The function of most proteins is regulated in some way by enzymatic post-translational modifications (PTMs). A large number of covalent PTMs exist in nature and can affect many aspects of protein function including enzymatic activity, binding partners, stability, structure, and cellular localization. Most PTMs are reversible, and therefore allow dynamic control of protein function. Reversible phosphorylation of serine, threonine, and tyrosine, controlled by the opposing activities of kinases and phosphatases, is the most abundant and heavily studied regulatory PTM in eukaryotic cells. At least 75% of human proteins are phosphorylated in vivo
] and many proteins are phosphorylated on multiple sites that can be subject to differential regulation [2
], often by more than one kinase and/or phosphatase. As an example, more than 100 in vivo
phosphorylation sites have been mapped on the human anaphase-promoting complex (APC), a large E3 ubiquitin ligase complex that regulates progression through mitosis [1
]. Evidence suggests that the APC is regulated, both positively and negatively, by at least four different kinases [5
]. This complexity makes functional studies of protein phosphorylation challenging. There is a need for improved methods for detection and quantification of individual PTM sites on multiply modified proteins.
Very few analytical methods employed to study phosphorylation, or PTMs in general, are effective at quantitatively detecting multiple sites independently on the same protein. Mass spectrometry (MS) is one exception. Reduction of proteins to peptides is an effective way to separate the PTMs on a single protein into distinct molecules for mass spectral analysis. Tandem MS can provide the sequence of individual peptides and the exact location of any PTMs. Although the complexity of biological systems poses a constant challenge to the detection and quantification of target molecules, the development of affinity enrichment strategies, multi-dimensional chromatographic separations, improved instrument designs, and various other methodological advances are steadily allowing more comprehensive study of the proteome. Selected reaction monitoring (SRM) MS, which relies on the filtering capabilities of a triple quadrupole mass spectrometer, has become a popular method for direct detection of specific target proteins within complex biological samples. Because of its sensitivity and specificity, SRM is also a useful approach for the quantitative study of PTM regulation in living systems.
To fully understand regulation and function of PTMs it is necessary to measure their dynamic changes under different conditions and in response to physiological and environmental signals. A variety of methods exist for the absolute and relative quantification of proteins and PTMs from peptide-based mass spectral data [13
]. Most quantitative MS approaches employ some form of stable isotope dilution, which has the advantage of allowing simultaneous analysis and comparison of multiple samples or a single sample with an internal standard [15
]. However, use of stable isotopes is not essential for quantification and a number of “label-free” approaches have also been developed [14
]. Label-free quantification requires comparison of separate MS experiments, which necessitates exquisite care in sample preparation and the appropriate controls or standards to account for run to run signal variations. Although stable isotope-based methods are generally thought to be more accurate than label-free approaches [13
], there are important advantages to label-free methods such as simplicity and general applicability. With label-free quantification there are no specialized reagents required, no extra chemical reaction and sample processing steps, and the methods are suitable for any experimental system. For these reasons, label-free quantitative MS methods have important advantages for the study of PTMs and are becoming more common.
A variety of strategies have been developed for quantitative MS-based analysis of phosphorylation and other PTMs (for example [18
]). The majority of these methods rely on stable isotope labeling or addition of specific standards for quantification. Some are limited to studies of synthetic peptide substrates or highly purified protein preparations or require phosphopeptide enrichment steps. We set out to establish a strategy that will be generally useful for the quantitative measurement of changes in phosphorylation on intact proteins. Our approach uses label-free quantification of phosphorylation stoichiometry [21
] for simplicity, compatibility with all experimental systems, and practicality for studies of large numbers of phosphorylation sites. We used SRM MS of peptides generated by enzymatic digest because the sensitivity, specificity, and dynamic range of SRM [27
] make it suitable for analysis of highly complex biological samples, and it is also capable of simultaneous monitoring of up to hundreds of target molecules in a single sample run. Related SRM MS methods were developed and used recently. In one case, changes in phosphorylation of the Pho4 transcription factor in budding yeast were measured [28
], but without a rigorous quantification strategy. In another case, autophosphorylation sites on the Lyn tyrosine kinase in mouse xenograft tumors were monitored [26
]. Our strategy is equally useful for studying phosphatases and kinases, and can be applied in vitro
as an enzymatic assay, or in vivo
to quantify changes in many phosphorylation sites in response to experimental variables. Among other potential applications, we predict it will be very useful for the exploration of kinase and phosphatase specificity. The general method is also suitable for studying other PTMs.
In this report we describe proof of principle experiments that validate this strategy. We demonstrate that SRM MS combined with label-free quantification is comparable to conventional assays for measuring reaction rates and steady-state kinetic parameters of phosphatases and kinases. We also demonstrate the utility of the method for simultaneously monitoring multiple phosphorylation sites on intact protein substrates, including in a whole cell extract, and the potential to reveal different responses of individual sites to experimental variables. Possible experimental designs and practical aspects of applying this method to large-scale studies in complex biological mixtures and in vivo are discussed.