B1 Intermediate US 4506 Folder Collection
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Professor Kevin Ahern: Happy Monday!
Monday, Monday, I have a song I'm writing
called Hyundai, Hyundai, I'm sure you can figure out
the tune it goes to.
Hope everybody had a good weekend.
Anyone remember it?
Alright, enough silliness.
We spent some time last time talking about techniques
and today I'm going to spend some additional time
talking about some new techniques,
some of which I think you'll find very medically relevant.
So, hopefully, you'll find this interesting what I have to say.
Last time I was talking I finished up
talking about polyacrylamide gel electrophoresis
and more specifically, I talked about SDS-PAGE and SDS-PAGE
is what we use to separate proteins by electrophoresis
and those proteins we have to manipulate
to make them behave like DNA molecules so we manipulate them
so they behave like DNA molecules and the manipulation
that we did was that we treated them with SDS.
The SDS coated those proteins and it made them lay out flat
like rods and the coating gave them a very negative charge
and the longer that rod was,
the more negative charge it would have.
The common question I get, and I got last time was,
"Well does that negative charge negate any other charge
"that's in that protein?"
And the answer is yes it does because there's a lot more of those SDS's
on there than there are charged residues within that protein.
So that effectively makes that a polyanionic substance
meaning there are multiple negative charges.
And it's not a perfect technique.
Some proteins behave a little anomalously but the vast majority
of proteins behave very well in SDS
and we can see their molecular sizes.
The next figure I want to show you is an example of a protein gel
where on both sides of this is called a ladder
and a ladder simply is a set of proteins of known size
that are there and then you can see a protein being purified
at various steps in the purification process
and there are various quantities of that protein
being present and we can see an estimated size from that.
As I noted last time, this technique really isn't useful
for isolating a protein that we want to use later,
it's more of an analytical technique that tells us
how much protein do I have here, what's its size
and how pure is it compared to the things
I'm trying to purify it away from.
That's what SDS-PAGE is very, very useful for.
SDS-PAGE as we shall see is very useful when combined
with another technique that I'm just getting ready
to talk to you about and so I'll do that in just a moment
but before I do that I'll remind you of the structure of SDS
and no you don't have to draw SDS but this is what it looks like.
It's a long chain fatty acid and these are all carbons
and hydrogens down here and at the end we've got a sulfate
and the sulfate is what gives it
the negative charge that is necessary for electrophoresis.
Well, as I said, if we combine SDS-PAGE with another technique,
we get a really, really powerful technique
and it's a powerful technique for studying things like cancer,
for studying how drugs work and so forth, so I want to spend
a few minutes talking about that because I think
it's a pretty cool and fascinating technique.
The technique I want to discuss first is called isoelectric focusing.
Isoelectric focusing and this technique uses some substances
that are interesting in their structure,
we don't have to talk really too much about them
but suffice it to say that it relies on a mixture of compounds
that have varying amounts of charge.
Some of them are very, very, very negative
and some of them are very, very, very positive
and some of them are in between, so we can imagine,
for example, that we might have a compound that might
have forty negative charges and another compound
in our mixture that might have thirty-nine, thirty-eight,
thirty-seven, thirty-six negative charges and on the other side,
we might have something that has forty positive charges
and thirty-nine, thirty-eight, thirty-seven.
And so, we've got a complete spectrum of charges
that are in this mixture that I have.
Well if I take this mixture and I treat it very carefully,
and I put it into a tube kind of like the columns
that we've been talking about and apply an electrical field
to that tube what will happen is that the things
that are the most negatively charged will move toward
the positive electrode and the things
that are the most positively charge
will move to the negative electrode.
And if I look at that tube when I've finished
doing this electrical treatment of the tube
what I'll see is that the things that had forty minus
will be closest to the end followed by the thirty-nine minus
and the thirty-eight minus,
the thirty-seven minus, etcetera, etcetera.
And in the very middle that's where I'll see things
that had a zero charge.
Make sense?
Okay.
So this tube turns out to be really nice
because if I take this tube and instead of just putting
these compounds in there
what if I put my mixture of proteins in there as well.
Will they separate according to those charges that are in there?
And the answer is yes they will.
The proteins that are the most negatively charged
will be the closest to the positive electrode and those
that are the most positively charged
will be closest to the negative electrode.
And I'll see a spectrum of proteins across that range.
Make sense?
Now these tubes have a sort of gelatinous-like material
in them so what I've done is I've separated
these guys on the basis of charge
and in essence what I've done is I've separated
them on the basis of their pI's.
We don't have to worry about that too much at the moment,
but that's basically what I've done.
Those that have the lowest pI's are on one side
and those with the highest pI's are on the other side.
All right, now, why do I tell you that?
I tell you that because first of all that technique
is a way of separating things by their pI's
but more importantly when I combine that technique with SDS-PAGE,
I get something really, really cool.
So let's imagine we do what I just said, we take that tube,
we take our proteins, we mix them we run that electrical current
so we separate those guys on the basis of their pI
with those with the highest pIs on one side
and those with the lowest pIs on the other side
and the others, a mixture in between.
Now what if we carefully take that tube
and I open the tube up and I take that gelatinous material out
and I lay it on top of a gel where I'm going to do SDS-PAGE?
So I add SDS this material that's in here, I coat it all with negative charges,
It's not starting out with negative charges
because I haven't put anything on it to start with,
I just have the proteins by themselves, I add the SDS to it
and then I run the material on that tube downwards
and separate on the basis of size.
What I've just done is something we describe as 2-dimensional gel
electrophoresis, two dimensions.
The first dimension separated on the basis of pI
and the second dimension separated on the basis of size.
This figure schematically shows you what I just described.
Here are some proteins that have been separated
on the basis of their pI on the top and then we've added SDS
and applied them to an SDS-PAGE gel
and separated them on the basis of,
first on the basis of pI, secondly on the basis of their size.
All right, so what do we see?
We see this gel if we were to look at the upper left
we would see the proteins up in here we would see the proteins
that have the most negative charge
and are the largest.
Down here we would see the ones that have the most negative charge
and are the smallest,
down here the most neutral charge and the smallest, et cetera.
You get the idea as we go across this two dimensional gel.
Well, this turns out to be an extraordinarily powerful
analytical tool as well.
Let me show you what an actual 2-D gel looks like.
An actual 2-D gel looks something like this.
Wow what a mess, right, what a mess this thing is.
Well it turns out this mess is pretty darn cool
because this mess in this case was taken from all
of the proteins in given cell.
I can separate all the proteins
and I can decide using a variety of techniques
which protein corresponds to which spot.
Every protein will have its own unique spot.
Right, that's one thing and second, this type of analysis
if you're very, very careful is very reproducible.
It's very reproducible.
So what does this mean?
Well, let's imagine I want to study a certain cancer
and I want to understand something about that cancer.
I might take and let's say I've got liver cancer.
I might take some cells from my liver that are not cancerous
and I might separate those proteins on a 2-D gel
and then I might take that liver tumor that I have
and I might take those proteins and apply them to a different 2-D gel
and then I would compare the 2-D gels and say which proteins are different?
which proteins are being made in the cancer
but not being made in the regular cell,
which proteins are being made in the regular cell
that are not being made in the cancer cell,
which proteins are being made more in one than the other
or less than one in the other, all kinds of things I can ask
but now as the result of this analysis I can understand
the complete picture of every protein
in a normal cell versus a cancer cell.
I'm not restricted to cancer.
I could say what happens I've got this new drug
that I want to treat people with and I want to understand
what affect this drug has on our cells.
I take one batch of cells over here, don't treat it with the drug
another batch of cells over here,
treat it with the drug, do 2-D gel electrophoresis
and then ask the question, which proteins are different?
That's a really powerful tool, an extraordinarily powerful tool
This type of analysis tells us very quickly
how we're affecting the expression of proteins inside its cells.
Since proteins are the workhorses of cells,
we know very, very quickly something about the cancer,
something about the treatment,
something about your cells versus my cells.
Maybe I'm making more of one protein than your cells are.
You get the idea.
2-D gel electrophoresis is a very powerful tool.
Questions on that, yes up in the top.
Very, very good question.
His question is, if I take a protein that has quaternary structure
and I treat it with SDS, will the subunits come apart.
Let's take a vote, how many people think they will come apart?
How many people think they won't come apart?
Majority rules, the subunits will come apart and there's two reasons.
One, hydrophobic interactions can in some cases help stabilize
a quaternary structure, but even if the quaternary structure
is not being stabilized by those interactions,
when the individual subunits themselves unfold,
it disrupts completely completely the interactions
between the subunits.
So, quaternary structure is completely disrupted
by SDS treatment.
Yes?
No, actually that's not true.
What he said was, "If it denatures, then the individual strands
"are going to run faster than the entire protein."
But remember, the same happens in any kind of denaturation.
When I know I've got a multi-subunit protein,
I know what the individual subunits are.
I do not know what the overall structure is.
Now, your question is very good because if we want to understand
the quaternary structure,
we have to use other kinds of analytical techniques
to understand that.
Sometimes we can be working with a protein and have no idea
that it's a dimer, for example.
If we just simply do SDS PAGE, we won't know
that we've got a dimer of two identical subunits.
We have to use other analytical techniques to do that,
but you're exactly right.
Okay, good questions, yeah?
Are you going to have to tell me
which spot is which, is that what you're saying?
Well, I'm not going to say which protein is that spot,
if that is the question, but I might point to something on here,
and say where would you find, for example,
the largest, most positive proteins?
That would be fair.
Yeah, yeah?
Okay, he says if it is vertical,
does gravity have an effect and no it does not and the reason it
does not is because the electromotive force that is driving it
through there is much stronger than anything gravitational effect it could have.
Okay, good questions, so that's 2D gel electrophoresis,
kind of cool stuff there.
Here's a comparison of normal colon cells versus a tumor.
And you can see,
here's a protein that's made in fairly small amounts
in a normal colon and in a tumor it's made
in very abundant amounts.
You can also see there's probably a difference here and here,
there are some differences we see between them
and there are some very powerful computer tools that will do that analysis
and say how much darker is this spot than that spot
so we can get some pretty good ideas from that.
Okay, well hopefully what I've told you last time is that,
and hopefully you remember, is that we have to do many steps
in purification to get a protein that's ultimately very, very pure.
And you might wonder, "Why do we care so much about
"getting the protein very, very pure?
"Is just fairly pure is that okay?"
Well, for some analyses, fairly pure is okay,
but for many analyses it's not.
So one of the analyses we commonly want to do
when we purify a protein is we want to determine its structure.
Remember we can't predict it from the computer,
so we actually have to determine it and to determine it
we have to have absolutely pure protein.
Techniques that allow us to do that,
I'll briefly mention in a little bit,
include nuclear magnetic resonance and X-ray crystallography
and I'll say something about those two in a little bit.
So those two techniques really require us to have protein
that is very, very pure and we can see on this gel
the analytical use of SDS-PAGE to help us sort of follow
what's happening during the purification process.
Here's the material we got out of the cell.
Here's after we use salt fractionation,
that's something I haven't talked about here,
you're not responsible for it but it's another purification step.
Here's some ion exchange chromatography.
Here's what happened after we did gel,
and then finally after we did affinity we got something
that looked like this.
So we can see our protein started out over here
and we finally got it to a place where it was fairly pure.
One of the things that we need to do
during that purification process is to follow it.
We have to follow it.
We have to test at each step, where's my protein at?
How much of it do I have?
Because if I don't do that, I really won't have any idea
of how much material I'm working with.
Further, I ultimately might like to go back into my cells
and determine how much material is in a given cell,
how many copies of that protein are found in a given cell.
So doing the kinds of analyses we see here in table 3.1
are very commonly done by researchers and they are done
partly also so they don't throw out the baby with the bathwater.
Remember, we're doing a step, let's say, centrifugation,
where we have a precipitate and we have a supernatant.
If we think all the material we want is in the pellet
and all we keep the pellet and throw away the supernatant
that water might have had the baby in it,
so we don't want to do that.
We've got to check everything and make sure that it is there.
Well, this scheme shows a purification going through
the steps you saw on the last figure and this step
in the purification follows the amount of protein we've got.
So we started out with a protein where we were able to determine
we had 15,000 milligrams, that's 15 grams of protein,
that's a lot of protein.
And after we fractionated by salt we ended up with 4,600.
You might say, "well, why do we lose protein?"
We lose protein because we're throwing away things
we don't want and hopefully keeping things we do want,
that's what we saw in the last gel
in fact we saw some of those disappearing.
After ion exchange chromatography we had 1,278 milligrams,
after gel filtration we had 68.8, and by the very end
we had 1.75 but you can recall from that gel
about the only thing on that gel was in fact our protein.
So out of that whole mixture I got 1.75 milligrams of protein.
Now, I have to follow the protein, I have to measure its activity.
So that protein most commonly might be an enzyme,
how much product does it produce per time, that would be a measure
of its activity so I can measure its activity in some unit
that I can decide what that is.
This 15,000 milligrams of protein had 150,000 units
of activity to start with.
This mixture I had to start with had a specific activity
of 10 units per milligram,
simply the number of units divided by the number of milligrams.
The yield I had was 100% because I haven't done anything yet,
I haven't thrown anything away and my purification level
was one because again I haven't done anything to it.
By the second step what I see is that I've reduced
the total protein by a considerable amount,
I've reduced the protein by more than a factor of a third,
it's less than a third of what I started with,
but I still have a pretty good amount of the units.
Now you'll say, "Well how come I don't keep all the units?"
Well, no purification technique allows you to keep everything.
You're going to lose some as you're going along.
So this is pretty good, this is pretty good retention.
We can see in fact, well how much retention was it?
Well it was 92%,
it was equal to the number of units I had here divided
by the number of units the units I started with, that's 92%.
We can see that the result of my purification helped to increase
the specific activity because the specific activity
went from 10 units per milligram up here
to 30 units per milligram up here,
remember this is the units divided by the total milligrams
so I've increased the specific activity by a factor of three
and my purification, therefore, has increased by a factor of three.
If I go all the way down I see I get less and less material
and I still lose material by the very end I've got one third
of my total activity but look how much junk I've gotten rid of,
an awful lot of stuff I don't want,
so I've actually done pretty well right here.
My retention, my yield was thirty five percent which was this number,
52,500 divided by the 150,000, of course times 100 to get percent
and I got thirty-five percent yield, so that was pretty good.
My specific activity went through the roof,
I went from ten here up to 30,000, and 30,000
was units divided by total protein again.
So my purification level went up 3,000,
I purified this guy 3,000-fold.
This is a pretty cool purification that I've done
and each step along the way gave me purer and purer protein
until I finally got down here to the very end.
Okay, questions about that?
Yes, Omar?
Professor Kevin Ahern: I'm sorry?
How do you quantify the purification level?
The purification level is equal to the purification
at any given step divided by what it was when you started,
so it's this 30,000 divided by ten which gives you my 3,000.
Yes?
Professor Kevin Ahern: Is that a pretty typical?
It's going to vary a lot from protein to protein,
I'd say this is a pretty good yield.
Other questions?
Alright, we're going to move from purification
to some characterization, I'm going to skip over a few things
in characterization, there are some things that your book
goes through that are really very old and nobody does anymore
so I don't think there's a reason we should mess with them.
One of the things I do want to talk about
are what are called polypeptide cleavage agents.
Polypeptide cleavage agents are substances
whether they're chemical or enzymes
that can break proteins into pieces.
That can break proteins into pieces.
You might wonder why you want to do that and the answer
to that question is because some of the analytical techniques
don't work very well on great big proteins but they work nicely
on smaller pieces, we'll talk about one in just a little bit.
So being able to break proteins
into specific smaller pieces is really helpful.
As I said there are some of these compounds
that work chemically and some that work enzymatically.
I will tell you about some in both categories
and one of the more common ones used for chemical cleavage
is a compound called Cyanogen bromide and what Cyanogen bromide
will do is it will break a protein on the carboxyl side
of methionine residues.
So we find a methionine in a protein
and we look on the carboxyl side of it,
that's where that Cyanogen bromide is going to break it.
I'm not going to discuss the other chemical agents
which means again I am not going to hold you responsible for those
but I will talk about some of the enzyme ones
because some of the enzyme ones we'll talk about later in the class,
trypsin is one of those.
Trypsin is one of the easier one to remember.
It works on the carboxyl sides of lysine
and arginine residues, so it's an enzyme.
It's actually an enzyme from our digestive system
and that enzyme in our digestive system breaks proteins
in the carboxyl side of lysine and arginine residues.
Another one we're going to talk about later in the term
is thrombin.
Thrombin is invloved in the blood-clotting process and thrombin cuts
on the carboxyl side of arginine.
If I look up here at the carboxyl side of lysine
and arginine residues, what do those two
have in common, anyone remember?
What's that?
They have an NH3 plus on their side chain
and it turns out, that is, in fact, what they're looking at
as they are doing their cleavage.
All these enzymes are examining the R group side chain
of the amino acids and then cutting appropriately.
Chymotrypsin is one we'll talk about,
I'm not going to hold you responsible for the specific amino acids
but I will note that these tend to be fairly nonpolar
and, in this case, they tend to be fairly large sidechains,
and we'll talk a little bit more about that later.
The last one I'll mention is Carboxypeptidase A
and it's a very easy one to remember.
It's the only one that cuts on the amino side
and it acts like a Pac Man.
You are all too young to remember what Pac Man was like
but Pac man of course started at one end
and he just started chewing, chewing, chewing, chewing in.
That's what this guy does.
It starts at the carboxyl end and it chews
in one amino acid at a time.
It says it doesn't work on arginine,
lysine, and proline and it actuallydoesn't work very well on those,
but for our purposes, we'll say it works on all of them
just to make it easy for you.
So it starts on the carboxy end
and it just starts chewing, chewing, chewing, chewing, chewing.
If I leave the Carboxypeptidase A in the presence of a protein
it will completely eat the protein down to amino acids
and you're probably thinking, 'Will it eat itself?
'Will it eat other Carboxypeptidase As?'
And the answer is, yes it will.
So proteases all have that property.
They don't have any distinction between is this a protease
or not a protease and act on it, they will eat up each other as well.
Yes?
Do these proteins become charged after these reactions?
Their charges will not change from what they were,
but we can imagine let's say we have a protein
that had a charge of plus seven and I might cut it
into a protein that has a plus four and a plus three,
for example, depending upon where I cut it.
So depending on where I cut it I'll make subunits, smaller pieces,
that might have individual charges
but that total charge is going to be the same.
We talked about this guy before, I didn't show you the reaction
for it but I talked about DTT
lecture before last, and DTT, I will remind you,
is a reagent that will reduce disulfide bonds to sulfhydryls.
It works like mercaptoethanol does.
In fact, it works identically to the way mercaptoethanol does.
It just starts out with a different structure
but the end result is the same, we get sulfhydryls
where we started with disulfides
and we see that the Dithiothreitol starts out as sulfhydryls
and ends up with disulfides, so DTT works like mercaptoethanol does.
Now I want to spend a little time-
Well actually before I do that I'll talk about-
Obviously you guys know the genetic code is the information
inside of cells that tells the cell
what sequence of amino acids to put together to make a protein.
That information is encoded ultimately in the DNA,
the DNA is converted into messenger RNA,
we'll talk about that next term but suffice it to say
that it is actually way, way easier to determine the sequence
of a protein by sequencing the DNA that codes for it
than it is to try to find the actual amino acid sequence
by breaking down a protein and then analyzing
each one of those amino acids.
That's the way they used to determine the sequence
of a protein and that's some of those older techniques
I was talking about that we're not going to discuss.
They're very, very tedious, they're very, very time-consuming.
To give you an idea to determine the amino acid sequence
by analyzing a protein, amino acid by amino acid, might take a month for one protein.
The way our DNA sequencing technology is today,
we can actually determine the entire genome
of an organism, if it's a bacterium, in a day
and that would include several thousand proteins, in one day.
DNA sequencing really has taken over our analysis of proteins.
So, what I want to turn our attention to at the moment
is a set of immunological techniques we use to analyze proteins
and they're used in combination with SDS-PAGE.
So what I'm going to describe is something called
Western Blotting and I'm going to come to that
in a second and show you.
but before I talk about western blotting
I need to say a little bit about the tool
we use to perform the analysis.
The tools we use to perform the analysis in western blotting
are antibodies and antibodies are proteins of the immune system.
They have a very useful property
in that they're designed to bind to things,
they're designed to bind to structures and more specifically,
they're designed to bind to specific structures.
So let's imagine for a moment I'm studying the protein insulin.
Insulin is a hormone that is found in our body,
and I want to analyze this protein by western blotting
which is what we're getting ready to do.
In order to do that, I would have to have an antibody
that recognized and bound to insulin.
I would have to have an antibody that recognized
and bound to insulin.
Turns out I can make one of those or, more specifically,
an organism can make one of those for me.
How does that occur?
Well, let's say I have a bunch of human insulin that I want to study,
I take that human insulin and I inject it into a bunny rabbit.
Our immune system is set up so that it recognizes invaders
and when it recognizes invaders it synthesizes antibodies
that bind to those invaders.
Since bunny rabbits don't have human insulin,
they have bunny insulin, they see the human insulin
as an invader and they start making gobs of antibodies.
I got a flu shot last week, somebody shot into my arm
a whole bunch of proteins from a flu virus and my immune system
is right now is recognizing those and making antibodies
against those so that when a real flu comes along,
it's going to bind to it and keep me from getting infected.
Well this bunny rabbit that's making my insulin
does this for a while and then I say,
"Okay I can really use those antibodies," and it turns out
to be very easy, I take a little of the rabbit's blood
and then in that blood contains
these antibodies that bind to insulin.
I can purify those antibodies
and now I've got a very powerful tool.
These antibodies will bind to insulin
and usually only to insulin, specifically human insulin.
And that's step one, we don't need to worry about
the various nomenclature, antigens and blah blah blah.
Antigen is simply what the antibody binds
to and this is what the structure schematically
of an antibody looks like.
We don't even need to worry about that too much.
That's really what it looks like more realistically,
instead of schematically.
Here is an antibody binding to an antigen.
So here's that protein that the antibody recognizes
and you can see it binding to it here.
I tell you that because this is essential
for something we call a western blot.
Why do I want to do a western blot?
Well, a western blot allows me to not only separate proteins
but also identify, in that separation which protein
is the one I'm interested in studying.
Let's go through the steps here.
So I've got a mixture of proteins,
I bust open some cells, I say,
"I'm interested in knowing how much human insulin
"is in these cells, or is there human insulin in these cells?"
So I take these proteins, I don't purify them,
I just separate them on an SDS-PAGE system.
I then take that gel that I've just done
and I lay it on top of a membrane.
That membrane is specially designed to bind to those proteins
that are in the gel.
So what I can do is use an electric field
and transfer them out of the gel and on to the membranes.
So now they become stuck to the membrane and that membrane
has all the proteins lined up
just like they were in the original gel.
Everybody with me?
I can take that membrane then and put it into a little Ziploc bag
and yes, we do use Ziploc bags for these sorts of things,
treat it with some buffer and with an antibody
that has been made against that insulin.
Well you can imagine what's going to happen.
The place where insulin is present in this mixture
is where the antibody is going to bind and the other places
which don't have insulin, it's not going to bind to.
I take out that membrane, I wash it off
to get rid of the loose stuff that isn't bound to anything
and then I treat it with something that tells me,
'Where are there antibodies?'
This can be a color reagent, this could be a light reagent
that would make things flash, it doesn't really matter
what the reagent is but we're simply asking the question,
'Where do I find antibodies?'
I get a color, I get a spot,
I get whatever but now I can see where in that original mixture
my protein was, I get an indication of how much I might have
and consequently I've learned something about insulin
in those cells that I was studying.
This is true, I can do this essentially for any protein
that I want so long as I have an antibody
that has been made against it.
Yes, question.
That's a very good question.
How does that antibody recognize that protein
if I've treated it with SDS?
First of all, it doesn't always.
There are sometimes the treatment with SDS
will disrupt that and not let that occur
but the treatments I'm doing here will,
A, partly get rid of the SDS
and second, the antibodies will often times recognize
the primary sequence, not tertiary structure.
So if it recognizes short segments
of sequence, those are always present in the protein
as long as I can expose them and let the antibody get at them,
it will recognize it.
It's not perfect, it won't always do that,
sometimes you'll do it and it just won't show up
but most of the time in fact it will.
Good question.
Other questions?
Clear as mud?
Anybody awake?
Pop quiz then, right?
No, no, no!
Alright, the next thing I want to talk about,
these are all cool techniques and in fact, as I'm going along,
we're seeing these techniques are getting more
and more sophisticated and they're actually getting,
in many cases, newer as well.
What I want to talk about that has really revolutionized
our ability to analyze proteins in recent years
is a technique called MALDI-TOF Mass-Spectrometry.
There's a mouthful of a name.
MALDI-TOF has a longer name than I'm going to go through here,
but it's a technique that involves proteins
within an evacuated chamber
that are accelerated by an electrical field.
Wow there's a mind-boggling kind of thing for you to think about.
What does it look like?
What do we do?
That's not what it looks like, that's what the result looks like.
And that's what the blah blah.
Oh, I'm missing my link, oh blast it!
What did I do?
Alright so I'm going to have to describe it to you.
You guys get to visualize, right?
Visual learners like to visualize,
so you guys are going to be visual learners,
you're going to visualize this.
Let's imagine I have a protein that I want to determine
the molecular weight of.
A protein's a pretty big thing.
A protein might have a molecular weight of 200,000
and I told you some techniques don't work real well on large proteins
so I have to break it into pieces
and I might take a piece of a protein
and analyze the molecular weight of that piece.
Maybe it has a weight of 8,000,
much easier for me to handle.
How would I determine what it's mass is?
Well MALDI-TOF allows me to do that surprisingly precisely.
Surprisingly precisely.
How do I do it?
Well, I take my protein and I put it into a little crystal
and I put it like on the head of a pin.
You with me so far, I got my protein it's in a crystal,
it's on the head of a pin.
This crystal I can volatilize very easily with a laser
meaning I shoot it with a laser,
it turns into vapor very quickly.
Well, if the crystal turns into vapor you can imagine
what happens to the protein, it's no longer bound by the crystal.
So I've got this crystal, I've got this protein,
it's on the head of a pin and I now take this head of a pin
and I stick it into an evacuated chamber meaning it's a vacuum,
envision that, I have this long tube,
I've put my stuff in at one end.
Here in there is the head of a pin,
it's got this crystal and it's got my protein in it.
Then this tube is set up,
A, it has no air in it so it's completely evacuated,
B, at the other end there is an electrical plate
meaning that I can generate an electrical field in here,
three, it's got a laser in there.
So, here's what I do.
I shoot the pin head with the laser,
the material volatilizes and that's time zero.
When the laser hits the pin, I have time zero.
I turn on the electrical field and what happens
when the laser hits the material?
The protein volatilizes in the chamber
now floating and the protein is going to be charged.
It turns out that ionizing a protein
when I shoot the laser at it,
it causes the protein to gain a charge,
it starts out with zero charge and it's gaining a charge.
My electrical field, now I have a negative here
and here I have a positive, guess what's going to happen,
it's going to be pulled by the positive-negative interaction.
It turns out that the time it takes to move
from where it starts to hitting the plate
is related to its molecular weight.
The MALDI-TOF, when I talk about MALDI-TOF the TOF stands
for Time Of Flight.
How long does it take to go from here over to here.
Each time it has a charge of plus one.
Let's see I have something that has a molecular weight of 8,000
and it has a charge of plus one,
it's going to move at a certain rate, right?
Let's say I compare that with something with a molecular weight
of 4,000 and a charge of plus one,
which one is going to move faster?
The smaller one, the weight to mass ratio is smaller
so therefore the smaller one will move twice as fast.
If it had 16,000 it would move half as fast.
Yes?
We can set it up so it has a positive or negative,
so we don't really need to worry about that,
but for our purposes we'll say it has a positive.
So now what's happened is by measuring the time it takes
I can very precisely measure the molecular weight,
and I can even do more really cool things with that
because it turns out that when I hit it with the laser
several things happen to it.
One of the things that happens to it is
that big 8,000 molecular weight piece
doesn't stay 8,000 molecular weight,
it frequently breaks into smaller and smaller and smaller pieces.
If I measure the mass of each of those pieces
I can actually determine the sequence of the protein.
How so?
Well, here's a protein that has a sequence of glutamic acid,
glutamic acid, glycine, methionine, arginine.
If I measure the whole thing it has a mass of 621.
If I lose one from this end,
I lose a glutamic acid, I know the weight of glutamic acid that's lost
the difference between this and this is glutamic acid
and only glutamic acid will have that mass.
I know it had a glutamic acid.
Next it loses another one, next it loses a glycine,
next it loses a methionine, you get the idea.
I can look at the weight of all those fragments
that are in there and say, "Oh, here's the sequence of my protein."
It's a really cool way and a really fast way
to determine the sequence of a protein
if that's what I need to do.
Now you might think, that's kind of tedious.
It is very tedious, it's computer intensive,
very computer intensive.
However, if it's set up properly I can determine the sequence
of 4,000 proteins a day using this technique,
that's why I'm not telling you about the old stuff.
4,000 proteins a day using this technique
because the computer just does it automatically.
There's a little robot that will punch,
I'll take a 2D gel and I'll take one of those spots
and a little robot will take this spot
and put it in the mass spec
and this spot and put it in the mass spec
and this spot and put it in the mass spec, each time determining
the sequence of each one as it goes along, a really cool technique.
So, MALDI-TOF allows us
to do some phenomenal analysis of molecular weight.
Questions about that?
Everybody's kind of blown away by that.
You guys look tired today, should we do a song and then finish early?
I just had a feeling, i don't know, I just had this feeling about that.
Okay, this is a song, I see people leaving
and that's interesting because people who are leaving
probably don't want to have extra credit questions I'm guessing.
Oh ok, there goes your voice, you're losing your voice.
This is a song I wrote for BB 350, but you can substitute 450 and it will still work.
It's called, "This Song's for BB 4-5-0."
[singing] It's one o'clock and Ahern's talkin'
Henderson and Hasselbach and pKa's
and Buffers I should know.
This song's for BB four five oh.
I hope that maybe
he'll think the way we
wrote our answers
wasn't crazy
I really need the
partial credit-so
this song's for BB four five oh!
It's really groovy
that it improves me
watching lectures
in Quicktime movies.
I really need to
go and download those
podcasts for BB four five oh!
I'm feeling manic
I'm in a panic
I'd better study
my old organic.
It has reactions
that I need to know.
This song's for BB four five oh!
I know he said it
that's why I dread it
'cause I skipped Friday's
extra credit
'twil prob'ly haunt me
that lonely zero
grade in BB four five oh!
It could be steric
or esoteric
that carbons get so
anomeric.
I'm too hysteric
better let it go.
This song's for BB three five oh!
[END]
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7. Kevin Ahern's Biochemistry - Protein Purification II

4506 Folder Collection
Scott published on January 14, 2015
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