A Review on Quantitative Multiplexed Proteomics

Abstract

Over the last few decades, mass spectrometry-based proteomics has become an increasingly powerful tool that is now able to routinely detect and quantify thousands of proteins. A major advance for global protein quantification was the introduction of isobaric tags, which, in a single experiment, enabled the global quantification of proteins across multiple samples. Herein, these methods are referred to as multiplexed proteomics. The principles, advantages, and drawbacks of various multiplexed proteomics techniques are discussed and compared with alternative approaches. We also discuss how the emerging combination of multiplexing with targeted proteomics might enable the reliable and high-quality quantification of very low abundance proteins across multiple conditions. Lastly, we suggest that fusing multiplexed proteomics with data-independent acquisition approaches might enable the comparison of hundreds of different samples without missing values, while maintaining the superb measurement precision and accuracy obtainable with isobaric tag quantification.

Keywords: analytical methods; isotopic labeling; mass spectrometry; proteins; proteomics.

Figure 1:. Outline of peptide identification with shotgun proteomics.

A) A sample of proteins is digested by trypsin, which cleaves peptide bonds at the C-terminus of lysine and arginine residues. The example peptide EIQTAVR, which we follow throughout this figure, is shown in blue. To reduce complexity, the peptides are separated by liquid chromatography (LC), ionized via electrospray, and injected into the mass spectrometer (MS). B) Plotted is the chromatogram of the most abundant peak at each retention time. The blue and green peptides elute at different retention times. C) At any given time, e.g., when the blue peptide elutes, multiple different peptides co-elute. The mass spectrometer can typically distinguish them by their mass to charge ratio (m/z). The mass spectrum of the intact peptides is called the MS1 spectrum. The peak corresponding to the peptide EIQTAVR is highlighted in blue. D) In complex mixtures, mass alone is not enough for peptide identification. Inside the mass spectrometer, a peak corresponding to a peptide is isolated and fragmented by collision with inert gas. The fragment ions’ m/z values, derived from the blue peptides, are recorded in the MS2 spectrum. By convention, peptide fragments containing the N-terminus are called b-ions, while fragments with the C-terminus are called y-ions. The characteristic masses of the fragment ions, together with the precursor mass from the MS1-spectrum, are typically sufficient to identify a peptide unambiguously.

Figure 2:. Outline of label-free and SILAC quantification.

A-C) Principles of label-free quantification. A) In label-free quantification, multiple protein samples are digested with trypsin (which cleaves after K or R). The resulting peptides are separated via liquid chromatography (LC) and ionized before entering the mass spectrometer (MS). Shown throughout are two peptides. The one ending in K has equal concentration in the two analyzed samples, and the one ending in R is concentrated 2-fold higher in the experimental sample compared to the control. B) The MS1 spectrum records the number of ions for various m/z values of the intact peptide eluting at a given time. C) The elution time of a peptide takes ~20 seconds. During this time, ~10 MS1 spectra are collected, each showing the peptide at potentially different intensities. The integration of this intensity over time approximates the total number of ions ionizing into the mass spectrometer. D-F) Principles of MS1 based quantification via heavy isotope labeling (e.g., SILAC). D) In SILAC, cell samples are grown either in media with amino acids with naturally occurring isotopes (light) or media where amino acids (K and R) contain heavy isotopes (here 6). Importantly, the heavy isotopes do not alter the chemical properties of the peptides. Cells are lysed and combined. The proteins are digested together, and the resulting peptides are simultaneously separated via LC and ionized before entering the mass spectrometer. E) Peptides in the heavy sample are shifted to the right on the MS1 spectrum compared to those from the light sample. Ratios between peak sizes within one spectrum can thus be used for relative quantification. F) To utilize all available information, typically the ion intensity is integrated over the entire elution profile.

Figure 3:. Comparison of Data-Dependent and Data-Independent Acquisition approaches.

A-E) Data-Dependent Acquisition (DDA). A) The goal of DDA is to identify as many peptides as possible, one at a time. The highest peaks in the MS1 spectrum are selected for isolation, with an isolation window of ~1 Th. Peptides in this window are isolated and fragmented for readout in the MS2 spectra. B) Shown is the sequence of MS1 spectra (black) and the data-dependent MS2 isolation windows (dark red), centered on the highest abundant peaks. Each MS1 is followed by multiple MS2 spectra - with current instrumentation and duty cycles of ~2 seconds this would be ~30 MS2 spectra following each MS1 spectrum. C) The MS2 spectrum resulting after isolation and fragmentation consists mainly of b- and y-ions from the target peptide, which allows for comparatively simple peptide identification. D) For quantification, e.g., with label-free approaches, the peptides in the MS1 spectrum are continuously monitored via the peak intensities in the MS1 spectra. Shown are the retention profiles for various peptides - the area under this curve is typically used for peptide quantification. The black line represents the single MS1 scan shown in B. E) Shown here are peaks for the b- and y-ions for the green and blue peptide. A peptide is typically only isolated once for MS2 analysis, the peak height cannot be used for quantification. Not all peaks in the MS1 will trigger the collection of an MS2 spectrum (peaks for the red peptide are missing). Additionally, low-abundance peaks might be below the detection limit in the MS1 spectrum and thus cannot trigger MS2 spectra. The dark red line represents the single MS2 scan shown in C. F-J) Data-Independent Acquisition (DIA). F) The goal of DIA is to continuously collect fragment ion intensities for all eluting peptides. To make this approach compatible with current MS speed requires significantly wider isolation windows (~10 Th) compared to the DDA approach (~1 Th). All the ions within this comparatively wide isolation window are isolated and simultaneously fragmented. G) Shown is the schedule of MS1 spectra (black) and the isolation windows of MS2 spectra (red). H) The simultaneous isolation and fragmentation of multiple peptides results in a complex MS2 spectrum consisting of b- and y-ions from all isolated peptides. I) Similarly to DDA, MS1 intensities of peptides are collected and can be used for quantification. The black line indicates the time for the MS1 spectrum in G. J) Unlike in the DDA equivalent, ion intensity information for b- and y-ions are available throughout the entire elution profile for each peptide. This makes it possible to use fragment ion intensities for quantification. Because the entire m/z space is continuously covered, information for more peptides than with the DDA approach is available. Here, the red peptide’s abundance can be quantified. The dark red line represents the single MS2 scan shown in H

Figure 4:. Outline of multiplexed proteomics with isobaric tags.

A) Isobaric tags have the same total mass, but differing distributions of heavy isotopes between the reporter group and mass balancer. Heavy isotopes are shown as asterisks. Peptides from 4 different samples are labeled with tags of the same mass, resulting in a single MS1 peak which can be isolated. With more tags (conditions), the complexity of the MS1 spectrum does not increase. This makes isobaric tags compatible with higher multiplexing (currently up to 11) compared to e.g., SILAC (see Fig. 2). B) After a peptide is isolated based on the MS1 spectrum, fragmentation will either cleave off the reporter ions, or lead to fragmentation of the peptide backbone. The reporter ions show different masses in the MS2 spectrum and can be used for relative quantification. Similarly, the intact peptide with the balancing groups, i.e., the complementary reporter ions, can also be used for quantification. The b- and y-ions are used for peptide identification (see Fig. 1).

Figure 5:. The problem of multiplexed proteomics: ratio distortion.

A) Even when using the smallest technically possible isolation window centered on a peptide of interest (red and dark blue), in a real experiment, other peptides with similar m/z and retention time will be co-isolated (pink and light blue). These interfering peptides will also be labeled with identical isobaric tags. B) Upon co-isolation and co-fragmentation, in the MS2 spectrum the low m/z reporter ions are identical, regardless of origin, and distort the quantification. Most background peptides tend to not change, showing a 1:1 ratio between control and experiment. The observed ratio for a peptide of interest, which changes 2-fold between control and experiment, will typically be compressed towards a 1:1 ratio.

Figure 6:. Strategies to overcome ratio distortion.

A-C) Overview of the MultiNotch MS3 approach A) The MultiNotch MS3 method acquires an MS2 spectrum similar to the standard approach by isolating a target peptide (red and dark blue), along with interfering peptides (pink and light blue). This spectrum is used for peptide identification. B) Instead of quantifying the reporter ions in the MS2 spectrum, the highest abundant peaks, which typically are b- and y-ions from the peptide of interest, are simultaneously isolated and further fragmented for an MS3 spectrum. C) Reporter ions in the MS3 spectrum are used for quantification. The additional gas-phase purification typically leads to removal of most interfering signal. While not perfect, the measured ratios are typically significantly more accurate than with a standard MS2 approach. D-E) Overview of the complementary reporter ion quantification strategy. D) For the complementary reporter ion method, a standard MS2 spectrum is acquired, which will co-isolate and co-fragment the peptide of interest and interfering species. E) The low m/z reporter ions show interference, as discussed in Figure 5. However, this method involves analyzing the complement reporter ions in the MS2 spectrum, where the peptide is still attached to the mass balancer group. Since the target peptide and interfering peptides typically have slightly different masses, this allows them to be distinguished with a high resolution mass analyzer like an Orbitrap. The results are significantly more accurate quantification compared to MS2 and even the MS3 approach. While interference still will occasionally lead to ratio distortion, to our knowledge this method currently generates the most accurate data.

Figure 7:. Overview of isobaric tags.

In each structure, asterisks denote heavy isotopes. The black part of the structure indicates the reporter part, the balancing group is blue, and the leaving group, which is removed after the tag reacts with the peptides, is red A) The original 2-plex TMT from Thompson et al. B) Current commercial TMT, which can encode up to 11 different conditions (See Fig. S1 for heavy isotope distribution). C) In the iTRAQ structure, the marked oxygen can be either ¹⁶O or ¹⁸O. D) The DiLeu-tag is a 4-plex tag developed by Xiang et. al. E) Braun et al. developed combinatorial tags. These tags generate multiple reporter ions, which allows for high multiplexing capacity for a given number of heavy isotopes. After fragmentation at the shown cleavage site reporter 1 forms. However, this further fragments into reporter 2 and a neutral loss. F) Stadlmeier et al. developed the sulfoxide-based tag, which is optimized for complement reporter ion formation due to fragmentation of the sulfoxide bond at lower energies. The two tertiary amines result in higher charge states of peptides after ionization and further facilitate fragmentation. G) The EASI-tag developed by Winter et al. similarly fragments comparatively easily. The “reporter” part of the EASI-tag is a neutral loss. Therefore, quantification with the EASI-tag is only possible via the complement reporter ions.

Figure 8:. Proposed fusion of DIA with multiplexed proteomics.

A) Peptides would be labeled with isobaric tags similar to a normal multiplexing experiment. For sake of simplicity, we only show two conditions. Like in a normal DIA workflow, all the peaks within a certain wide m/z window in an MS1 scan are co-isolated. This window contains multiple peptides, which will all be simultaneously isolated and fragmented into an MS2 spectrum. For simplicity, only two peptides are shown in detail, depicted with solid and dashed outlines. B) In the MS2 spectrum, the low m/z reporter ions cannot be used for quantification since the reporter ions of all co-isolated peptides will be identical. However, simultaneous quantification is possible via the peptide complement reporter ions. Additionally, complementary b- and y-ions that additionally lost their reporter group can also be used for peptide specific quantification. C) The continuous monitoring of peptide complement reporter ions and b- and y-fragment complement reporter ions allow the relative quantification of multiplexed abundances even between various runs. Additionally, the number of missing values in samples larger than the multiplexing capacity of a single tag should be drastically reduced.