#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 ***. 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]