Analyzing proteins is about trade-offs. Do you want to take a top-down approach or a bottom-up? Will you get the information you need from looking at the proteins or by looking at the peptides? If you look at peptides, you lose isoform information, but if you look at the proteins you may have a lower throughput. It's a balancing act.
The two well-established, well-known methods in protein analysis, particularly for the separation step of the process, are two-dimensional gel electrophoresis and liquid chromatography, coupled to mass spectrometry. While those approaches are still in use, they have both been updated and have given birth to newer methods that allow for an even better peek into the proteome. Some researchers are loyal to one or the other approach, though it often depends on their end goal. "We have to fit the best method to whatever problem we have," says the University of Reading's Rainer Cramer. "We use both, basically."
"When you are thinking about what strategy to use, you really have to think about the question you are trying to answer," the Scripps Research Institute's John Yates adds.
If you're interested in a first look at the proteins or protein isoforms in a given proteome, a gel-based approach may be the best bet, though if you want to get a global view of the peptide, a shotgun, LC-based method could be a better choice. Also on the horizon are newer techniques: one combines the gel and gel-free approaches, while another is a method co-opted from the small molecule world.
For their part, mass specs have become increasingly more sensitive. Yates says that since he started work in this area in the 1980s, mass specs have become "impressively sensitive."
"Mass spectrometry is one of the most sensitive techniques," Cramer says. "The problem is more how am I going to get my sample into the mass spectrometer in a form that is actually very nicely amenable for mass spec analysis."
The University of Missouri's Jay Thelen agrees. "It's all about getting sample complexity reduced to a level that's compatible with online nano LC/MS."
Running proteins out on gels to get a look at their components has been on the scene for quite some time. Two-dimensional gel electrophoresis has been in use since the late 1950s. It's a routine analysis, says ¬Greifs¬wald University's Jörg Bernhardt. He estimates that his group runs about 100 gels a week and that they only face minor problems, most of which are easy to handle.
Though well established, 2D gels suffer from reproducibility issues and an inability to make inter-gel comparisons easily. Samples run on different gels are subject to slightly different running conditions during separation, making it difficult to compare spots on different gels.
To overcome that drawback, Carnegie Mellon University's Jonathan Minden developed difference gel electrophoresis, or DIGE, that uses dyes to distinguish proteins run on the same gel. "The basic premise was if you want to find what proteins are different between two samples, it's good to run them together on the gel rather than two separate gels," Minden says.
Along with Alan Waggoner, also at Carnegie Mellon, Minden designed the dyes they'd need. The first set of dyes attached to lysine residues but labeled less than 3 percent of proteins; the unlabeled proteins would run slightly faster on the gels, Minden says. Then, they tried cysteine-reactive dyes, and with those they were able to achieve 100 percent saturation.
Naturally, Minden uses DIGE in his lab. "We use that for initially identifying proteins that are changing under whatever biological circumstance we're testing. Once we see those differences, we cut the protein spot out of the 2D gel, and use MALDI-TOF to identify the protein within that spot," he says.
DIGE and 2D gels also lend themselves to examining the proteome at the protein level. "The advantage of it is that you can resolve post-translationally modified proteins very nicely on 2D gels. That's pretty tricky to do with other approaches," says Missouri's Thelen.
"One of the great things about 2D gels is being able to look at isoforms of proteins, so whether that's proteolytically processed or whether it has modifications of some kind, you can really potentially see those patterns on the 2D gel. It's not always straightforward to pull the spot and figure out what's causing those changes, but you at least see them and you can work from there," Scripps' Yates adds. "I think that one of the big advances that's come along in that field has been the DIGE technique. That's really, I think, helped a lot."
Though DIGE compensates for the reproducibility problem of 2D gels, it still shares other disadvantages with 2D gels. "Some of the problems are still inherent just to two-dimensional gel electrophoresis that the labeling method won't really help. That's being able to focus membrane proteins," Minden says. "The other issue is very high molecular weight proteins also don't focus very well."
In that case, Bernhardt says that a slightly different approach may help. "With 2D gel it's very difficult to separate very small proteins. You need different running buffers and a completely different buffer system for that. The same problem we have faced very hydrophobic proteins coming from membranes," Bernhardt says.
That particular problem may extend beyond gels. Minden says that even straight mass spectrometry methods aren't terribly good at getting membrane proteins. "They are just very sticky, hard-to-deal-with proteins," he says.
Another disadvantage to 2D gels is the range they cover. "2D gels are a good place to start when you are quantifying and characterizing the proteome for the first time," Thelen says. Once the study has moved beyond the initial steps and needs to dig deeper into the proteome, he says it becomes much more challenging.
To get a look at the peptide level of the proteome, researchers often turn to chromatography. In the early 1990s, Scripps' Yates worked on a method to study protein complexes using single-dimension liquid chromatography. At the same time, his group also developed a software program to help researchers to identify peptides from a digested mixture coming off a tandem mass spec.
"I'd always been enamored with multi-dimensional liquid chromatography," Yates says. They then began to look at more and more complex mixtures, multi-dimensional LC, and followed their "natural inclination to push to the limit." From that work came the multidimensional protein identification technology approach, or MudPIT, that researchers now use to resolve peptides from complex mixtures of proteins.
MudPIT lends itself to shotgun, bottom-up proteomics. "You take your really complex proteome sample, you just digest the whole lot of it and basically you put it in MudPIT … and you separate, in a multidimensional fashion, the peptides," Reading's Cramer says. "It has a major advantage because, from the technical point of view, it seems to be one of the most sensitive methods. If you really want to go out there and do a global proteomics exercise, look at as many proteins or protein identifications as possible, then that is a very powerful method."
"It wasn't exactly clear how much better it would be at the time versus two-dimensional gel electrophoresis, but it was clear that what it really brought to the table for that type of analysis was this ability to be really highly automated," Yates says, adding that 2D gels can be labor-intensive.
In his lab, Yates has studied a variety of proteomes using this approach. "I think some of the most productive experiments that we've done have looked at constrained structures within the cell, so protein complexes, organelles, things like that, where you can ask much more specific questions," he says. "Protein expression analysis — people call it discovery but it's sort of an experiment without a question in a sense because you don't really know what you are looking for and you hope that something will pop out."
The disadvantage of MudPIT, however, is that since the proteins are broken down to their component peptides, any information about modifications and isoforms is lost. Yates says this is common to these types of analyses. "You lose that information for the most part in these bottom-up shotgun proteomics strategies," he says. "If you just want to catalogue them, bottom-up is certainly good for doing that. If you want to look at what the relationship among the various modifications are on a histone, you would probably do a top-down or middle-down approach."
A different method that Jay Thelen's lab at Missouri is evaluating combines the gel and gel-free approaches. This new approach, called GeLC/MS, couples SDS-PAGE to liquid chromatography and mass spectrometry. Here, the SDS-PAGE works as a pre-fractionation step. "What it allows us to do is dig deeper into the proteome in a way that we can't really accomplish using 2D gel or MudPIT approaches," he says.
This way, he says, they can load a lot of protein — half a milligram — onto the SDS-PAGE. That then goes through the LC and mass spec before things get tricky. "Once you've identified the peptides in the mass spectrometer, then quantitation is certainly achievable," Thelen says. His group has been comparing spectral counting results to peak integration results, and he says that spectral counting looks promising. "For now, spectral counting is easy and it seems to be semi-quantitative," Thelen says.
The disadvantage to GeLC/MS is speed. With SDS-PAGE as the first step, it all but eliminates the possibility of being a high-throughput method. "When you do that and then section it into smaller pieces, the throughput goes down pretty quickly there. It's a low-throughput technique but it allows you to dig deeper," Thelen says.
Besides just kicking the tires on the GeLC/MS approach, Thelen's group has also been putting it through its paces and comparing it, as best can be done, to other techniques, particular to 2D gels. They've compared molecular weight versus pI and versus hydrophobicity. "The indications are that SDS-PAGE, as well as gel-free MudPIT, yield higher frequency of membrane proteins, high and low molecular weight proteins, high/low pI proteins," he says. "So, basically, 2D gels are useful but they have their well-known biases." This combination GeLC/MS approach appears to counteract some of those biases.
While it might not be a new technique, a method called multiple reaction monitoring, or MRM for short, is a fairly new to the field of proteomics. "It's a very promising, though not new, technique," says Cramer from Reading. MRM has been used for years in small molecules, he adds. It's based on mass spec and it can be used to absolutely quantify biomolecules or biomarkers.
To apply the MRM approach, researchers need to know what they are looking for. Thus, if you want to do a global analysis or survey, Cramer says, MRM wouldn't be your method of choice. However, if you have one or a few peptides of interest, you can focus on those with MRM. Basically, he says, researchers watch for those specific peptides and specific transitions as they pass through the mass spec. That way you can get quantitative values as well. He adds that scientists "can monitor how [the peptide(s) of interest] might change from one stage to another if you want to do clinical proteomics, for instance."
"It's a way of tracking where a peptide should be during the LC/MS/MS process," adds Minden. "By doing the typical mass spec analysis, which takes intact proteins and cleaves them with trypsin, you increase the complexity 50-fold. To then start sifting through that for important changes is very, very difficult. Knowing what you are looking for from the beginning often helps."
In that case, MRM is best as a secondary analysis, when a few interesting proteins or peptides have been identified and need to be investigated further. That way, the researcher already has the fragment ion mass information needed. "You basically fix on these specific masses on the MRM. With new samples, you find out and monitor with the MRM method your peptide or peptidic biomarker, how it is actually expressed in these various different samples," Cramer says. He adds that MRM looks promising as a method for use in clinical proteomics.
Though there are many methods to choose from when analyzing proteins and each of them may give slightly different results, comparative studies have shown that all those results are valid. Whatever approach or combination of approaches you choose, keep in mind the goal of your study and what it is that you are trading off to get those results.