to. And I take those
proteins in that gel. Gels are kind of hard to handle. They fall apart real readily, like working with
Jello or something. So I can take and I can
actually use an electric current to transfer those
proteins onto a membrane, Onto a membrane, like a sheet of, sometimes you can use like
a specialized sheet of paper. So you would transfer
all the proteins that’s on there onto
this sheet of paper. And the proteins would
then be stuck to that paper. And we can use techniques to
make them stick quite strongly. So now I’ve got my paper that
was the exact match of my gel, and it’s got those
proteins attached to it. I take the paper and
I transfer it to a bag that has some buffer in it, and I add my antibody. I add my antibody. The antibody is given some time, a few hours, to bind to
what it’s going to bind to. Let’s say my protein
is right there. The other ones aren’t
proteins of interest. The antibody binds and
I take the piece of paper out. I wash it so that all the
things that aren’t bound specifically to my protein, all the other antibodies, come off. And then I use a reagent
that basically tells me, I ask the question,
“Where are the antibodies?” So there are reagents
that will light up color where there’s an antibody. And now, by identifying
the color, I can say, “There’s my protein.” And I can go all the
way back here and say, “There’s where my protein was.” So it’s a very useful technique
for specifically identifying a protein of interest in a
mixture of other proteins. Very, very useful. A very important technique
for me to be able to identify a protein after I’ve
done an SDS-PAGE. So I’m going kind of fast there. I’ll slow down and take any
questions you might have. Yes. Back in the back. Student: So does
the protein, like, do all the other
proteins come off of that? Kevin Ahern: Do all the
other proteins come off? No, they don’t. But the antibody
doesn’t bind to them, so when I treat
to find antibodies, only the one that’s got
antibodies on it will light up. Yes, sir? Student: Could you treat
those antibodies beforehand. Kevin Ahern: So his question was, could I treat the
antibodies beforehand? And the answer is, yes, I could. Sometimes people will take, and it’s actually easy to
put a color onto an antibody, so I don’t have to do
the treatment afterwards. So now I can just look again, and say, “Where’s the color?” There’s a variety of ways of
visualizing this thing right here. Student: Can you
assay the amount? Kevin Ahern: Can
you assay the amount? Western blotting gives
you a rough idea of amount, but it’s not real good
for overall quantitation. But it gives you
a ballpark idea of the amount, yes. Yes, sir? Student: Okay. You’ve identified which one
out of your original SDS-PAGE is a protein of interest. Kevin Ahern: Yep. Student: Is there a way
to dissolve away the gel? Or to isolate the protein
for experimental use? Kevin Ahern:
Oh, very good question. So he says I’ve
identified my protein. Is there a way I can recover
that protein and use it? The answer is, you can
extract a protein from there, but frequently you
won’t want to do that. Anybody know why you
wouldn’t want to do that? Student: You’ve
denatured the protein. Kevin Ahern: You’ve already
denatured the protein. So what’s in there is already
probably not of any use to you if you’re interested in
a protein that’s active. Yes, you can, and commonly what people
will do is take that and analyze it in another way. So they’ll cut it out and use
a technique of spectroscopy to further identify it. So the answer is, yes, you can, but it depends on what
you want to do with it in terms of whether that’s going
to be practical for you or not. Student: But you could
recover it through like crystalize for an X-ray exam? Kevin Ahern: You
would not take this and use it for crystallography or for other analysis
like that, no. No. It wouldn’t be of use to you. But it is of use in other ways. Yes? Student: So after you identify
it and know where it is on there if you want to get more
of it without denaturing it Kevin Ahern: You probably didn’t
put your whole sample on here. So you probably took a
pretty small fraction. Student: And then
you just do it again and you don’t denature
it the next time? Kevin Ahern: Uh, no. Because it’s not denaturing it. You don’t know where
it’s going to go, right? So there’s other things
that you have to do. I’m just showing you one way of
doing that isolation separation. Yes, sir? Student: You talk about
developing antibodies in rabbits. Does that work for
non-animal proteins? Kevin Ahern: Can I葉he question, I think, is, will a rabbit
make antibodies against a non-animal protein? The answer is, yes. They are specific for structures. So they’ll work on proteins. You can make ’em against DNA. You can make ’em against RNA. You can make ’em
against carbohydrate, So it’s structure that’s
the important thing, not the source of the molecule or even the type of
molecule that it is. Good question. Yes? Student: We were talking
the other day about prions. Kevin Ahern: Uh-huh. Student: And you said
that immune systems generally don’t recognize them. What if you did that
cross-speciesation and you injected that like
into a different species and it recognized
it as a non-native… Kevin Ahern: Okay, so, yeah. The question is, that’s
getting a little involved, but basically his question is, can I make an immune
system recognize a prion. The answer is, yes I can. But that doesn’t mean it’s going
to be an effective treatment, and that is what we
were talking about with the immune
system the other day. So, yes, I can make
antibodies against that, but that’s not necessarily
going to be something that’s going to
be useful in terms of helping to treat the disease. In other words, my immune
system is not going to protect me against that protein. So… yeah. Ah-bah-dah. Let’s see here. So let’s spend just
a couple of minutes I’m not too far from
finishing stuff here spend a couple of
minutes talking about, you asked the
question earlier about, can I pull this protein out of
this gel and use it for something? And I can. So let’s imagine I’ve taken
and I’ve identified my band. I have cut out that band. It could either be from a gel, like I showed here, or it could be from a 2D gel. Either way, I could pull out
the band of interest and say, “Okay. Here is my protein. I’ve separated it by
gel electrophoresis. I’d like to know, what is it? What’s the sequence
of it, for example?” Well, I don’t know where
in the DNA it came from. I have to actually sequence, in this case, the protein itself. So I take this purified
protein that I’ve got and I might treat
it with some enzymes to break it into smaller pieces. I might treat it with a
chemical like cyanogen bromide to break it into smaller pieces. And breaking it into smaller
pieces is going to be important because the next technique
I’m going to describe to you works very well on relatively
small pieces of polypeptides. This technique is
called MALDI-TOF. And I’m going to show
you the image first, okay? Oh, blast it. I’ll start with this. MALDI-TOF is a, MALDI,
M-A-L-D-I, stands for “matrix assisted
laser desorption ionization.” You don’t need to know that. Okay? It’s a mouthful. I always have to
look it up each time, so I remember what it is. Matrix assisted laser
disruption ionization. What does that mean? It means this
technique uses a laser to make a sample volatilize. That’s the first part of
what happens in mass spec. How many people have done
mass spec in a chemistry lab? So mass spec tells us mass. And mass specs work
in vacuum chambers. And ions in the vacuum
chamber get accelerated and they get accelerated
to move up to a detector which detects when
things hit them. So MALDI-TOF is a specialized
form of mass spectrometry that allows me to analyze
relatively large molecules, like polypeptides. To do MALDI-TOF, one takes
a protein sample that’s purified, that little band that I had, and mixes it with some material that makes it form
a sort of a crystal. And that would be
on the end of a, let’s say a pinhead that
I could put into a chamber that would be evacuated. So I’ve got my sample
that’s on a pinhead. I put it in this evacuated
chamber and it’s sitting there, waiting for the analytical
process to begin. The “L” said “laser,” and laser plays an important
role in this process. The laser, I thought I had that figure
here and I just don’t see it. Okay. Well, it’s in the book. I’ll post the figure later, to show you. The laser is, there’s
a laser that’s pointed, and the laser hits that sample
in the evacuated chamber. So the evacuated chamber
has my crystallized material. When the laser hits it, that crystallized
sample volatilizes, meaning it leaves the pinhead
and goes into a gaseous phase. In the process of that happening, my sample becomes ionized. It becomes charged. Okay? Because it’s charged, an
electric field will attract it to the detector
at the other end. Okay? Let’s imagine I’ve got
a sample that’s got, let’s say, two things in it. One that has a molecule
with a mass of 500 and one that has a mass of 1,000. And they’re both charged +1. Which one’s going to make it
faster through the chamber? The 500, right? Because it’s got the same charge. It’s got less mass. It’s going to have less inertia, and it’s going to
be accelerated faster and is going to arrive
at the detector faster. It means that, if we measure
carefully the time it takes from volatilization over to the
time the detector detects it, that time interval
actually tells us the size. Because it’ll take something
twice as long that’s 1,000 in molecular weight as it
will for something that’s 500. That’s really useful for us because with that technique we
can determine molecular masses with amazing efficiency. Because of that, we can take a
sample that has a polypeptide, and that polypeptide, when it
ionizes, will break into pieces. The places where it will
break are peptide bonds. So here’s a full-length
polypeptide that’s there. When it ionizes, some of
them will be broken here, and I’ll get one amino acid. Some will be here, I’ll
get two amino acids. Some will be broken here,
I’ll get three amino acids. And if I determine
the masses of all those the difference between
the peaks is the difference of each amino acid that comes off I can actually determine the
sequence of the thing I started with. Yes, it’s complicated and yes, it takes a computer to do it. I’m not going to sit
and stare at it, myself. But the beauty is that,
in a single mass spectrometric analysis, I can determine the sequence of
amino acids of that polypeptide I put into the chamber. Now that is really powerful. Because of this technology, because of this technology, a scientist in a modern
mass spec lab can determine the sequence and, thus, the identity of
4,000 proteins a day. We could take all the
proteins that was on a 2D gel, cut out each spot,
have each one analyzed, have each one identified,
in a single day. That is an incredibly
powerful technique. Question? Student: How long have
they be able to do that? How old is this technology? Kevin Ahern: How what? Student: How old
is this technology? Kevin Ahern: This
technology dates to the ’90s. Yeah. So it’s relatively new. Yes? Student: So why do you, how do you know which
end it starts at, so you have the… Kevin Ahern: Okay. So there are at both ends, and you can actually
get this one, as well. That’s what that
computer has to sort out. Okay? Okay. Kevin Ahern:
A lot of stuff there. Let’s call it a day and
I will see you on Wednesday. Student: [inaudible] Kevin Ahern: I’m sorry? Student: Will we get to talk about photo [inaudible]. Kevin Ahern: We’re not going
to talk about [inaudible]. Sorry. Yeah. [END]

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