BIOCHEM DISCUSSION 2

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Okay. Now we're going to take a look at protein biochemistry and some of the techniques that are used in the laboratory. And I will tell you that a lot of these techniques are not necessarily relevant to biological situations, but they will help you to understand more about the proteins that we're talking about.

The amino acid sequence of tenecteplase, a fibrinolytic agent for the acute treatment of myocardial infarction. [After X. Rabasseda, Drugs Today 37(11):749, 2001.]

Here is a diagram of a protein and it has 527 amino acids, which is not uncommon. Nothing particularly remarkable about this one, except, well, you can see the extensive cross-linking between the cysteine residues in many different places. And sometimes they crosslink between cysteine residues that are close to each other. And sometimes they crosslink between cysteine residues are far away from each other. Obviously, this is far away from each other in sequence, but not in physical space. They have to be right next to each other to be cross-linked.

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CHAPTER 5 Techniques in Protein Biochemistry

As the outline of the chapter, I wanted to point out that the first two statements I don't necessarily agree with, and we'll go into more detail on that. But the last two portions are especially important and we will go into in some detail.

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Chapter 5: Outline

5.1 The Proteome Is the Functional Representation of the Genome

5.2 The Purification of Proteins Is the First Step in Understanding Their Function

5.3 Immunological Techniques Are Used to Purify and Characterize Proteins

5.4 Determination of Primary Structure Facilitates an Understanding of Protein Function

We've talked about the relationship of the DNA to RNA to proteins. And we can see here in this diagram that the proteome represents all of the proteins in a cell or organism, whereas the genome represents the DNA and the transcriptome represents the RNA (and of course, the metabolome at the bottom, but you don't need to worry about that).

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Section 5.1 The Proteome Is the Functional Representation of the Genome

https://en.wikipedia.org/wiki/File:Metabolomics_schema.png

So protein purification is one of those things that is actually very difficult to do. And they don't really mention this so much in the textbook, but getting purified samples of proteins is something that some scientists make their whole career out of. Proteins all have different qualities and they all have different structures. And therefore you have to do a lot of trial and error sometimes to figure out what is the best way to get a pure sample. And sometimes they're not able to get a pure sample, especially in cases of membrane proteins, where they don't dissolve readily in our normal solutions. But let's just talk about if you're trying to get a purified portion of a protein, you need to have some sort of test to know if your protein is in fact pure.

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Section 5.2 The Purification of Proteins Is the First Step in Understanding Their

Function

Learning objective 4: Explain how proteins can be purified.

• Proteins can be purified on the basis of differences in their chemical properties.

– Protein purification requires a test, or assay, that determines whether the protein of interest is present.

One of the most important techniques that's used for this, and for lots of different things, by the way, is spectrophotometry. It's long word, but it just means "colored light measuring". And you take a color of light and you shine it through a sample. And then you have a detector that tells you how much of the light has been absorbed. 2.0 here just being an example. It'll tell you the amount of light that reaches the detector. And the idea is that in this sample you will have your substance that will absorb light. So the more of that substance that you have in that sample, the more light will be absorbed, and the less light will reach the detector. And by measuring that, you can determine how much of whatever that substance is in your sample.

One of the things I'd like to mention at this point is that spectrophotometry is one of the most useful techniques for determining anything. You can use it to measure DNA, RNA. And there's a difference between the two of them. So you can tell which is which, and protein as well. But protein itself does not absorb a lot of light. So usually what they do is what's called a Bradford assay. And I've done a ton of these, which is you have a reagent, it's called a Bradford reagent. And I don't remember what it is right now, {mainly Coomassie G-250} but that doesn't really matter. It reacts with protein, and I believe it reacts with, I don't know, the backbone of the protein or something... {no...it's complicated} anyway, it reacts with protein in a quantitative way. So the more protein that you have in a sample, ... the darker it gets, so the more light it absorbs. And it doesn't matter what the protein is so much. So that's what they use to actually measure the amount of protein in a sample. And a special word about the term "sample". It's actually important because when you are doing these kind of measurements, you're never doing it, I mean, as far as I can recall, ever doing it on all of your protein because a lot of these things will screw up your protein and then you can't use it for anything else. So you take a homogeneous mixture and take a small piece out of that, and that's your sample. And then that's the bit that you test.

So you can also run other assays which can create products that can be detected by spectrophotometry, such as NADH. And they show this here in this lactate dehydrogenase assay. So if you are measuring the absorbance of NADH, NADH itself actually absorbs light, then you can put your lactate in with your lactate dehydrogenase and measure how much NADH gets produced. And that tells you the activity of the protein, or the activity of the sample, let's say.

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Lactate Dehydrogenase

• Proteins can be purified on the basis of differences in their chemical properties. – An assay for the enzyme lactate dehydrogenase is

based on the fact that a product of the reaction, NADH, can be detected spectrophotometrically.

And then you can compare the activity of the sample to the amount of protein in the sample. And get an idea of how much of your sample is actually the target protein that you're looking for. They call that the "specific activity". It's a ratio of enzyme activity, to protein concentration. But that term, specific activity, is not something that you need to be worried about.

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Specific Activity

• Proteins can be purified on the basis of differences in their chemical properties.

– Protein purifications are monitored in part by determining the specific activity of the protein being purified.

– In the case of an enzyme purification, specific activity is the ratio of enzyme activity to protein concentration. Specific activity should increase with each step of the purification procedure.

So why is an assay required for protein purification? Well, it's because you gotta measure it somehow. There's gotta be a way that you can figure it out, figure out what's in there. And the way you do that is with an assay.

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Quick Quiz 1

QUICK QUIZ 1 Why is an assay required for protein purification?

"The purification of a protein is the first step in understanding its function." I disagree with that. A lot of genetic techniques are being used these days to understand the functions of proteins and there's no need to purify them. But if you're going to be doing tests on a protein in a laboratory, then you do need to purify the protein as much as possible so that you can know that the results that you're getting have to do with that protein. And the first step in extracting the proteins from your cells is to disrupt the cells to form a homogenate. And then you centrifuge them and form pellets.

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Proteins Must Be Removed from the Cell to Be Purified

• Cells are disrupted to form a homogenate, which is a mixture of all of the components of the cell but no intact cells.

• The homogenate is then centrifuged at low speed to yield a pellet consisting of nuclei and a supernatant. This supernatant is then centrifuged at a higher centrifugal force to yield another pellet and supernatant. This process, called differential centrifugation, is repeated several more times to yield a series of pellets enriched in various cellular materials and a final supernatant called the cytosol.

(Section 5.2 The Purification of Proteins Is the First Step in Understanding Their Function)

Figure 5.1 Differential centrifugation. Cells are disrupted in a homogenizer, and the resulting mixture, called the homogenate, is centrifuged in a step-by-step fashion of increasing centrifugal force. The denser material will form a pellet at lower centrifugal force than will the less dense material. The isolated fractions can be used for further purification. [Photographs courtesy of Dr. S. Fleischer and Dr. B. Fleischer.]

Up here on the left, you can see the creation of the homogenate. You start with a tube full of cells and fluid, and you grind up the cells using some sort of grinding apparatus. And what that'll do is it'll break down the plasma membranes, and it will disrupt the cellular cytoskeleton, and all of the organelles and everything will be floating around. And it will wreak havoc, basically, on the cell, but it will leave some things intact. And one of those things is usually the nucleus. The nuclei of the cells. Nuclei of cells are very dense and so you put them in a centrifuge, which spins them around and the denser things move to the tip of the tube. And using 500 times the force of gravity up here for ten minutes, it will take all of the nuclei and put them down into the tip of the tube. The remaining part here is called the supernatant. And in this case, you want to take the supernatant then, and spin it again for 10 thousand times the force of gravity for 20 minutes. And then the mitochondria will pellet out into the bottom. And everything else is still going to be floating up in the supernatant. And you can put it again through, at higher, and longer intensity. And you end up getting rid of the microsomes, which are the tiny bits and pieces, the lysosomes, the Golgi apparatus, stuff like that. And in the

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Model of Differential Centrifugation

supernatant here you will have the soluble proteins and of course the salts and any other small molecules, glucose and stuff. Now this is an idealized example. It hardly ever works this cleanly. When you spin down the mixture, you're gonna get a lot of crud in with the nuclear fraction, including a lot of mitochondria, and bits of cytoskeleton, and the proteins that are attached to the cytoskeleton, and the proteins that are in the nuclei. So it's not as clean as it appears to be. Likewise, when you do the second spin, you're getting a lot of crud in here too. And the third spin. So you're going to lose a certain amount of the proteins that you are actually trying to get. But it's one of those situations where we just do the best that we can.

So now I'm going to tell you about chromatography. And I just grabbed this from Wikipedia. It's a laboratory technique for the separation of a mixture. Originally, the mixtures that they were using were mixtures of pigments like chlorophyll, carotenes, and xanthophylls. And you could see where the colors were on your "chromatograph". "Chromo" means color, and "graph" is written. So this is like your "picture of the colors". And you could determine how much of each one of these substances was in your mixture by the intensity of these bands. That terminology, chromatography, has been applied now to a whole bunch of different techniques where the name is no longer appropriate; but don't get confused - it's all just separating out the parts of a mixture.

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Chromatography

Chromatography is a laboratory technique for the separation of a mixture. The mixture is dissolved in a fluid called the mobile phase, which carries it through a structure holding another material called the stationary phase. The various constituents of the mixture travel at different speeds, causing them to separate. ... Chromatography was first employed primarily for the separation of plant pigments such as chlorophyll, carotenes, and xanthophylls. Since these components have different colors (green, orange, and yellow, respectively) they gave the technique its name.

https://en.wikipedia.org/wiki/Chromatography

https://en.wikipedia.org/wiki/Laboratory_technique

https://en.wikipedia.org/wiki/Separation_process

https://en.wikipedia.org/wiki/Pigment

https://en.wikipedia.org/wiki/Chlorophyll

https://en.wikipedia.org/wiki/Carotene

https://en.wikipedia.org/wiki/Xanthophyll

https://en.wikipedia.org/wiki/Chromatography

One thing that they talk about in the textbook is "salting out". I really haven't done anything with this, but by changing the concentrations of various salts in your mixture, by putting salt into the mixture, for example, you can cause certain proteins to precipitate and others to stay in the supernatant. And that's really all you're trying to do is to create two populations. There's one in the pellet and then there's one in the supernatant. And whichever population has the protein that you're looking for, that's the one you keep and you throw the other one away.

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Proteins Can Be Purified According to Solubility, Size, Charge, and Binding Affinity

• Salting out takes advantage of the fact that the solubility of proteins varies with the salt concentration. Most proteins require some salt to dissolve in water, a process called salting in. As the salt concentration is increased, different proteins will precipitate at different salt concentrations, a process called salting out.

If you want to remove salt from a mixture - and like I said, when you do that centrifugation, you're going to have all the salts and small molecules glucose and stuff in there - dialysis is a procedure that is used commonly to remove the salts and small molecules. And because they're so small, you can put them in this membrane. And the membrane has small holes in it and the small things will diffuse out of your solution and into the surrounding solution. But the proteins will not be able to pass through. So you'll end up having proteins in the bag and all the other stuff is equally spread throughout. And you can do the same process again and again to get rid of all these little remaining bits and get a pure mix of protein. It's the same technique that's used when you're dialyzing blood, using it to get rid of the stuff that you don't want in the blood. You send it through a filter and the small things diffuse out, and the large things, like the cells, the blood cells that you need, they stay in and those get put back into your body.

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Dialysis • Proteins can be purified according to solubility, size, charge, and

binding affinity. – The salt can be removed from a protein solution by dialysis. The

protein solution is placed in a cellophane bag with pores too small to allow the protein to diffuse but big enough to allow the salt to equilibrate with the solution surrounding the dialysis bag.

https://www.niddk.nih.gov/health-information/kidney- disease/kidney-failure/choosing-treatment

https://www.niddk.nih.gov/health-information/kidney-disease/kidney-failure/choosing-treatment

So the remaining techniques that we're going to be talking about here are basically variations on filtration. Gel filtration chromatography, they say here, allows the separation of proteins on the basis of size. They talk about a glass column, and that is one way that it is done. It is not the only way that it is done. You can use a gel as well, but we will discuss those more in the future. When you're doing a large- scale separation of proteins, columns are usually the ones that you use for that.

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Separation by Size

• Proteins can be purified according to solubility, size, charge, and binding affinity. – Molecular exclusion chromatography (gel filtration

chromatography) allows the separation of proteins on the basis of size. A column is filled with porous beads. When a protein solution is passed over the beads, large proteins cannot enter the beads and exit the column first. Small proteins can enter the beads and thus have a longer path and exit the column last.

COLUMNS are commonly used for this.

Figure 5.4 Gel-filtration chromatography. A mixture of proteins in a small volume is applied to a column filled with porous beads. Because large proteins cannot enter the internal volume of the beads, they emerge sooner than do small ones.

So in this case we have three columns, (this is a column), and you put your solution in through the top. And when it gets to the column, this is what a close- up of the column looks like. This particular column has carbohydrate polymer beads in it, and small molecules will get stuck in here, will get into the little cracks in the beads. The larger molecules will not, so they will pass through more quickly. And you take serial samples of this solution out of the column, and your initial samples won't have any proteins. And then later on you'll be able to get all of the yellow proteins here, the large ones, into one tube (hopefully), and then the green ones into another tube, the red ones into another tube. Now, you're never really sure which tube they are going to be in, and so you usually will do several tubes, many, many tubes, and then test them to see which tubes have the sample that you're looking for. Once you've got that figured out, then you can narrow it down a little bit and make it a little easier for future filtrations.

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Model of Gel-filtration Chromatography

So ion exchange chromatography is similar, but instead of using beads that separate on the basis of size, it has beads that separate on charge. And the image on the next page will show you more like that.

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Ion-exchange Chromatography

• Proteins can be purified according to solubility, size, charge, and binding affinity.

– Ion exchange chromatography allows separation of proteins on the basis of charge. The beads in the column are made so as to have a charge. When a mixture of proteins is passed through the column, proteins with the same charge as on the column will exit the column quickly. Proteins with the opposite charge will bind to the beads and are subsequently released by increasing the salt concentration or adjusting the pH of the buffer that is passed through the column.

Figure 5.5 Ion-exchange chromatography. This technique separates proteins mainly according to their net charge.

So proteins all have slightly different charges. Some of them are slightly positively charged, some of them are slightly negatively charged. If you put beads in the column that are negatively charged than the ones that are positively charged will stick to those, and the ones that are negatively charged will pass through. So you can separate your different charges based on using this column.

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Model of Ion-exchange Chromatography

Now, affinity chromatography, this is actually one of the most useful and most powerful techniques that people do to get specific proteins.

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Affinity Chromatography

• Proteins can be purified according to solubility, size, charge, and binding affinity.

– Affinity chromatography takes advantage of the fact that some proteins have a high affinity for specific chemicals or chemical groups. Beads are made with the specific chemical attached. A protein mixture is passed through the column. Only protein with affinity for the attached group will be retained. The bound protein is then released by passing a solution enriched in the chemical to which the protein is bound.

Figure 5.6 Affinity chromatography. Affinity chromatography of concanavalin A (shown in yellow) on a solid support containing covalently attached glucose residues (G).

And the idea is that you have a protein that will attach to a very specific target. In this case, this protein attaches to glucose. And so you attach glucose to these beads. And when you send the protein through, the protein that you're looking for will stick to that glucose and the other proteins will all go through. So you'll end up with a column that is full of the protein that you're looking for. Then you add some free glucose and it will compete away the binding sites for glucose on those proteins and release them from the beads, and - into a new tube now - you dispense all of the proteins that you're looking for.

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Model of Affinity Chromatography

So this first sentence here basically is saying that the more surface area you have on your beads, the more proteins can bind to them. And the way to get that surface area as large as possible is to make the beads small. But if you have very small beads, then it becomes harder to get the proteins through. So they developed this technique called HPLC. And you might have heard about that. It's very popular, it's working very well, and it's essentially just the same as your ordinary column, except you're pushing everything through with high pressure.

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High-pressure Liquid Chromatography

• Proteins can be purified according to solubility, size, charge, and binding affinity.

– The resolving power of any chromatographic technique is related to the number of potential sites of interaction between the protein and the column beads. Very fine beads allow more interactions and thus greater resolving power, but flow rates through such columns are too slow.

– High-pressure liquid chromatography (HPLC) uses very fine beads in metal columns and high-pressure pumps to move the liquid through the column. Because of the increased number of interaction sites, the resolving power of HPLC is greater than normal columns.

Figure 5.7 High-pressure liquid chromatography (HPLC). Gel filtration by HPLC clearly defines the individual proteins because of its greater resolving power. Proteins are detected by their absorbance of 220-nm light waves: (1) thyroglobulin (669 kDa), (2) catalase (232 kDa), (3) bovine serum albumin (67 kDa), (4) ovalbumin (43 kDa), and (5) ribonuclease (13.4 kDa). [After K. J. Wilson and T. D. Schlabach. In Current Protocols in Molecular Biology, vol. 2, suppl. 41, F. M. Ausubel, R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl, Eds. (Wiley, 1998), p. 10.14.1.]

And then you can see as the things come out, this assay here will check the absorbance at 220 nanometers, which is just your general absorbance wavelength for protein. And as time goes on, you see some proteins coming out here, and then some proteins coming out here - anywhere there's a spike is a different protein.

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Graph of Gel Filtration by HPLC

And now we get to something that I have done a lot of, is gel electrophoresis. And the idea here is that you put the proteins in a gel, not a column. It's a similar principle, but a gel, much like Jell-O, is a cross-linked mass of polymers - some sort of polymer - and proteins will migrate through that gel, often based on their size. And they're driven through the gel with an electrical field. Now it says that proteins will migrate an electrical field because they're charged. OK... Not all proteins are charged, but some of them are, some of them aren't. But if you take this substance, sodium dodecyl sulfate, SDS, which is this molecule down here at the bottom, and you mix that in with your protein and heat it up, it will denature the proteins and it'll stick to the backbone of the protein. At least that's the general belief. That's what people think. The actual detailed, atomic nature of all this stuff is really actually not known. But in general it sticks to the protein, one molecule of SDS for every two amino acids. And so the longer the chain is, the more SDS will stick to it. And each SDS molecule has a negative charge. So these proteins become more and more negative. And then you can push them through the gels using an electric current. They say here it's on the basis of mass only. Well, that's not exactly true. The mass of the proteins can be different depending on the amino acids in the protein. But it generally works out, because proteins are made up of a jumble of amino acids, that the higher mass proteins move through more slowly.

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Proteins Can Be Separated by Gel Electrophoresis and Displayed

• Proteins will migrate in an electrical field because they are charged. When the migration occurs in a gel, the process is called gel electrophoresis.

• Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) allows accurate determination of mass. SDS denatures proteins, and for most proteins, 1 molecule of SDS binds for every two amino acids. Thus, proteins have the same charge-to-mass ratio and migrate in the gel on the basis of mass only.

Figure 5.8 Polyacrylamide-gel electrophoresis. (A) Gel-electrophoresis apparatus. Typically, several samples undergo electrophoresis on one flat polyacrylamide gel. A microliter pipette is used to place solutions of proteins in the wells of the slab. A cover is then placed over the gel chamber, and voltage is applied. The negatively charged SDS (sodium dodecyl sulfate)–protein complexes migrate in the direction of the anode, at the bottom of the gel. (B) The sieving action of a porous polyacrylamide gel separates proteins according to size, with the smallest moving most rapidly.

So here's an apparatus that one would use to do electrophoresis. There's a tank of fluid at the top and the bottom, and then the gel is in between them. Now, the gel also is full of fluid. So there's a complete circuit through the apparatus from the negative pole to the positive pole. And using a small pipette you load your sample into one of these little tiny wells, you turn on the electricity, and it'll push everything through the gel. And they show, as I mentioned, the smaller proteins will move faster than the larger proteins.

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Diagram of Polyacrylamide-gel Electrophoresis

Figure 5.9 The staining of proteins after electrophoresis. Proteins subjected to electrophoresis on an SDS–polyacrylamide gel can be visualized by staining with Coomassie blue. The lane on the left is a set of marker proteins of known molecular weight. These marker proteins have been separated on the basis of size, with the smaller proteins moving farther into the gel than the larger proteins. Two different protein mixtures are in the remaining lanes. [Wellcome Photo Library.]

This is an example of one of those gels that has been stained with a dye called coomassie blue. It just stains the proteins. And the proteins at the top would be the longer proteins and the proteins at the bottom would be the shorter ones. And the intensity of the band can tell you how many of the proteins are there. So looking at this, I can see that this band here, it's very intense, whereas this one is not. And these smaller bands are actually also proteins. So one of these faint bands, there's very little of that protein there.

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Image of Stained Proteins After Electrophoresis

Proteins separated by SDS–PAGE are visualized by staining the gel with dyes such as Coomassie blue.

Figure 5.10 The principle of isoelectric focusing. A pH gradient is established in a gel before the sample has been loaded. (A) The sample is loaded, and voltage is applied. The proteins will migrate to their isoelectric pH, the location at which they have no net charge. (B) The proteins form bands that can be excised and used for further experimentation.

Isoelectric focusing is a different method which I personally have never used. And this is more a proteomics based method. But the idea is that proteins have different charges, and they can settle in at different pHs. So you have a gel here at the top, in A, where the pHs are different. So the high pH areas over here have fewer protons, so those would be more negatively charged. And the low pH areas have a positive charge, like a lot of protons hanging out there. And so when you put them in, they basically would protonate the proteins. So a protein that can be protonated will end up migrating to the right. A protein that is not being protonated will migrate to the left. And they will continue to migrate until they reach a point where they have no net charge.

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Model of Isoelectric Focusing Isoelectric focusing allows separation of proteins in a gel on the basis of their relative amounts of acidic and basic amino acids. If a mixture of proteins is placed in a gel with a pH gradient and an electrical field is applied, proteins will migrate until they reach their isoelectric point (pI), the pH at which they have no net charge.

Figure 5.11 Two-dimensional gel electrophoresis. (A) A protein sample is initially fractionated in one direction by isoelectric focusing as described in Figure 5.10. The isoelectric-focusing gel is then attached to an SDS–polyacrylamide gel, and electrophoresis is performed in the second direction, perpendicular to the original separation. Proteins with the same pI value are now separated on the basis of mass.

That can be combined with two-dimensional electrophoresis. And the first step is an isoelectric focusing, and the second step is a regular SDS polyacrylamide gel, which separates them on basis of size. So these are different "dimensions" of the protein, and they are independent. So the isoelectric point of this protein is where it is here. And then the sizes of the proteins that are contained in that field are here. So this is a mixture of proteins that are all at the same isoelectric point. And then you can separate out that mixture by their length, using regular gel electrophoresis.

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Diagram of Two-dimensional Gel Electrophoresis

In two-dimensional gel electrophoresis, proteins are separated in one direction by isoelectric focusing. This gel is then attached to an SDS–PAGE gel, and electrophoresis is performed at a 90° angle to the direction of the isoelectric focusing separation.

Figure 5.11 Two-dimensional gel electrophoresis. (B) Proteins from E. coli were separated by two-dimensional gel electrophoresis, resolving more than a thousand different proteins. The proteins were first separated according to their isoelectric pH in the horizontal direction and then by their apparent mass in the vertical direction. [(B) Courtesy of Dr. Patrick H. O’Farrell.]

And this is kind of what that would look like. And you can see the isoelectric focusing would occur on the top, and the proteins would be arranged by their isoelectric points. And then SDS page would be applied. And the proteins that are higher up are the proteins that are longer. Now the effect of this is that you can see pretty much a different spot for every protein. Now of course, there's some overlap, but it's quite remarkable. Each one of these spots represents a different protein. So you can get an overall view of all of the proteins inside your sample using this technique.

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Image of Two-dimensional Gel Electrophoresis

Figure 5.12 Alterations in protein levels detected by two-dimensional gel electrophoresis. Samples of (A) normal colon mucosa and (B) colorectal tumor tissue from the same person were analyzed by two-dimensional gel electrophoresis. In the gel section shown, changes in the intensity of several spots are evident, including a dramatic increase in levels of the protein indicated by the arrow, corresponding to the enzyme glyceraldehyde-3-phosphate dehydrogenase. [Courtesy of Lin Quinsong © 2010, The American Society for Biochemistry and Molecular Biology.]

And then you can compare two different dots from, for example, cells that are normal and cells that are cancerous. And you can see here that this big dot is more or less the same in both of them. This dot here's more or less the same in both of them, but these two dots and this dot have increased. So that means that in the tumor tissue, there is a lot more of this protein.

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Images of Alteration in Proteins Levels Detected by Two-dimensional Gel

Electrophoresis

Now, immunological techniques are used to purify and characterize proteins. Immunological techniques are among the most powerful and widely used techniques. And I mean, we could focus most of our time on that. I've used them a lot and we're gonna study them. But we're going to start out with this talk about estradiol and the estrogen receptor, which is interestingly not actually an immunological technique, but it'll introduce us to some of the ideas. So estrogen receptor is a protein that binds to the steroid hormone estradiol. And it binds very, very strongly. In fact, this is one of the most strong binding, most specific interactions between proteins and their hormones that are known. So this is a good example for this. You put in estradiol into a mixture of proteins, it will only stick to the estrogen receptor and it will stick strongly. So we were talking earlier about assays that you would do. So if you were going to do an assay to see if you had the estrogen receptor in your sample, how could you do it? And it has no enzymatic activity. So this estradiol method is another way that you can use to look for it. And we use gradient centrifugation.

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Section 5.3 Immunological Techniques Are Used to Purify and Characterize Proteins

Learning objective 5: Explain how immunological techniques can be used to purify and identify proteins.

• The estrogen receptor binds the steroid hormone estradiol tightly and with great specificity.

• The estrogen receptor has no enzymatic activity but can be purified by immunological techniques and the use of gradient centrifugation.

Now we've talked about centrifugation already. Ultracentrifugation is just really fast centrifugation and it is able to separate proteins by their... not necessarily their size, not necessarily their density, but by something which is what they called the "sedimentation coefficient". And there is a mathematical formula that we can use to determine that, but it's not really that important. Not many scientists even really know what that is. But all we know is that certain proteins have a certain place that they tend to settle in a gradient, in gradient centrifugation. And that gives them this number, "S", the Svedberg units. The larger the Svedberg units, the farther it will move in the centrifugal field.

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Centrifugation Is a Means of Separating Proteins

• Ultracentrifugation can be used to examine proteins. When subjected to a centrifugal force, the rate of movement of the particle is defined by the sedimentation coefficient, s.

• Sedimentation coefficients are usually expressed as Svedberg units (S) equal to 10−13 s.

And it starts off with a density gradient.

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Gradient Centrifugation Provides an Assay for the Estradiol–Receptor Complex

• A density gradient is formed in a centrifuge tube, and a mixture of proteins in solution is placed on top of the gradient.

• To identify the estradiol receptor, the protein mixture is first incubated with radioactive estradiol, which is readily detected. Only the estradiol receptor will bind to the steroid. Moreover, the steroid alone is too small to be influenced by the centrifugal force.

• After the centrifugation is complete, a small hole is made in the bottom of the centrifuge tube and portions of the gradient are collected and tested for radioactivity.

Figure 5.14 Zonal centrifugation. The steps are as follows: (A) form a density gradient, (B) layer the sample on top of the gradient, (C) place the tube in a swinging- bucket rotor and centrifuge it, and (D) collect the samples. [After D. Freifelder, Physical Biochemistry, 2d ed. (W. H. Freeman and Company, 1982), p. 397.]

And I will just show you here what a density gradient is. And I have actually made these myself. It starts with two things, low density and high density, and a mixing bar that mixes them up. And the high density solution tends to go to the bottom faster and then slowly fills up. And the low density solution fills up to the top and you end up with something that's dense at the bottom and {gradually changes to} not as dense up at the top. And when you spin this in a centrifuge, that density gradient doesn't really change because the denser stuff is already at the bottom, the not so dense stuff is at the top. But if you put something on top of it, then it will go down and find its place in that gradient. And it will stop moving when it gets to that density, which is where the Svedberg units are.

[Go back to previous slide, #32] So first you do radioactive estradiol. And just a note - why do we use radioactive stuff? Because radioactive stuff is exactly the same chemical composition as the original stuff. The radioactive nuclei of an atom does not change its chemical

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Diagram of Zonal Centrifugation

properties. So you can use radioactivity to mark things without changing how they behave. And that's very important. So what you do is you put the radioactive estradiol in there, and it binds to the estrogen receptor.

[Back to slide #33] And then you put your whole sample on there and it gets separated out. And this is an example. There's three different bands in this tube that are being shown. And only one of them is the estrogen receptor, which has the Svedberg units which makes it come to right there, and then it gets separated out. So you do this similarly as you did with the columns. You just drip it out - you put a hole in the bottom and then ten drips into here, ten drips into here, ten drips into here, starting from the right in this case for - I don't know why.

Figure 5.15 Gradient-centrifugation analysis of the estradiol–receptor complex. The receptor protein bound to radioactive estradiol migrates into the gradient on centrifugation. Although some unbound estradiol diffuses into the gradient, the steroid does not migrate to any significant extent. Abbreviation: DPM, disintegrations per minute.

But the idea is that the tubes will have different amounts of your target substance. And in this case, the red one is the estrogen receptor that we're looking for. And you can see the radioactivity here is highest here in the middle where the estrogen receptor is. And so you can just use it at the detector. And the steroid only, of course, will also show up with radioactivity. But that'll be right at the top of the {centrifuge} tube because it's very small and it doesn't migrate. So there will be radioactivity in here. But you're like, oh, that's not what we need. You'll see radioactivity here. That is definitely the tube that contains our estrogen receptor. And then there wouldn't be as much radioactivity in there. So that's one way of separating these.

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Diagram of the Gradient Centrifugation Analysis of the Estradiol–receptor Complex

Alright, so now we're getting to the actual immunological techniques here when we start talking about antibodies. And the idea is that antibodies to specific proteins can be generated - and are generated in your body all the time. So there are actually lots of antibodies in your blood right now, and most of them are just sitting around doing nothing because you're not infected with anything. But if you get infected with something, then one of those antibodies will stick to that thing because it's a foreign substance. And the antibodies are more or less random. But once one of them sticks to something, that particular antibody gets amplified and starts being made in large quantities in order to fight off the infection. They talk about "an antibody is a protein synthesized" - it's synthesized in your body. It's not synthesized in a lab, and it represents a particular structural feature. The antigen is the thing that is the foreign substance, and the epitope is the structure that the antibody recognizes. And the idea is that the antibodies don't recognize any of the structures that are normally in your body. So you would all have, we all have epitopes, but the antibodies don't recognize those because those are our own and they are protected.

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Antibodies to Specific Proteins Can Be Generated

• An antibody is a protein synthesized in response to the presence of a foreign substance called an antigen.

• The antibody recognizes a particular structural feature on the antigen called the antigenic determinant or epitope.

Figure 5.16 Antibody structure. (A) Immunoglobulin G (IgG) consists of four chains, two heavy chains (blue) and two light chains (red), linked by disulfide bonds. The heavy and light chains come together to form Fab domains, which have the antigen- binding sites at the ends. The two heavy chains form the Fc domain. Notice that the Fab domains are linked to the Fc domain by flexible linkers. (B) A more schematic representation of an IgG molecule. [Drawn from 1IGT.pdb.]

So this is the structure of the antibody, and you'll usually see it in this form, which looks like a Y. The end bits here are the ones that bind to the epitopes. And it's four chains connected by disulfide bonds. And this is a more realistic look at what it would look like.

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Structure of an Antibody

Figure 5.17 Antigen–antibody interactions. A protein antigen—in this case, lysozyme—binds to the end of an Fab domain of an antibody. Notice that the end of the antibody and the antigen have complementary shapes, allowing a large amount of surface to be buried on binding. [Drawn from 3HFL.pdb.]

And just taking a look at this portion right here, which is the epitope-recognizing, antigen-recognizing site. Here would be the antigen, and it has a structure with atoms in a particular shape and the antibody fits that shape. And so when they come together, they bind tightly. And that's the idea.

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Model of Antigen–Antibody Interactions

Now each antibody-producing cell in your body only produces one type of antibody. So you have a ton of different cells that are producing slightly different antibodies. When you only have one type of antibody, that's called "monoclonal". And if you have a whole range of different antibodies, that's called "polyclonal".

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Monoclonal vs Polyclonal Antibodies • Antibodies to specific proteins can be generated.

– Any antibody-producing cell synthesizes antibodies that recognize only one epitope. Each antibody-producing cell thus synthesizes a monoclonal antibody.

– Any antigen may have multiple epitopes. The antibodies produced to the antigen by different cells are said to be polyclonal.

Figure 5.18 Polyclonal and monoclonal antibodies. Most antigens have several epitopes. Polyclonal antibodies are heterogeneous mixtures of antibodies, each specific for one of the various epitopes on an antigen. Monoclonal antibodies are all identical, produced by clones of a single antibody-producing cell. They recognize one specific epitope. [After R. A. Goldsby, T. J. Kindt, and B. A. Osborne, Kuby Immunology, 4th ed. (W. H. Freeman and Company, 2000), p. 154.]

So if you look at a single antigen, you would get perhaps, in this case, three different antibodies that are produced in response to the three epitopes here - the red triangle, and the blue square, and the gray circle or whatever. So each one of these antibodies would be produced by a different cell, and would recognize only that epitope. But all together, if you're looking for a specific protein, polyclonal antibodies are ... very useful because they have multiple antibodies in them that recognize multiple epitopes. And so they will do a better job of finding that antigen. Monoclonal antibodies, though, are more specific, and they only have one type of antibody.

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Model of Polyclonal and Monoclonal Antibodies

So this here says monoclonal antibodies with virtually any desired specificity can be readily prepared.

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Monoclonal Antibodies with Virtually Any Desired Specificity Can Be Readily Prepared • Immortal cell lines

producing monoclonal antibodies can be generated by fusing normal antibody- producing cells with cells from a type of cancer called multiple myeloma.

• A monoclonal cell line is isolated by screening for the antibody of interest.

DID YOU KNOW? Clinical laboratories use monoclonal antibodies in many assays. For example, the detection in the blood of enzymes that are normally localized in the heart points to a myocardial infarction (heart attack). Blood transfusions have been made safer by antibody screening of donor blood for viruses that cause AIDS, hepatitis, and other infections diseases. Monoclonal antibodies also find uses as therapeutic agents. Trastuzumab (Herceptin), for example, is a monoclonal antibody useful for treating some forms of breast cancer.

Not true! The creation of monoclonal antibodies is actually a really big business. And anybody who's used antibodies extensively will tell you that a lot of the time they don't even work, or they don't work well, and it's often very hard to get one that does what you want. It's a big business. So I don't know who said this, but that's actually probably somebody who didn't have a lot of experience working with actual antibodies. But the idea behind it is fair enough to say. Immortal cell lines producing monoclonal antibodies can be generated by fusing normal antibody producing cells, and we'll talk about that in this next thing.

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Monoclonal Antibodies with Virtually Any Desired Specificity Can Be Readily Prepared • Immortal cell lines

producing monoclonal antibodies can be generated by fusing normal antibody- producing cells with cells from a type of cancer called multiple myeloma.

• A monoclonal cell line is isolated by screening for the antibody of interest.

Figure 5.19 The preparation of monoclonal antibodies. Hybridoma cells are formed by the fusion of antibody-producing cells and myeloma cells. The hybrid cells are allowed to proliferate by growing them in selective medium. They are then screened to determine which ones produce antibody of the desired specificity. [After C. Milstein. Monoclonal antibodies. Copyright © 1980 by Scientific American, Inc. All rights reserved.]

First of all, the way this is done - and this is done usually in a mouse if you want monoclonal antibodies, or a rabbit if you want polyclonal antibodies - ... and I'm not sure what the reason is for that. But you inject the antigen into the mouse - the protein that you're trying to get the antibody to, like the estrogen receptor, for example. And then the mouse will recognize that as foreign and it will start producing antibodies to it. And the cells that produce the antibodies are in the spleen. So then you take those cells that are producing these antibodies - of course, you don't know which one is which - and you fuse them with the cell culture from a myeloma line. Myeloma is a cancer, so these cells are considered immortal because they will continue to reproduce. That's what cancer does. They just reproduce uncontrollably. And then you can combine those two and create these hybrid cells. And then using these cells, you will separate them out and put them into different wells in a 96 well plate or whatever and grow them, check them, screen them to find out which one of these cells is making the antibody that you want. After

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Diagram of the Preparation of Monoclonal Antibodies

you find that, you take those cells and you grow them up, and you can get the antibodies directly from those cells in culture, or I guess you can inject them into another mouse and induce tumors. That sounds awful, but I guess people do that. Anyway, moving on...

Alright. So the estrogen receptor can be purified by immunoprecipitation. And this is a technique that is very common and is used all the time. So this is something that, if you're interested in this type of stuff, you should know about. So you start off by creating a monoclonal antibody for the estrogen receptor using the methods that we just talked about - injecting it into a mouse, et cetera, et cetera. Find the cell that produces the antibody, and you grow up a bunch of antibodies.

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The Estrogen Receptor Can Be Purified by Immunoprecipitation

• A monoclonal antibody for the estrogen receptor can be isolated by searching for cell lines that produce an antibody that binds to the receptor.

• If an antibody for the receptor is present, it will bind to the receptor and alter the sedimentation constant of the receptor.

Figure 5.21 Purification by immunoprecipitation. Monoclonal antibody to the estrogen receptor is added to a preparation of cytosol containing the receptor. The antibody is attached to an insoluble bead. The mixture is then gently stirred for several hours to allow the interaction of the receptor and antibody. The mixture is centrifuged, and the supernatant is discarded. The pellet is resuspended in buffer to wash out any trapped cytosolic components. The pellet is again collected and the supernatant discarded. The antibody–receptor complex is treated with a denaturant. On centrifugation, the supernatant contains pure estradiol receptor.

And then you attach those antibodies to beads, just like we had the glucose attached to the beads in a previous slide. We have a bunch of beads with these antibodies stuck to it. And so what's going to happen then is the estrogen receptor will stick to the beads, mix them together, and you end up with beads attached to estrogen receptors, which makes them very large complexes. And you can centrifuge them down and they all go down to the bottom. Then you discard the supernatant ... and you can resuspend this, and you might want to fill it with fluid again and then centrifuge it again just to get rid of all the crud that's down there. But eventually you end up with a pretty pure solution of beads attached to the estrogen receptors. Then you add a denaturant, which denatures all of the proteins. And remember, an antibody has a specific structure that is necessary for it to recognize its antigen. So if you denature everything, then they're not gonna stick

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Diagram of Purification by Immunoprecipitation

anymore. So that will separate the estrogen receptor from the beads. Centrifuge again, and the beads, now, all go down to the bottom, and the estrogen receptor stays floating in the mix. So you discard the pellet, keep the supernatant, and now you have a pretty pure collection of your estrogen receptor. Now one possible problem with this is that now you've denatured everything. The estrogen receptor might not fold back into the correct form and therefore it will be inactive. But, you know, maybe you don't need to know that. I don't know. I don't know specifically how that's going to work with this.

Okay. So building on this concept, we can use antibodies to detect and quantify the amount of protein in your sample. And one of the ways that's done is what's called ELISA - Enzyme-Linked ImmunoSorbent Assay. And this is a technique that is used a lot, but I don't know that it really has any relevance to your biology. But I think it's used in all kinds of applications like criminology, {HIV diagnosis, pregnancy detection, etc.} Anyway, so here's the idea.

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Proteins Can Be Detected and Quantified with the Use of an Enzyme-Linked

Immunosorbent Assay • Antibodies are used as a reagent to determine the amount

of a protein or other antigen present. Enzyme-linked immunosorbent assay (ELISA) quantifies the amount of protein present because the antibody is linked to an enzyme whose reaction yields a readily identified colored product.

Figure 5.22 Indirect ELISA and sandwich ELISA. (A) In indirect ELISA, the production of color indicates the amount of an antibody to a specific antigen. (B) In sandwich ELISA, the production of color indicates the quantity of antigen. [After R. A. Goldsby, T. J. Kindt, and B. A. Osborne, Kuby Immunology, 4th ed. (W. H. Freeman and Company, 2000), p. 162.]

We're going to talk about two different kinds. There's actually more than just two. There are several. But let's start with the indirect ELISA here. Put the antigen that you're looking for at the bottom of the well. So you put in your protein sample and then you add your specific antibody. And this is the monoclonal antibody that you had created. And so - this is just a trick that you do with antibodies a lot, and I'll explain that in a second. But you add an enzyme- linked antibody that binds to that specific antibody. And then you use a substrate that the enzyme can process into a colored product. And then you can measure how much of that colored product there is, and therefore determine how much of your protein there is in the sample. So why do they do this this way? Why can't they just take a monoclonal antibody and attach the enzyme directly to that? Well, they could actually, but it's a pain. It's a lot of work to actually attach an enzyme. And the process is not perfect. And you've got your monoclonal antibodies, and sometimes these monoclonal antibodies are really precious - like, if you lose the cells that make them, you might never be able to make the same antibody again. It's a random

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Diagram of Indirect and Sandwich ELISA

process. They all develop in different ways. And so it's very difficult to predict what's going to happen. So these antibodies can be very precious, but these are a mouse antibody. ... So this part, the end part here, it's always the same in all of the monoclonal antibodies. It's only the tips that are different. So if you get an antibody that attaches to this part, then you can make a ton of that antibody and attach enzyme to all of that, which is tough, but you only need to do it once, and then you can use that enzyme- attached antibody to detect the mouse antibodies, any kind of mouse antibodies that you want. So it just makes it a lot easier and cheaper for everything to work. Another way here of doing this is a sandwich ELISA, where you coat the well with your monoclonal antibody. Then you put your antigen into there, your sample, and the antigen sticks. And then you use a second monoclonal antibody linked to an enzyme ... I don't know, that's kinda weird. And then you use that to detect it. I mean, these techniques are really not that important to know, but it's just important to know that they exist because they are very commonly used.

And that brings us to Western blotting. This is a technique that is very near and dear to my heart. It is something that I've done a lot of, so much so that it's probably the technique that I know better than anything else. And so I can describe it to you.

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Western Blotting Permits the Detection of Proteins Separated by Gel Electrophoresis

• In western blotting or immunoblotting, proteins are separated in an SDS–PAGE gel, transferred to a sheet of polymer, and then stained with a fluorescent antibody.

Figure 5.23 Western blotting. Proteins on an SDS–polyacrylamide gel are transferred to a polymer sheet and stained with fluorescent antibody. The fluorescent antibody is excited by light, and the band corresponding to the protein to which the antibody binds is visualized with an appropriate detector.

I don't think this is a great example, a great diagram of it, but it'll do. So - we've talked about gels and - denature your proteins, load it into a gel with SDS, and push it through with an electric current, and the proteins will separate based on their length. Primarily. And each protein, hopefully, will separate into a different band. And that way you can tell them all apart. Sometimes proteins have very close, similar weights or sizes, and they overlap, and that can be a pain, but in general it doesn't matter too much. And then you take and use an electric field to push all of those out of the gel sideways onto a polymer sheet. And now you've got all your proteins stuck to this piece of paper, basically. And that's important because when they're inside the gel, they're hard to get to. If you wanted to use an antibody to detect them, the antibodies can't really get into the gel. But once they're pushed onto this polymer sheet, the antibodies can attach to them because they're all kinda just sitting on top of the sheet. You mix it with your antibody that you got, your specific antibody, and then wash it all off and only the protein that you're looking for will end up with antibodies stuck to it. Then you would use, normally, a secondary antibody, again {like in indirect ELISA}, that is specific to all of the mouse antibodies, and

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Diagram of Western Blotting

that has a fluorescent probe attached to it. This is just generally how it's done. You can also use other techniques, but we're talking here about fluorescent probes. You can see over here the fluorescence. So that's what we're going to be talking about. And then you've got your protein with the antibody stuck to it, with the fluorescent probe attached to that. And then you can look at it and under fluorescent light and you can see where your protein is. And you can also tell how much of it there is. And that's the most important thing a lot of the time. If you're going to want to know whether your protein of interest has been increased or decreased in amount, then this technique is really good. So I think that's essentially it - separate the proteins, put them on a piece of paper, use an antibody to attach to the protein you're looking for, and then a secondary antibody with a fluorescent probe attached to it. And then you can look at that under fluorescent light and see where your protein is, and what size it is, and also how much of it there is.

So determination of primary structure facilitates an understanding of protein function. And I totally agree with that. I think that's a very good point. And I'll talk about the primary structure, which is the amino acid sequence. And yes, very important. The rest of this I do not agree with. "A preliminary step is to determine the amino acid composition of the protein" - as far as I know, nobody really does that.

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Section 5.4 Determination of Primary Structure Facilitates and Understanding of

Protein Function • A key step in understanding protein function is to

determine the primary structure, or the amino acid sequence, of the protein. A preliminary step is to determine the amino acid composition of the protein.

• The protein is hydrolyzed, and the constituent amino acids are separated on an ion-exchange column. The amino acids are visualized by reaction with fluorescamine.

There is this thing called Edman degradation, where you take and determine what the amino acid at the end of the protein is, and then cut it off, then determine what the next one is, then cut it off, and it's a process that goes on and on like that until you know what the primary sequence is.

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Edman Degradation

• The amino acid sequence can be determined by Edman degradation.

They have special reagents that cleave protein chains in various places.

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Table 5.3 Specific Cleavage of Polypeptides

Reagent Cleavage site Chemical cleavage Cyanogen bromide Carboxyl side of methionine residues

O-Iodosobenzoate Carboxyl side of tryptophan residues

Hydroxylamine Asparagine-glycine bonds

2-Nitro-5-thiocyanobenzoate Amino side of cysteine residues

Enzymatic cleavage Trypsin Carboxyl side of lysine and arginine residues

Clostripain Carboxyl side of arginine residues

Staphylococcal protease Carboxyl side of aspartate and glutamate residues (glutamate only under certain conditions)

Thrombin Carboxyl side of arginine

Chymotrypsin Carboxyl side of tyrosine, tryptophan, phenylalanine, leucine, and methionine

Carboxypeptidase A Amino side of carboxyl-terminal amino acid (not arginine, lysine, or proline)

Figure 5.27 Overlap peptides. The peptide obtained by chymotryptic digestion overlaps two tryptic peptides, establishing their order.

And then you can get fragments and reconstruct them to make the overall sequence.

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Diagram of Overlap Peptides

And those are things that you can do. However, I think for the most part this type of stuff is done now in the bioinformatics realm. And you get the DNA sequence of the protein of interest. And then you can infer the rest of that without having to do an Edman degradation. I don't think people do many Edman degradations these days. However, if you get a protein that you don't know what it is, that might be one way that you can find out. And you can compare. So if you find a protein, and you don't know what it is, then you can look at its primary structure. And if it's similar to another protein's primary structure, then you can guess that it does the same thing or something similar. You can also look at primary structures of proteins from different animals. And if their primary structures are very similar, then that would suggest that the animals might be more closely related evolutionarily. Certainly if the primary structures are vastly different, then you probably would not be evolutionarily related as closely. However, most of that is done with DNA sequences these days. But it's kind of important to remember, you know - you get your DNA sequence and you don't actually know exactly what's going to happen to that DNA sequence. You can guess that it might be spliced in different ways and put together. But once a protein is created, then parts of it sometimes get cleaved off, new things get added to it, and you really can't predict that (yet) from the DNA sequence. So it is important to be able to look at proteins themselves, even though that's not done very often because it's actually much more difficult. And if you look for internal repeats in a protein, you can get some insight into that.

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Amino Acid Sequences Are Sources of Many Kinds of Insight (1/2)

1. Primary structures from different proteins can be compared to infer knowledge about structure and function.

2. Primary structure comparison of similar proteins from different species provides information about evolution.

3. Primary structure can be searched for internal repeats that may yield information on the history of the individual protein.

Figure 5.28 Repeating motifs in a protein chain. Calmodulin, a calcium sensor, contains four similar units (shown in red, yellow, blue, and orange) in a single polypeptide chain. Notice that each unit binds a calcium ion (shown in green). [Drawn from 1CLL.pdb.]

This is a calmodulin and it basically is the same sequence repeated four times. And you can see the red bit here and the yellow bit. They are very similar in their structure and also very similar in their function. They all bind to calcium. And the blue and the orange. So that you can determine a little bit about what the function might be. And also you can guess that maybe it started out as one segment and then somehow that segment was reproduced.

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Model of Repeating Motifs in a Protein Chain

And you can also use the amino acid sequences to find out things about the protein of interest. But usually that's just by comparing them to other proteins that you already know stuff about. So... ... That's about all we're going to say about protein structure and the biochemical techniques that we're going to use.

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Amino Acid Sequences Are Sources of Many Kinds of Insight (2/2)

4. Primary structure can reveal the presence of amino acid sequences that regulate protein function and location.

5. Primary structure can provide insight into the molecular basis of disease.

6. Primary structure can be used as a guide to explore nucleic acid information.