MBios 478: Mass Spectrometry Applications [Dr. Wyrick] Slide #1 Lecture 25: Mass Spectrometry Applications Measuring Protein Abundance o ICAT o DIGE Identifying Post-Translational Modifications Protein-protein Interactions o Co-Immunoprecipitation Welcome again to bioinformatics, in this lecture we ll be talking about proteomics applications that one can do with mass spectrometry, we ll talk about three different aspects of these applications. One is measuring protein abundance; we ll talk about two different methods that can be used to do this, ICAT, and DIGE. The second topic will be proteomics methods for identifying post-translational modifications to proteins, and then the third topic, which we will begin talking about in this lecture, and finish in the next lecture is protein-protein interactions, so how we can use mass spectrometry and other proteomics technologies to measure protein-protein interactions. Slide #2 Proteome Measurement Protein abundance 1) Isotope-Coded Affinity Tags ICAT 2) 2D-Difference Gel Electrophoresis - DIGE. So in the previous two lectures we talked about ways we can identify proteins using peptide mass fingerprint, and ms/ms, tandem ms peptide sequencing but that information typically doesn t give us quantitative information about how much of that particular protein or peptide is present in our sample. And so often we want to be able to measure quantitatively what the abundance is of each protein in our sample. And there are two methods that are commonly used to do this. One is called ICAT, which stands for Isotope-Coded Affinity Tags, and the second is DIGE, or 2D-Difference Gel Electrophoresis, and we ll talk about both of these methods that we could use to determine protein abundance levels in a sample. Slide #3 ICAT Technology Measures relative abundance of proteins between two samples o e.g., normal cell and cancer cell samples o or proteins isolated from cells grown at normal temp. vs heat shock Proteins are labeled with heavy ICAT tag; the other sample is labeled with the
light ICAT tag Heavy and light ICAT tags differ by 8 daltons o Peptides labeled with heavy or light ICAT tag can be distinguished by mass in a mass spectrometry experiment By comparing abundance of heavy vs light ICAT peptides, one can estimate the relative abundance of a protein in each sample So the first method we ll talk about is the ICAT technology, and this is a method for measuring relative abundance of proteins between two samples. An example would be say we isolated proteins from normal cells, and then from a parallel set of samples from cancer cells. So we have a protein sample that is derived from normal cells, and then a protein sample that is derived from cancer cells. And we want to identify which proteins show changes in abundance between those two samples. Alternatively, another example application would be look at protein samples from cells grown at normal temperature versus heat shock, so we can then identify proteins whose abundance changes in response to heat shock. So again these would be examples of potential applications that we could use the ICAT technology in which to measure protein abundance. The way the ICAT technology works is it uses what are called, ICAT tags, and in the next slide we ll talk a little bit more about how these tags work. First of all, these ICAT tags are labeled onto proteins at cysteine residues, so the free thiol group on a cysteine residue can react with the ICAT tag and make a covalent linkage between the tag and the protein, so because we have two different samples we use two different tags so we can distinguish these two samples from each other even after mixing, and so one sample, say the normal cell protein extract is labeled with the heavy ICAT tag, and then the other sample, say the cancer cell sample of proteins is labeled with the light ICAT tag. And the heavy and light ICAT tags differ by mass, so there are 8 Daltons difference in mass between the two tags. And thus we can distinguish peptides that are labeled with the heavy or light ICAT tag based on their mass in a mass spectrometry experiment, and that s the key step we ll talk about that a little bit more in the next slide. And so by comparing the abundance of heavy versus light ICAT peptides we can estimate the relative abundance of a protein in each sample. So if we see twice as much signal from the heavy ICAT tag for that particular peptide or protein versus the light ICAT tag, then we know the heavy ICAT tag sample, say the normal cell, there s twice as much of that particular protein compared to our light ICAT sample, say the cancer cell sample. Slide #4 Isotope-Coded Affinity Tags Different ICAT tags are used to label two different protein samples One sample is labeled with the heavy ICAT tag (X=deuterium) Other protein sample is labeled with the light ICAT tag (X=hydrogen) Heavy vs Light ICAT tags differ in mass by 8 daltons Biotin is used for affinity purification of ICAT-labeled peptides Image at bottom
And so, ICAT depends on this ICAT tag, the Isotope-Coded Affinity Tag, Isostope coded is, you use different isotopes of hydrogen. So the heavy tag, the heavy ICAT tag has deuterium in the place of hydrogen at eight positions in the tag, and that gives it a mass 8 daltons more than the light sample, in which those 8 positions are encoded with hydrogen, a different isotope, the lighter isotope. And so one sample is labeled with a heavy ICAT tag, well with a deuterium ICAT tag, and the other protein sample is labeled with the hydrogen ICAT tag, and so the heavy versus light ICAT tags differ in mass by 8 daltons, and so the schematic of the ICAT tag is shown below. You can see there is a thiol reactive group on the far right side, that s the part of the tag that reacts with the cysteine residue in a protein or peptide, and then there s a linker which acts as coding the tag for either being heavy or light each of those x positions, it will either be hydrogen or deuterium depending on if it s the heavy or the light ICAT tag, and then finally at the left end of the tag is the biotin tag, which is used for the affinity purification of the ICAT labeled peptides. Slide #5 ICAT Experimental Strategy 1. React ICAT tags with free thiol groups in proteins (cysteines) Heavy ICAT tag with Sample 1; Light ICAT tag with Sample 2 2. Mix samples, and digest with protease (e.g., trypsin) to generate peptides 3. Affinity purify ICAT-labeled peptides with Avidin Biotin moiety in ICAT tag interacts strongly with Avidin 4. Compare relative MS signals for heavy- and light-labeled peptides Heavy and Light peptide peaks will be separated by 8 daltons 5. Identify peptide and corresponding protein by MS/MS protein ID And so the experimental strategy is you react the ICAT tags with the free thiol groups in proteins on cysteine residues typically, and you use the heavy ICAT tag with one sample, and the light ICAT tag with the second sample. We then mix the samples together, so we combine both samples, now we can still tell which sample is which based on whether it has the heavy ICAT tag, or the light ICAT tag associated with it, so we don t lose information about which sample the protein was derived from. And then we digest the proteins with protease such as trypsin to generate peptides because it s easier, again, to work with peptides by mass spectrometry, particularly when we want to determine these small mass differences, these 8 dalton mass differences due to the tags. Step 3 is we affinity purify the ICAT labeled peptides with Avidin because the biotin moiety in the ICAT tag has a very strong interaction with Avidin, and so you can, with high efficiency, purify these ICAT labeled peptides, and the reason we want to affinity purify them is because the ICAT peptides will only be a fraction of the total peptides in the sample because only a fraction of the peptides will have cysteines and be labeled, and so we don t want all those other peptides to generate a lot of noise because we only can measure protein abundance with the ICAT labeled peptides. So we purify those away from the other unlabeled peptides for ICAT experiments. We then do mass spectrometry, and
compare the relative signals we get from mass spectrometry for the heavy and light labeled peptides, and the heavy and light labeled peptide peaks will be separated by eight daltons, that s how we can identify pairs of peptides, and then use the relative abundance of those peptides to determine the abundance of the protein in each sample. Then we need to identify what the protein or peptide is that s the peptide or protein that the peptide is derived from, and we can do that by doing things like MS/MS-based protein ID like we talked about in the previous lecture, so we can sequence the peptide, and that will tell us what protein those peptides are derived from. Slide #6 ICAT Experimental Strategy Diagram of ICAT This figure shows an illustration of this ICAT experimental strategy, we just discussed, shown on the left are the two protein samples, in this case sample 1 is labeled as mixture 1, sample 2 is labeled as mixture 2. We react them with the different ICAT tags, so sample, or mixture one is labeled with the light ICAT tag shown here with circles, and sample, or mixture 2 is labeled with the heavy ICAT tags shown here as cubes, and these again react with cysteine residues in proteins. We then mix the two samples of proteins, and add a protease to digest them such as trypsin. We then do an Avidin affinity enrichment to purify the peptides that have an ICAT tag associated with them. And then we do mass spectrometry, and typically there s two steps. The top right panel shows a mass spectrum, m/z spectrum of the ICAT peptides, and you see two peaks here, a light peak and a heavy peak, and typically again they will be separated by about 8 daltons, if you deconvolute the m/z spectra, and that s how you can find these pairs of peaks. And then the relative intensity, or abundance, the size of the peak of the light versus heavy tells you how much, what s the relative abundance of that particular protein in the two samples in mixture one versus mixture two. So that s how we get our relative abundance. Then we need to identify what the protein is, in the next step we take one of those peptides, and do an ms/ms spectrum so a tandem mass spectrometry experiment, and basically sequence that peptide, and shown below is where they have sequenced the heavy peptide, and identify its peptide sequence, and based on that then they can identify what protein that peptide was derived from, so we can then get quantitative information about the relative abundance of that particular protein. Slide #7 ICAT allows: Relative quantitation between 2 systems Protein ID from MS/MS data Can be done without gel electrophoresis So ICAT allows relative quantification between two systems, so if we are comparing
protein abundance between two sets of cells, or different experimental conditions. It allows those types of experiments, we get protein identification from the MS/MS data, and a big advantage of ICAT is you don t have to do a lot of experimental work like gel electrophoresis, which can be time consuming and technically challenging. It s almost entirely done following the reaction with the ICAT tags and the purification, and so on, it s all done in the mass spectrometer. Slide #8 ICAT Advantages Relatively fast LC/MS/MS Reduced peptide complexity Quantitation and ID in one experiment Disadvantages Not all proteins have cysteine PTMs not usually visualized Difficulties if protein is absent in one sample So the advantages is that it s relatively fast if you do an LC, liquid chromatography, MS/MS experiment it s relatively fast if you separate those peptides, and analyze them by tandem mass spectrometry, you get reduced peptide complexity because you are just purifying ones that have cysteine residues. That have been associated with this ICAT tag, and you get quantification, and protein identification in a single experiment. Disadvantages, are, first of all, not all proteins have cysteines, post-translational modifications are not usually able to be visualized as is the case with DIGE technique we ll talk about next, and you can have difficulties if the protein is present in one sample, and absent in another sample because in that case you won t get the pair of peaks, you ll just get one of the peaks, and so you won t be able, typically, to measure relative abundance of that particular protein. Slide #9 2D Difference Gel Electrophoresis (DIGE) Measures relative abundance of proteins between two samples o E.g., normal cell and cancer cell samples o E.g., proteins isolated from cells grown at normal temp. vs heat shock Proteins are labeled at amine groups (N-terminus, lysines) with fluorescent dyes (Cy3, Cy5) One sample is labeled with Cy3, the other sample labeled with Cy5 o Labeled protein samples are mixed prior to 2D gel analysis Proteins are separated by charge (pi) and molecular weight on 2D gel Relative abundance of samples is determined by comparing Cy3 vs Cy5 fluorescence intensity in 2D gel
Identify protein spots of interest by mass spectrometry So the second technique we mentioned for measuring protein abundance was 2D Difference Gel Electrophoresis, or DIGE, and this again measures relative abundance of proteins between two samples, just like with the ICAT experiment, so for example, if we want to compare normal cells to cancer cells, the proteins in those samples, or the proteins isolated from those cells grown at normal temperature versus heat shock, we can do all those types of experiments. The 2D Difference Gel Electrophoresis depends on labeling proteins with a fluorescent dye, and so the technique labels proteins at amine groups such as lysine residues with fluorescent dyes like Cy3 and Cy5. Again, note that these are very similar dyes that we discussed in the microarray section, so in this case instead of labeling DNA or RNA we are labeling proteins with these fluorescent dyes, but the idea is somewhat similar. So one sample will be labeled with say the Cy3 dye, the other sample will be labeled with the Cy5 dye, and then you mix those two protein samples prior to separating the proteins by 2D Gel analysis. So the proteins after labeling and mixing are separated by charge or isoelectric point, pi, and by molecular weight on a 2D gel. Then once we have separated the proteins on this 2D Gel we measure the relative abundance of each protein in the samples by comparing the signal we get from the Cy3 versus Cy5 fluorescence intensities in our 2D gel, and once we have identified proteins that are potentially changing in our experiment in their abundance, we can identify those protein spots of interest by mass spectrometry. Slide #10 DIGE Strategy Illustration of DIGE Strategy So this is a graphic illustrating the DIGE strategy, DIGE strategy. So again we have protein sample one, protein sample two isolated from different types of cells or whatever. You label one sample with the Cy5 dye, one sample with the Cy3 dye then we combine the samples. We then do a 2D gel where we separate proteins in 2 dimensions first we separate them based on charge, that s the pi or isoelectric point, the charge, the intrinisic charge of the protein, and so we use what is called isoelectric focusing to separate the proteins based upon their pi or isoelectric point, and then we do a second dimension, we separate based on molecular weight using a simple SDS PAGE type apparatus. So we ve separated proteins based both on their charge, and on their molecular weight, so where they follow on this 2D gel will depend on those two intrinsic properties of the proteins. Note that things like post-translational modifications will alter, potentially, things like the isoelectric point of a protein, so that you can actually get multiple spots for a single protein if a protein is phosphorylated, say for example. Once we have done the 2D gel analysis we then scan it for the different fluorescent dyes, very similar to how we scan microarrays with different dyes. So we do a Cy5 scan, a Cy3 scan, then we do an overlay of the two scans to look for protein spots that are changing in their abundance, or are of interest for some reason. We then identify specific spots we may be interested in and identify what the protein is by mass spectrometry, so we cut the spot out, do a trypsin
digestion to generate peptides, and then identify the protein by mass spectrometry. Slide #11 CyDye Labeling of Proteins Amine reactive (lysine residues and N-terminus) 3 Dyes available (Cy2, Cy3, and Cy5) ~3% labeling Constraints on labeling buffer So, the key step for this technique is the CyDye labeling of proteins, so we use amine reactive dyes that will label things like lysine residues and the N-terminus of proteins typically 3 dyes are available, Cy2, Cy3, and Cy5. Usually a small fraction of the protein is labeling, that s fine, and there are some constraints on labeling buffer, you have to switch buffers in your samples based on how you do the labeling. Slide #12 Analyze Multiple Gels for Statistics Image Typically you don t do a single DIGE experiment, but you do analyze multiple gels so you can generate statistics because there can be variability between individual experiments, and how much proteins separate and so on as far as the runs go. So you do multiple gels, and then you do statistics very similar to how we do like microarray analysis, your looking for proteins that are changing in their abundance, or their protein levels between the two samples. Slide #13 (No Title) Print Screen from 2D gel analysis So this is an example of a software analysis where we are looking at 2D gels, so then top left is two sets of scans, one s Cy5 on the left there, and we re looking at different types of scans, two different gels we have scanned, and we are trying to compare the protein samples look at abundance, and things like that. Slide #14
Spot-Picked 2D Gel Image image of 2D gel Once we have identified our proteins we would then pick spots using either manualing, or using a spot picking machine, that will pick those spots that we identify from our software as being significantly changed, and then once those spots are picked, and you can see those white holes correspond to spots that have been picked or identified in this experiment, we can then ID what those spots are by using mass spectrometry. Slide #15 Sypro Ruby Stained Image, 200 ug Total Protein image of 2D gel So in this case, the experimenter has gone through and identified a bunch of different spots, and in some cases you can see multiple spots corresponding to the same protein. For example, enolase, there s two different spots associated with that, vimentin, and so on. And the ones in red are the ones that are being upregulated in one sample versus the other or downregulated, now these multiple spots corresponding to the same protein could be for example proteins that are post-translationally modified, say they are phosphorylated or they are cleaved, or a variety of different things can cause multiple spots. So you can actually see for example, changes in post-translational modification so in some case you ll see one spot, perhaps corresponding to the phosphorylated form of the protein, going up in one sample, whereas the unphosphorylated form is going down in that same sample, and vice versa in the other sample. So you can see changes in modification levels through this technique. Slide #16 DIGE Advantages Relative quantitation via Fluorescence Replicate gels statistical info Pick spot based on confidence level ID with peptide mass fingerprint and MS/MS Visualize changes in some PTM s Disadvantages Gel based time, labor and money intensive Not all proteins run uniformly on gels So the advantages of the DIGE method is you get relative quantification of the protein
abundance levels via the fluorescence, since you can do replicate gels you can get statistical information and do some of the analysis methods we talked about with microarray analysis. You pick spots to identify the protein based on the confidence level. you re using your statistics to identify differentially expressed proteins. You can ID the protein with peptide mass fingerprinting, or by tandem MS. And you can also, this is a key advantage of DIGE as opposed to ICAT you can visualize changes in posttranslational modifications PTM s. Disadvantages, are that since it is gel based it is time, labor, and money intensive, and not all proteins run uniformly on gel, so you can get gel to gel variability for particular proteins. Slide #17 Other information we need to extract from proteome Post- Translational modifications Protein-Protein interactions Information not encoded in genome or transcriptome So in addition to looking at protein abundance, there s other information that s important to extract from the proteome. These would be post-translational modifications, and protein-protein interactions, this information is not encoded in the genome or the transcriptome, we have to have some sort of proteomics method for deriving this type of information. Slide #18 Common Post-Translational Modifications Phosphorylation: Mostly S, T, and Y residues Proteolysis: pro- to active species Methylation: K, R, possibly others Acetylation: K residues, N-terminus of protein Glycosylation Most modifications result in mass change Mass change can be detected by MS The first type we ll talk about is post-translational modifications so proteins are commonly post-translationally modified, they can be phosphorylated, namely on serine, threonine, or tyrosine residues. They can proteolyzed, so you can get specific cleavage of proteins that may activate or inactivate them. We have methylation of lysine, or arginine residues, this is particularly important for say, histone proteins, where histone methylation plays a role in gene regulation, also have acetylation of proteins, such as histone proteins, and other proteins involved in gene regulation. Again, acetylation typically occurs on lysine residues, and on the N-terminus of the proteins. You have glycosylation, and many other modifications. Most of these modifications result in a mass change, and mass changes can be detected by mass spectrometry, so, mass spectrometry
is a good method for identifying post-translationally modified proteins, and where those modifications occur. Slide #19 Mass changes associated with post-translational modifications Chart of post-translational modifications So, what are the mass changes associated with post-translational modifications? So Acetylation, when you add an acetyl group to a lysine residue, that gives you a mass change of 42.0373 daltons. Methylation of lysine or Arginine residues, it depends on the methylation state, so if you just mono-methylate a single methyl group is added the mass change is 14.0269, di-methylation is twice that, tri-methylation is three times that, adding three methyl groups. And then, another common modification is phosphorylation, which mainly occurs on serine, threonine, or tyrosine residues, and that mass change is about 80 daltons. Note that the mass change for acetylation and tri-methylation are very similar, and so you need to have good mass accuracy to determine which of these modifications is occurring on, say, a lysine residue because the mass difference is very small, you know.05 or so daltons, and so mass accuracy is very important for distinguishing between acetylation and tri-methylation. Other modifications tend to have more unique mass changes, but that s an important one to keep in mind, acetylation versus tri-methylation. Slide #20 Phosphorylation is one of the most well studied reversible covalent modifications Signal transduction Molecular activation/deactivation Dynamic modifications-kinases and phosphatases We ll mainly talk in this section about phosphorylation because it s one of the most well studied reversible covalent modifications. It s involved in signal transduction, map kinase, and other kinase cascades, involved in cell signaling. It s also proteins can be activated or deactivated by phosphorylation. Many Kinases for example are activated by phosphorylation, and it s a dynamic modification. Kinases add phosphate groups, phosphatases remove them from proteins. Slide #21 Proteomics methods to detect Phosphorylation Charge phosphorylated peptides preferentially interact with positively charged ions Immobilized-metal affinity chromatography (IMAC) Mass Measure mass change by +/- phosphatase treatment
So, there are at least two proteomics methods that can be used to detect phosphorylation. And they exploit different aspects of the modification, so, different chemical aspects. So phosphorylation obviously adds charge to the peptides, so it adds a negative charge, so phosphorylated peptides preferentially interact with positively charged ions, and that allows one to purify phosphorylated peptides using what is known as immobilized-metal affinity chromatography, which exploits that charge difference in phosphorylated peptides. The second method is by mass where you simply measure the mass change in a peptide due to phosphorylation. And one additional step one can do is take that peptide and treat it with a phosphatase, if a phosphatase removes that phosphate group, then we ll see a mass change corresponding to the loss of that phosphate group. So we can confirm that a peak we suspect may be due to a phosphorylated peptide is in fact corresponding to a phosphorylated peptide by adding a phosphatase to the sample, which should cause then a shift of the peak decreasing its mass by about 80 daltons. Slide #22 Immobilized Metal Affinity Chromatography (IMAC) Flow Chart of IMAC So this graphic illustrates how the Immobilized Metal Affinity Chromatography, or IMAC, method works for identifying phosphorylated proteins and the sites of phosphorylation. So if we have our phosphorylated protein, or mix of proteins, we start out by doing a trypsin digestion, which will cleave the proteins into peptides. And what we are looking to isolate are peptides that have a phosphorylation site associated with them, so isolating those proteins or peptides, and so to do that, which in this case, in the graphic, is shown with a big P associated with them. We use an IMAC column, which will then affinity purify peptides that have a phosphorylation site, and so essentially how that IMAC column works is that it has beads inside the column that have metal ions associated with them, in this case the IMAC matrix, and these will be let s say for example, iron or gallium metal ion coated beads. And the phosphorylated peptides, which have a strong negative charge due to the phosphate modification will strongly interact with these positively charged metal ions, and so they will stick to the column whereas the unphosphorylated peptides will flow through, and so we elute the phosphorylated peptides from the column we can then identify what these peptides are by mass spectrometry using the methods we talked about in the previous lectures. Slide #23 Summary of post-translational modifications Most modifications result in mass change Mass changes can be identified by MS Some mass changes are difficult to distinguish (e.g., tri-methylation vs. acetylation) Not all modifications have unique character to exploit
Most current work phosphorylation Significance Most commonly known Unique properties And so to summarize most post-translational modifications result in a mass change, and these mass changes can be identified using mass spectrometry, some mass changes are difficult to distinguish for example, tri-methylation versus acetylation have very similar masses. We can also in some cases add, for example a phosphatase to confirm that the peak we think is modified is due to that modification, so if we add a phosphatase to a phosphorylated peptide peak that peak should shift in mass upon phosphatase treatment, the shift should equal the mass change due to the loss of a phosphate group, about 80 daltons. Not all modifications have unique character to exploit. Phosphorylation is the most commonly studied modification because you can take advantage of the highly negative charge on the phosphate group to do things like IMAC and so on. Slide #24 Protein-Protein Interactions Protein-protein interactions contain important biological information Protein-protein interactions can be critical to function o Multi-subunit protein complexes (e.g., ribosome) o Protein signaling cascades Many purification methods disrupt protein-protein interactions Systems-level methods are needed to detect protein-protein interactions So last topic we ll talk about in our discussion of mass spectrometry applications are protein-protein interactions. Protein-protein interactions are important because they contain important biological information. Many protein-protein interactions are critical to cellular function, or protein function for example, many proteins function in multisubunit protein complexes such as the ribosome, or RNA polymerase II, so knowing which proteins that protein interacts with is important to understanding the function of the complex, and the components of the complex. And in addition you can have more transient interactions between proteins that form, for example, protein signaling cascades that are involved in cell signaling, and signaling networks in cells. However, many purification methods used to isolate proteins disrupt protein-protein interactions, so it s difficult to extract that type of information. Thus systems level type methods are needed to detect protein-protein interactions, particularly if we want to identify protein-protein interactions for many proteins at once. Slide #25 Protein-Protein Interactions No chemical, charge, or mass difference that is a common marker of protein-
protein interactions We need some bioanalytical tricks Common Methods: Co-Immunoprecipitation (Co-IP) o Tandem affinity purification (TAP) tags Yeast two hybrid Protein and Antibody arrays So, there s no chemical, charge, or mass difference due to a protein-protein interaction, as we had with post-translational modifications, so we have to have some additional tricks to isolate these interacting proteins. We ll talk about one method today, which is Co- Immunoprecipitation, and then we ll discuss two other methods in the next lecture, so the method we ll be talking about today is co-immunoprecipitation, which can be facilitated using what are called tandem affinity purification tags, or TAP tags. Slide #26 Co-Immunoprecipitation (Co-IP) Methods Flow chart So this is how co-immunoprecipitation works, we have a cell extract, we re we ve isolated proteins from a cell, and we ve done a gentle protein isolation, so protein interactions remain in the extract, and so in this case we have four different proteins shown by the different shapes. And we have our target protein, which is a green rectangle, and then the protein that interacts with that target protein, our interacting protein, which is shown with a red elipse, or oval. So the co-immunoprecipitation method relies on using an antibody that specifically interacts with our target protein, and so we add the antibody, which is typically associated with some kind of bead or resin. It will bind specifically to your target protein, and isolate it, fractionate it, and in addition, any interacting proteins with our target protein will also be isolated, and so we ll isolate those from the non-interacting, or light blue proteins, and then we ll analyze the immunoprecipitated sample on an SDS PAGE gel and so we can separate the proteins in this complex, and so in this case we see two bands on our SDS PAGE gel, a target protein, the one we immunoprecipitated, which we expect to see, and then in this case one interacting protein, which is potentially associated with our target protein. Then simply what we want to do is identify the interacting protein, so we can cut that band out of the gel do a trypsin digestion, and then do mass spectrometry to identify coimmunoprecipitated proteins. And so by this method we can identify which proteins interact with our target protein of interest. Slide #27 Difficulties for Co-IP
Non-specific binding on resin can be a problem Harsh elution conditions elute specific and nonspecific interacting proteins Dynamic range limited nonspecific interacting proteins are much more abundant Often a limitation can be the co-immunoprecipitation step, so we may not have antibodies to our protein, or we may get non-specific binding of proteins to our resin. And when we try to elute our protein from the resin, from the antibody we can elute both specific and nonspecific interacting proteins, we don t want to be getting these non-specific interacting proteins that are not associating with our protein of interest, but just with the antibody or the beads. And so it s helpful to have a different strategy for separating these interacting proteins. And again, finally the dynamic range can be limited with this method if you have very abundant proteins that are non-specific interacting like ribosomal proteins, which are really abundant in protein extracts, you can see these by mass spectrometry even though they are not really specifically interacting with our target protein. Slide #28 Tandem Affinity Purification (TAP) Image at left Label protein with cleavable region between two affinity tags Affinity pull down complexes (Protein A IgG beads) Elute with TEV protease 2 nd Affinity capture (CBP-Calmodulin beads) Only proteins bound to TAP-tagged proteins are purified nonspecific signal reduced So a way to get around these problems is to use a Tandem Affinity Purification strategy, or TAP tag strategy. In this strategy we label our protein of interest with a TAP tag. On the picture on the left this shows, on the top left is a protein shown in blue, a blue oval with a green TAP tag associated with it, and there are three parts to this TAP tag. There are two affinity tags, the calmodulin binding peptide, or CBP region. In the middle, on the far right of the tag is protein A, and then between those two parts, the calmodulin binding peptide, and the protein A subunit is a TEV protease cleavage site. And so there is a cleavable region between two affinity tags, and so we take our protein of interest with the TAP tag in the cell extract, and we want to see which proteins interact with our protein of interest so in this case you can see multiple proteins are interacting with our blue oval binding protein of interests, and so first we would do an affinity pull down a first affinity purification step where the protein A part of the TAP tag binds specifically to IgG beads, so that s our first affinity column, and that separates the specific binding proteins with the contaminant proteins. We can then specifically elute our TAP tag protein by adding the TEV protease, which will cleave at that linker region between the protein A and calmodulin binding peptides, so we elute it with the TEV protease, that
will cut off the Protein A part, so the TAP tag protein, and the associated proteins will come off the column, and then we ll do a second affinity column where we bind the calmodulin binding peptide part of the protein to calmodulin beads, and that will again capture the protein with any associated, proteins, interacting proteins, and then we elute them by removing calcium with EGTA, which disrupts the calmodulin-calmodulin binding peptide interaction, and that gives us the TAP tag protein with other proteins that are interacting, and that reduces this problem of non-specific signals, and so that s the strategy of identify interacting proteins using co-immunoprecipitation, or TAP tag purification. In the next lecture we ll talk about a couple of different methods, protein arrays, antibody array method, and the yeast two hybrid method for identifying interacting proteins.