Proteomics has become a vital tool for biological scientists performing research on the healthy and diseased states of living things. It involves the large scale and systematic analysis of all proteins within a given cell, tissue, or organism. Because proteins are regulated by many different internal and external stimuli, the proteome is dynamic and quantities of proteins can change from one state to the next. Therefore, in order to be of the highest utility, proteomics experiments need to both identify and quantify proteins so that comparative studies can be done, such as between healthy cells and tumor cells, or the comparison of different treatment regimens.

Proteomics has evolved significantly over the last 20 years. Proteomics was first performed using 2D gel electrophoresis and amino acid sequencing¹. In this approach, proteins were identified by comparing their amino acid composition, estimated isoelectric point, and molecular weights to a database of proteins generated from a genomic database. It wasn’t long however, before scientists realized that mass spectrometry (MS) and tandem mass spectrometry (MS/MS) could provide a more streamlined approach for global protein identification. Proteomic samples were enzymatically digested and the entire peptide mixture was subjected to analysis. Similar to the 2D gel technique, proteins were identified by comparing peptide MS and/or MS/MS data to information within protein databases that had been created by translating the databases of partially or fully sequenced genomes. The technique rapidly gained acceptance and became a vital tool within the mass spectrometry community.

In the early days, simply detecting and identifying proteins was enough and laboratories would generate lists of proteins identified from a particular organism or sample. The concurrent rush in the genomics world to sequence the genomes of humans and other organisms served as the foundation upon which the protein identification portion of MS based proteomics experiments could further evolve. Additionally, continuous improvements in software, workflows, and hardware, have now resulted in an ever-increasing number of proteins that can be identified by mass spectrometry. 

While protein identification was fairly straightforward, using mass spectrometry for protein quantitation proved to be more difficult initially. Because different peptides can have different ionization efficiencies and detectability depending upon experimental variations, comparison of peak heights and areas between different samples was challenging. Accuracy and reproducibility can suffer if instrument and sample preparation variations are not strictly controlled. Even with state-of-the art instruments of the time, direct untargeted quantitation of complex proteome samples was unreliable and often not sensitive enough. As a result, scientists started to investigate the use of stable isotope labeling for quantitation which had been used previously for other applications. Up until this time, scientists had been using more tedious and laborious techniques for protein quantitation such as antibody-based methods and density comparison of 2D gel spots.

With stable isotope labeling, one sample is derivatized with a “light” version of a chemical tag while another sample is labeled with a version of the tag that incorporates a “heavy” isotope. The samples are then mixed together and analyzed in the same experiment. Identical compounds from the different samples co-elute as pairs of peaks and can be distinguished by the mass difference between the heavy and light isotope labels. Quantitation is performed on the pairs of peaks in the MS data and identification is performed using the MS/MS fragment data. This technique eliminates much of the bias that can be introduced when comparing peaks between different experiments since the data from all samples are collected within the same experiment.

Isotope coded affinity tags, or ICAT® Reagents^2,3, were the first stable isotope reagents synthesized specifically for quantitative proteomics. These reagents were developed in 1999 and made commercially available in 2001. ICAT Reagents are available in a light and heavy version and specifically bind to cysteine residues on proteins. The reagents also have a biotin moiety attached to the tag that is used to specifically capture the labeled peptides out of a complex mixture. In an ICAT experiment, samples are labeled with either the heavy or light version of the reagent. The two samples are then mixed together and the proteins are digested into peptides. All cysteine-containing peptides are then isolated and enriched using an avidin column that binds to the biotin portion of the ICAT Reagent. Only these cysteine-containing peptides are analyzed by mass spectrometry. The benefit of this technique is that by focusing only on cysteine-containing peptides, the overall data complexity is greatly reduced. Although any protein that does not have a cysteine is not detectable, most proteins will have at least one cysteine containing peptide, if not more. Thus, with ICAT, researchers could seamlessly quantitate large numbers of proteins by simply adding a few extra steps to their proteomics sample preparation protocol.

Stable Isotope Labeling by Amino Acids in Cell Culture, or SILAC, is a technique that was first described in 2002⁴. In this approach, two populations of a cell line are cultured under identical conditions with the exception that one is fed a growth medium that contains a natural or “light” form of a particular amino acid and the other contains a “heavy” form of the amino acid (e.g., incorporation of ¹³C into lysine). Cells within each population incorporate the light or heavy version of the amino acid into the proteins they synthesize. Afterwards, the light and heavy populations are mixed and analyzed by MS where the two samples can be distinguished by the mass difference of the labeled amino acid. While SILAC is typically used for cells growing in cell culture, it has also been applied to higher organisms such as flies and mice by feeding them heavy labeled food, but the process can be time consuming.

ICAT and SILAC are technologies that provide a mass difference in the molecular weights of differentially labeled peptides. Proteomics samples are inherently already complex, and mixing samples together further increases the complexity in the MS space. Thus, in practice these experiments were limited to comparing only 2 or 3 sample types. In order to compare a greater number of samples simultaneously, a different approach was required. Isobaric tags provide a means to increase the number of samples that can be simultaneously analyzed without increasing the sample complexity. Different versions of isobaric tags are identical in mass but differ in their chemical structure with respect to the sites for incorporation of heavy isotopes. Thus, differences are observed in the fragmentation spectra of any peptides derivatized with these compounds, whereas the MS spectra are all identical.

Tandem Mass Tags or TMT Reagents are a type of isobaric multiplexing tag and were first reported in 2003.⁵ Around this same time, iTRAQ® Reagents, were also developed and rapidly commercialized.⁶ The commercialization of TMT Reagents followed several years later. In the isobaric multiplexing tagging strategy, all versions of each tag have the same molecular mass but the positions of heavy and light isotopes are adjusted in order to affect the mass of a “reporter ion” region and “balance mass” region within the compound. With iTRAQ Reagents, heavy isotopes of carbon, nitrogen, and oxygen are incorporated into the structure in order to shift the reporter and balance masses. Since all versions of the reagent tag are identical in molecular weight, the same peptides originating from different samples will have the same mass in MS space regardless of which reporter ion is attached. Upon fragmentation, the reporter ion can be clearly distinguished, and the identity of the peptide determined from the sequence of the larger MS/MS peptide fragments, and the quantity of that peptide from each biological sample determined from the areas of each respective reporter ion peak.

iTRAQ Reagents are capable of analyzing up to 8 samples simultaneously.⁷ They have been extremely successful for quantitative proteomics and it is easy to see why. The reporter ions for iTRAQ Reagents span a region of MS/MS space that is empty in the fragmentation spectra of peptides, providing a very ‘clean’ area for quantitative accuracy. The reagents fragment well and provide good signal for both identification and quantitation – getting this balance right is a key factor in the design of the reagent chemistry. The experimental procedure for using iTRAQ Reagents is straightforward and involves only one additional step in the sample preparation protocol. Additionally, the reagents label all peptides and work with all matrices.

These attributes, and more, have contributed to the widespread adoption of iTRAQ Reagents by the scientific community iTRAQ reagents have been used to pioneer discoveries in disease research, in systems biology research, and across many different biological species, providing countless insights into the field of life science. Without the ability to multiplex to such a high degree, certain applications would be much more tedious, such as proteomics time course studies and analysis of many replicates. The importance of this technology was recognized at HUPO 2014 where Science and Technology Award was given to the teams involved in the commercialization of isobaric labeling for protein quantification. In summary, iTRAQ Reagents have helped to transform the utility of quantitative mass spectrometry for the biological sciences and are probably the most extensively used labeling reagent for quantitative proteomics with well over 1000 papers published on their use.

References:

1. Marc R. Wilkins, et. al., (1996). “From Proteins to Proteomes: Large Scale Protein Identification by Two-Dimensional Electrophoresis and Amino Acid Analysis”. Nature Biotechnology. 14 (1): 61–65.
2. http://www6.appliedbiosystems.com/press_releases/2001pittcon/
3. S. P. Gygi, et. al., (1999). “Quantitative Analysis of Complex Protein Mixtures Using Isotope-Coded Affinity Tags”. Nature Biotechnology, 17(10), 994-999.
4. S. E. Ong, et. al., (2002). “Stable Isotope Labeling by Amino Acids in Cell Culture, Silac, as a Simple and Accurate Approach to Expression Proteomics”. Molecular & Cellular Proteomics, 1(5), 376-386.
5. A. Thompson, et. al., (2003). “Tandem mass tags: a novel quantification strategy for comparative analysis of complex protein mixtures by MS/MS”. Analytical Chemistry, 75, 1895–1904. 
6. P. L. Ross, et. al., (2004). “Multiplexed Protein Quantitation in Saccharomyces Cerevisiae Using Amine-Reactive Isobaric Tagging Reagents”. Molecular & Cellular Proteomics, 3(12), 1154-1169.
7. L. Choe, et. al., (2007). “8-Plex quantitation of changes in cerebrospinal fluid protein expression in subjects undergoing intravenous immunoglobulin treatment for Alzheimer’s disease”. Proteomics, 7(20), 3651-3660.