Low-pH Solid-Phase Amino Labeling of Complex Peptide Digests with

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Low pH Solid Phase Amino-Labelling of Complex Peptide Digests with TMTs Improves Peptide Identification Rates for Multiplexed Global Phosphopeptide Analysis Gitte Böhm, Petra Prefot, Stephan Jung, Stefan Selzer, Vikram Mitra, David Britton, Karsten Kuhn, Ian Pike, and Andrew Hugin Thompson J. Proteome Res., Just Accepted Manuscript • DOI: 10.1021/acs.jproteome.5b00072 • Publication Date (Web): 04 May 2015 Downloaded from http://pubs.acs.org on May 5, 2015

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Journal of Proteome Research

Low pH Solid Phase Amino-Labelling of Complex Peptide Digests with TMTs Improves Peptide Identification Rates for Multiplexed Global Phosphopeptide Analysis

Gitte Böhm1, Petra Prefot1, Stephan Jung1, Stefan Selzer1, Vikram Mitra2, David Britton2, Karsten Kuhn1, Ian Pike2, Andrew H. Thompson*2 1

Proteome Sciences R&D GmbH & Co. KG, Altenhöferallee 3, 60438 Frankfurt am Main, Germany

2

Proteome Sciences Plc, Coveham House, Downside Bridge Road, Cobham KT11 3E, UK

ABSTRACT: We present a novel Tandem Mass Tag Solid Phase Amino Labelling (TMT-SPAL) protocol using reversible immobilization of peptides onto octadecyl-derivatised (C18) solid supports. This method can reduce the number of steps required in complex protocols saving time and potentially reducing sample losses. In our global phosphopeptide profiling workflow (SysQuant) we can cut 24 hours from the protocol while increasing peptide identifications (20%) and reducing side-reactions. Solid phase labelling with TMTs does require some modification to typical labelling conditions particularly pH. It has been found that complete labelling equivalent

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to standard basic pH solution phase labelling for small and large samples can be achieved on C18 resins using slightly acidic buffer conditions. Improved labelling behaviour on C18 compared to standard basic pH solution phase labelling is demonstrated. We analysed our samples for histidine, serine, threonine and tyrosine-labelling to determine the degree of over-labelling and observed higher than expected levels (25% of all Peptide Spectral Matches (PSMs)) of overlabelling at all of these amino acids (predominantly at tyrosine and serine) in our standard solution phase labelling protocol. Over-labelling at all these sites is greatly reduced (four-fold to 7% of all PSMs) by the low pH conditions used in the TMT-SPAL protocol. Over-labelling seems to represent a so-far overlooked mechanism causing reductions in peptide identification rates with NHS-activated TMT-labelling compared to label-free methods. Our results also highlight the importance of searching data for over-labelling when labelling methods are used.

Keywords: Mass spectrometry; Proteomics; Chemical labelling; Phosphorylation; Isobaric tags; TMT Introduction: Phosphopeptide workflows typically have a relatively large sample requirement to enable detection of low abundance phosphopeptides. Samples are often fractionated by Strong Cation Exchange (SCX) or Basic Reverse Phase chromatography with subsequent phosphopeptide enrichment. Immobilised Metal Affinity Capture (IMAC) with various metal ions such as Fe3+ 1, Ga3+ 2, Ti4+

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and Titanium dioxide (TiO2) resins4 have been shown to be highly effective and

complementary strategies for phosphopeptide enrichment that are often used in tandem5. It is not unusual for a phosphopeptide analysis workflow to combine analysis of SCX or high pH Reverse Phase fractionated and IMAC and TiO2 enriched samples with parallel analysis of unenriched


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fractionated sample material5. This may amount to 20 or more LC-MS/MS analyses for a single sample and obviously this requires a significant amount of sample material. Isobaric mass tagging6 with TMT® is a useful tool for the simultaneous analysis of up to 10 samples in one multiplexed experiment7-9, making long complex workflows more cost-effective. The ability to multiplex samples enables the efficient use of instrument time and reduces the occurrence of missing data when making comparisons between multiple samples. However, isobaric mass tag reagents are relatively expensive and labelling large amounts of sample becomes a limiting factor in such analyses. Furthermore, the large sample requirement requires multiple steps to reduce the large sample volumes, after cell lysis, reduction, alkylation and trypsin digestion, to provide the peptide digest in a volume suitable for cost-effective use of the labelling reagent. Our own phosphopeptide analysis workflow combines TMT-labelling with SCX fractionation and parallel IMAC and TiO2 phosphopeptide enrichment10 (see Figure 1). This workflow currently achieves this effectively but requires a C18 desalting step after the reduction, alkylation and trypsin digestion steps followed by an overnight drying step to reduce solvent volumes and enable peptide solubilisation in a buffer composition and volume suitable for efficient and economical TMT labelling. A further C18 desalting step is required after TMT labelling. The motivation for the work presented here was to find an abbreviated protocol that reduces the number of steps in our workflow thus reducing the time required, consumable use and potential for sample loss. Since a number of publications have shown successful labelling of peptides using reversible solid phase immobilization of peptides onto C18 resins for guanidination of lysine epsilon amino groups11, sulphonation of amino groups12, methylation of amino groups13


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and Beta-elimination/Michael addition at phosphate groups14, we expected that solid phase labelling might shorten our workflow. Solid phase labelling reactions on C18 are reported to concentrate both the target (peptide digest) and label11, if the label is sufficiently hydrophobic like TMT reagents, in a thin film of organic solvent on the C18 surface. This concentration effect can increase reaction rates and, for our purposes, it allows us to avoid the overnight drying step of the current protocol by allowing concentration of large volumes of peptide digest directly on the C18 resin followed by in situ buffer changes, TMT-labelling and desalting. In this way we replace two desalting processes, an overnight

drying

step

and

a

separate

labelling

step

with

a

single

on-column

concentration/labelling/desalting process with no overnight drying. To our knowledge, however, solid phase labelling of amino groups with NHS-activated reagents has not been described in the literature so far. Such an approach has to deal with the use of aqueous stock solutions of label reagents, to ensure a proper binding of the peptides during loading and incubation. However, aqueous stock solutions of NHS-activated label reagents are prone to hydrolysis of the reagent before the desired labelling of amino groups actually takes place. Thus, it was important, as with any new protocol, to ensure that the new protocol performed at least as well as the previous protocol. In addition to a shortened protocol, our results highlight a number of interesting discoveries in relation to labelling reaction pH, overlabelling and the use of solid phase materials, which are analysed and discussed in depth. Materials and Methods:


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Figure 1: Schematic showing both our standard Solution Phase TMT-labelling conditions and our shortened TMT-SPAL labelling conditions for global phosphopeptide analysis. Sample Materials: a single homogenous, pooled MCF7 cell line sample (48mg protein) was generated to provide enough starting material to directly compare our SysQuant® protocol with


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either standard solution phase TMT-labelling or our new TMT-SPAL labelling protocol. The sample material was split into 3 mg aliquots for further analysis. Tissue cell lysis: Frozen clinical tissue samples were pulverized using a pestle and mortar in the presence of liquid nitrogen and then transferred to Eppendorf tubes containing 1.3ml of icecold lysis buffer (8M urea, 75mM NaCl, 50mM Tris-pH 8.2, one tablet of protease inhibitors cocktail (complete mini, Roche) per 10ml of lysis buffer, and one tablet of phosphatase inhibitor cocktail (Roche) per 10mL of lysis buffer). Samples were then sonicated (Branson Digital Sonifier W450D) at 20% amplitude for 20 x 1 second, pulsing on and off, on ice (4°C). Following centrifugation at 12,500 g for 10 min at 4oC, the protein concentration of each sample was then determined using by Bradford protein assay. In-Solution Trypsin Digestion: Reduction, alkylation of cysteines, and digestion was performed on each lysate by following the Villén and Gygi, Nature Protocol5. The digested samples were spun for 10 minutes at 2,500 g and de-salted on 200 mg SepPak tC18 cartridges (Waters, Milford, MA, USA). Peptides were eluted with 50% acetonitrile (ACN)/0.1% trifluoroacetic acid (TFA) and then dried down in a Speedvac system comprising the following hardware: Speedvac: Univapo 150H from Uniequip; Cold Trap: Alpha 1-4 from Christ and Vacuum Pump: RC5 from Vacuubrand. Solution Phase TMT Labelling at basic pH: Digested peptides from all samples were separately re-suspended in 200 mM triethylammonium bicarbonate (TEAB)/10% ACN, mixed with their respective TMT (15 mM final concentration) in an 8-plex experimental design, and left to incubate for 1 hour at room temperature. TMT reactions were terminated by addition of hydroxylamine to give a final concentration of 0.25%. Quenching was allowed to proceed for 15 minutes. Samples were pooled into a TMT 8-plex and left to incubate for another 15 minutes.


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Each TMT 8-plex sample was acidified and the acetonitrile concentration diluted to below 5%, then divided into three aliquots each of which were desalted on a 200 mg SepPak tC18 cartridge and eluted. Individual TMT-labelled samples were pooled at this point and lyophilized. TMT Solid Phase Amino Labelling (TMT-SPAL): Waters SepPak C18 cartridges with 200 mg of resin were activated with a single 3 ml wash with Conditioning Solvent (95% ACN, 4.9% water and 0.1% TFA). Samples of peptide digest were diluted to less than 5% organic solvent/0.5% TFA and loaded onto the C18 cartridges. The C18 cartridges were washed with 1 ml of 50 mM KH2PO4 (pH 4.5). For solid phase labelling, a 300 mM TMT stock solution in fresh/dry acetonitrile was prepared. Immediately prior to use, the TMT stock was diluted in sufficient 50 mM KH2PO4 to reduce the acetonitrile content down to 5% (15 mM) TMT to avoid peptide elution during tag loading. This TMT solution was loaded onto the column immediately. For 3 mg of peptide digest on a 200 mg SepPak cartridge, 15 µmol of TMT was loaded (50 µL of 300 mM TMT stock diluted with 1000 µL 50 mM KH2PO4). Each of the 8 samples was labelled with a different TMT tag. The TMT reagents bind to the resin and should not be present in the flow-through. Samples are left on column for 1 hour at RT taking care to make sure the resin stays wet. No quenching step was applied for the solid phase labelling. After one hour, the C18 cartridge was washed with 1 ml of a standard C18 desalting Wash Solvent (5% ACN in water with 0.1% TFA). Labelled peptides were eluted with 1.5 ml of Elution Solvent (50% ACN in water with 0.1% TFA). Individual TMT-labelled samples were pooled at this point and lyophilized. Strong Cation Exchange-HPLC: All aliquots of lyophilized peptides were re-suspended in SCX loading buffer C (7 mM KH2PO4, pH 2.65, 30% ACN (vol/vol)), then separated into 12 fractions by SCX-HPLC chromatography. The fractionation was carried out using a


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polySULFOETHYL-A column (PolyLC) in a Waters Alliance 2695 HPLC system according to Villén and Gygi 5. Fractions were pooled according to our scheme shown in Supplementary Table 1 to give 8 fractions for the enrichment arm and 7 fractions for the non-enrichment arm. For subsequent enrichment of phosphopeptides, 10 mg of digested, TMT-labelled and pooled peptide digest were loaded per enrichment approach. A third SCX run with 3 mg peptide material was performed for the analysis without enrichment of phosphopeptides. Phosphopeptide Enrichment: Each SCX fraction for phosphopeptide enrichment was split into two aliquots. The two aliquots of each SCX fraction were enriched for phosphopeptides in parallel by IMAC (Thermo Scientific Pierce product code 88300) and TiO2 (Thermo Scientific Pierce product code 88301), in accordance with manufacturer’s instructions. Graphite Spin Columns. Following phosphopeptide enrichment, peptides were purified using graphite spin columns (Thermo Scientific Pierce product code 88302), according to manufacturer’s instructions. Liquid Chromatography – Tandem Mass Spectrometry (LC-MS/MS). Peptides from non-enriched fractions were re-suspended in 100 µl of 2% ACN/0.1% formic acid (FA), then 5 µL injected onto a 75 µm x 2 cm nanoViper C18 Acclaim PepMap100 precolumn (3 µm particle size, 100Å pore size; P/N 164705; Thermo Scientific) with an additional sample loading volume of 12 µL 0.1% FA in H2O, using the Thermo Scientific EASY-nLC 1000 system. Peptides were then separated over a 180-minute gradient starting with 0.1% FA in ACN (8-30% over 160 minutes, then 30-100% between 160-170 minutes, continuing at 100% up to 180 minutes) through a 75 µm x 50 cm PepMap® RSLC analytical column @ 40oC (2 µm particle size, 100Å pore size; P/N ES803; Thermo) at a flow rate of 200 nL/min.


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Peptides from all phospho-enriched fractions were re-suspended in 30 µl of 2% ACN/0.1% FA, then 5 µL of each sample were injected onto the pre-column with an additional sample loading volume of 12 µL 0.1% FA in H2O. The peptides were then separated using the same 180-minute gradient described above for non-enriched fractions. Mass spectra were acquired on an Orbitrap FusionTM TribridTM Mass Spectrometer (Thermo Scientific) throughout the entire chromatographic run (180 minutes), using top speed higher collision induced dissociation (HCD) FTMS2 scans at 30,000 resolving power (FWHM at m/z 200), following each FTMS scan (120,000 resolving power (FWHM at m/z 200)). HCD was carried out on the most intense ions from each FTMS scan within a user defined cycle time of 3 seconds, then put on a dynamic exclusion list for 30 secs (10 ppm m/z window). AGC ion injection target for each FTMS scan were 500,000 (100 ms max injection time). AGC ion injection target for each HCD FTMS2 scan were 10000 (100 ms max ion injection time). Instrument settings are summarized in Supplementary Table 2. Peptide identification and quantification. Proteome Discoverer In total there were 30 Raw data files (2 x TMT 8-plex samples (Solution Phase vs TMTSPAL) X 15 fractions X 1 replicate from the Fusion instrument. The 15 raw data files for the TMT-SPAL and the 15 from the Solution Phase 8-plex samples were analysed together as single MudPIT searches using Proteome Discoverer, as described below. Total Peptide Identifications: Supplementary Figure 1 shows a screenshot of our Proteome Discoverer workflow for determining peptide identifications. Raw data were submitted to the Thermo Scientific Proteome Discoverer 1.3 software, using the Spectrum Files node. Spectrum selector was set to its default values, while the Mascot node15, was set up to search data against


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the uniprot_sprot database, taxonomy homo sapiens (UniprotKB Homo Sapiens complete proteome canonical + isoforms/Release 2014_01). This node was programmed to search for tryptic peptides (two missed cleavages: missed cleavage data for both protocols is shown in Supplementary Figure 2 indicating very little difference between the two protocols in this respect) with static modifications of carbamidomethyl (C), TMT 6/8-plex (K), and TMT 6/8-plex (N-Term). Dynamic modifications were set to deamidation (N/Q), oxidation (M), and phosphorylation of STY. Precursor mass tolerance was set to 20 ppm and fragment (b and y ions) mass tolerance to 20 mmu. Spectra were also searched against SEQUEST16, using the same database, modifications, and tolerances as the Mascot node. For total peptide identification purposes, SEQUEST and Mascot identifications were merged and FDRs homogenized using the Percolator17 node. Spectra were also searched using the PhosphoRS 2.018 (fragment mass tolerance of 20 mmu, considering neutral loss peaks for CID and HCD). For speed, additional searches were carried out with just SEQUEST to determine rates of labelling efficiency as well as over-labelling. In these searches, the SEQUEST node was programmed to search for tryptic peptides (two missed cleavages) with the following modification settings: Labelling efficiency: static modifications of carbamidomethyl (C) and TMT 6/8-plex (K). Dynamic modifications were set to TMT 6/8-plex (N-Term). Over-labelling: static modifications of carbamidomethyl (C), TMT 6/8-plex (K), and TMT 6/10-plex (N-Term). Dynamic modifications were set to TMT-labelling of HSTY together, to determine total as well as individual over-labelling. Site-specific over-labelling was quantified by an in-house R script that separated out the individual amino acid over-labelling rates from the total over-labelling data.


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The reporter ions quantifier node was set up to measure the raw intensity values of TMT 8plex mono-isotopic ions, from all identified PSMs, at; 126.12773 m/z (126), 127.12476 m/z (127e), 127.13108 m/z (127), 128.13444 m/z (128), 129.13147 m/z (129e), 129.13779 m/z (129), 130.14115 m/z (130), 131.13818 m/z (131), using a tolerance of 20ppm after centroiding. No filters were applied at this stage using Proteome Discoverer, therefore all raw intensity values were exported to excel for later processing and filtering using in-house software. Results and Discussion: New Reaction Conditions: For comparison, the same total amounts of TMT reagent were used for both the solution phase and TMT-SPAL reactions. However, since high organic solvent concentration will elute peptides bound on C18, the TMT stock solution for Solid Phase labelling had to be made up in acetonitrile at a much higher concentration (300mM) than the solution phase reaction (60mM) to ensure that the final 15mM TMT loading solution comprised less than 5% acetonitrile. Fortunately, the TMT reagent is soluble in acetonitrile at this high concentration. Immediate use of the TMT solutions, after dilution with aqueous buffer, prevented significant hydrolysis of the NHS-activated TMT reagents prior to peptide exposure. It is worth noting that the TMT reagents bind to the C18 resin when loaded in a buffer with 5% acetonitrile. This means the TMT reagent could be loaded onto the C18 column with lower concentrations of TMT in larger loading volumes if that is desirable allowing a less concentrated stock solution or a lower final acetonitrile concentration to be used, if desirable. Initial experiments on a synthetic peptide showed that the standard basic buffer (100 mM TEAB, pH 8.5) does not result in successful TMT-labelling on the C18 solid phase. Figure 2 shows HPLC traces (UV detection at 214 nm) from solid phase labelling of a synthetic peptide


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(Sequence: VATVSLPR). We use the VATVSLPR peptide extensively for checking labelling: the HPLC peaks in Figure 2 at 12.1 and 13.1 minutes have been MS-confirmed as unlabelled and correctly labelled respectively (data not shown). In Figure 2a, it can be seen that on-column labelling with TEAB resulted in inconsistent labelling (putatively over-labelling of serine and threonine since this peptide does not comprise tyrosine) without the intended reaction going to completion even with repetitive loading of TMT onto the C18 column. We hypothesized that lower pH conditions might improve labelling and a brief pH scouting experiment (See Supplementary Figures 3 and 4 and Supplementary Table 3) indicated that pH 4.5 was preferred. In Figure 2b, it can be seen that a slightly acidic buffer (50 mM KH2PO4, pH 4.5) produces consistent and complete TMT-labelling equivalent to the standard solution phase labelling without measurable over-labelling after 1 hour on column with the same amount of TMT reagent as the solution phase protocol. No hydroxylamine quenching was used in the TMT-SPAL reactions as hydroxylamine does not bind to C18 and washing the hydroxylamine through the column appeared to be ineffective in reversing any over-labelling in our pH 8.5 experiments (data not shown). Reaction times (~1 hour) and reagent quantities required for TMT-SPAL are essentially the same as for the Solution Phase reaction as long as appropriate buffer is used. 2a

2b


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Figure 2: HPLC traces of peptide labelling reactions detected with UV absorbance (214 nm). Using the standard TEAB buffer, we observe incomplete labelling of peptide even with repeated incubations with fresh TMT reagent and we also see evidence of over-labelling. In contrast, labelling at pH 4.5 gives complete labelling of peptide in 60 minutes with the standard amount of TMT without evidence of significant over-labelling. Use of Solid Phase Supports for labelling of large amounts of peptide digest: Following our initial results showing that we can get complete TMT-labelling on a single synthetic peptide using the TMT-SPAL approach, we repeated the experiments on the same peptide using a similar amount of the peptide as we typically use of peptide digest for our SysQuant phosphopeptide enrichment protocol, i.e. about 3 mg of peptide. We demonstrated that C18 cartridges can readily capture 3 mg of synthetic peptide from a large volume of buffer and retain the peptide on the solid support (data not shown) for labelling. Optimal peptide retention and recovery performance was found using 200 mg SepPak tC18 cartridges for desalting of the TMT-labelled peptides. 100mg tC18 cartridges sometimes resulted in sample losses despite the stated binding capacity being apparently adequate. Similarly, we


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observed that HLB columns lose some peptide when the pH is changed (data not shown) so HLB cartridges are not appropriate for the TMT-SPAL protocol. This is similar to a previous report from the Heck group who used C18 for reductive methylation of peptides and they suggest allowing at least twice the manufacturer’s stated capacity over the expected sample amount to ensure robust labelling behaviour13. TMT-SPAL labelling using 200 mg tC18 SepPak cartridges on 3 mg of peptide using the pH 4.5 conditions produced clean HPLC traces like Figure 2b indicating that labelling of large samples on the solid phase is feasible. Robustness and Completeness After demonstrating complete labelling of synthetic peptide at large scale with TMT-SPAL and high capacity cartridges, we further tested the TMT-SPAL protocol on a complex biological sample (Human MCF7 cell line material) at 3mg labelling scale comparing this with Solution Phase labelling in our complete phosphopeptide workflow10. After sequence analysis of the raw data from all fractions (phosphopeptide enriched and un-enriched fractions from both labelling approaches), we observed that we are getting virtually complete labelling of a complex sample too with similar overall labelling rates for both the standard Solution Phase and TMT-SPAL protocols (see Figure 4). We also assessed ‘over-labelling’, i.e. unwanted labelling at the reactive amino-acid sidechains histidine, serine, threonine and tyrosine (HSTY) using a SEQUEST search with labelling at these side-chains included along with the canonical amino groups. Clearly, the search space increases dramatically when modification at multiple side-chains is considered but we used a standard decoy database, 1% FDR approach19 to control for false positives. If the SEQUEST searches were finding a lot of false positive over-labelling hits one might expect the over-


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labelled peptide XCorr scores to be lower (for a given intensity) than correctly labelled peptides. Furthermore, the mean XCorr scores and mean intensities would be lower for the over-labelled peptides than the correctly labelled peptides. However, it can be seen in Supplementary Table 3 that these means are approximately equal. Similarly, Supplementary Figures 5a and 5b show scatter-plots of the XCorr values versus the precursor intensities for the Solution Phase data and the TMT-SPAL data respectively. Correctly labelled and over-labelled scatter-plots are overlaid and it can be seen that the distributions are equivalent indicating that the over-labelled species are not likely to be false positives or artefacts of the search. Figure 3 shows the proportions of labelled and over-labelled (HSTY-labelled) peptide spectral matches (PSMs) in the Solution Phase labelled MCF-7 cell line material (bottom bar) and in the MCF-7 cell line labelled by TMT-SPAL (top Bar). Note that only the unenriched fractions from the data set are included in the data (Figure 3) to avoid distortion of the results by phosphopeptide enrichment. In Figure 3, we observe a significantly increased yield of correctly labelled PSMs with TMT-SPAL. We observe a certain proportion of over-labelled PSMs in both samples with over-labelling at all the potential HSTY over-labelling sites. However, we see a major reduction in over-labelling at all the HSTY sites with the TMT-SPAL reaction conditions even without the hydroxylamine quenching step used in our Solution Phase labelling protocol. In total, 25.6 % of PSMs in the Solution Phase data had over-labelling at one or more of the possible HSTY sites while only 7.6% of the PSMs had over-labelling in the TMT-SPAL data.


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Figure 3: Bar Chart showing the total numbers of PSMs identified in our TMT-SPAL data (top) and in our Solution Phase data (bottom). The number of correctly labelled PSMs (dark blue) is considerably increased in the TMT-SPAL sample. Relative proportions of PSMs that were improperly labelled with labelling at histidine (red), labelling at serine (green), labelling at threonine (purple) labelling at tyrosine (light blue) are shown. Figure 4 shows log10 ion intensity histograms for the Solution Phase and TMT-SPAL protocols. The ratio of correctly labelled species to over-labelled species is substantially higher in the TMT-SPAL protocol compared to the Solution Phase protocol. Moreover, the overall numbers of correctly labelled peptides in the low intensity bins are particularly increased with TMT-SPAL, while over-labelled species, which are typically lower abundance species are reduced compared to the Solution Phase labelling. Thus, with appropriate pH on the C18


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substrate, we get improved labelling behaviour from the TMT-SPAL protocol (Figure 4b) compared to the Solution Phase labelling protocol (Figure 4a). We speculate that the effect of over-labelling on relative peptide intensities is based on the fact that ionisation in mass spec is competitive and that the TMT structure comprises a Hunig’s base, i.e. a very strong base. Over-labelled species with two or more tags are thus likely to protonate even more strongly than correctly labelled species with one or two tags and could competitively ionise more strongly than correctly labelled species. Conversely, a reduction in over-labelling should, in theory, reduce competition allowing greater protonation of correctly labelled species and consequently producing higher signal intensities. In principle, reducing over-labelling could therefore increase the chance of DDA selection of correctly-labelled species and certainly the proportion of peptides identified as correctly labelled does appear to go up while the number of species identifiable as over-labelled seems to be reduced in TMT-SPAL. In addition, newer instrumentation with higher sensitivity (UHPLC interface coupled to an Orbitrap Fusion) is able to detect lower abundance species and is likely to detect a higher proportion of lower abundance species than older machines. 4a


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Figure 4: Figures 4a and 4b show log10 ion intensity histograms for the Solution Phase labelling protocol and the TMT-SPAL protocols respectively. Separate overlaid histograms are shown for TMT-labelled (blue), unlabelled (red) and over-labelled peptides (green). Scales are the same in both Figure 4a and 4b. The two phosphopeptide analysis workflows reported here have been assessed without replication. Given the large sample requirement for our workflow, multiple replicates were not practical. We have however, compared the TMT-SPAL and Solution Phase labelling conditions on two other different sample types, rat cortex and human plasma digest (Supplementary Table 5). These two different sample types were run as 1-dimensional LC-MS studies in contrast to the 2-dimensional protocol used for the phosphopeptide analysis. We observe the same overall trend in relation to increased identifications and over-labelling as in our phosphopeptide analysis leading us to conclude that the low pH reaction conditions reducing over-labelling is a genuine and consistent effect. However, in the human plasma digest we do observe slightly less complete labelling with TMT-SPAL (~94%) than the standard conditions. We speculate that this may reflect the more complex protein composition and wide dynamic range of protein abundance of plasma and that higher tag concentrations may be required for complete labelling of plasma samples by TMT-SPAL. Peptide, phosphopeptide, protein and phosphoprotein identification rates: To do determine the overall performance of both protocols, we carried out a standard analysis of the MCF7 enriched and unenriched fractions for both protocols using SEQUEST and Mascot followed by post-search re-analysis combining the SEQUEST and Mascot results with Percolator at 1% False Discovery Rate (FDR) searching for correctly TMT-labelled peptides with phosphates on serine, threonine and tyrosine as variable modifications (Supplementary Figure 1).


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Post-search analysis with PhosphoRS was also used to validate site localization of phosphate groups in phosphopeptides. This analysis shows that the TMT-SPAL protocol results in a higher level of overall peptide and phosphopeptide identifications (Figure 5 (left)). These numbers treat variants of a peptide as different as this represents the total identification rate of the labelling chemistry rather than being representative of the biology.

Figure 5: The Bar Chart shows the total numbers of species identified along with the total number of phosphorylated species and non-phosphorylated species in our Solution Phase protocol compared with our TMT-SPAL protocol. The Bar Chart shows the total numbers of peptides identified on the left with the total number of ungrouped proteins, phosphoproteins and non-phosphorylated proteins identified shown in the centre and with the total number of grouped proteins, phosphoproteins and non-phosphorylated proteins shown on the right.


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TMT-SPAL SysQuant detects about 20% more unique peptides and about 18% more phosphopeptides compared to the Solution Phase protocol over the whole dataset (Figure 5 (left)). The numbers of proteins identified without grouping of protein variants shows a smaller increase in proteins ids where ~2% more proteins and ~11% more phosphoproteins are seen in the TMT-SPAL data (Figure 5 (centre)). The final numbers of protein groups identified translates into a slightly lower increase in protein identifications in TMT-SPAL where ~7% more proteins and ~11% more phosphoproteins are detected compared to the Solution Phase (Figure 5 (right)). However, the sequence coverage in the TMT-SPAL data for each protein is higher on average with a mean number of 13.2 peptides per protein identified in the TMT-SPAL data compared to 11.6 peptides per protein in the Solution Phase data. The higher average number of peptides identified per protein and the higher overall peptide and phosphopeptide identification indicates that we are obtaining better protein coverage with TMT-SPAL and the higher phosphoprotein identification rates suggest we have a slightly higher chance of recording phosphate regulation with the new protocol. Figure 6a shows a Venn diagram with the overlap of identification rates of unique peptides identified by each protocol as shown in Figure 5a while Figure 6b shows the corresponding ungrouped protein identification rates between the TMT-SPAL protocol and the Solution Phase labelling protocol corresponding to the ungrouped proteins in Figure 5b. It shows that overall the peptide identification rates (~70%) and specific proteins (~80%) identified largely overlap, demonstrating that there is no loss from using the new approach. However, there is still a significant number of proteins (~20%) identified by the Solution Phase protocol that are not seen in the TMT-SPAL data. Conversely, there are about 22% of


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proteins seen in the TMT-SPAL data that are not present in the solution phase data and there are substantially more peptides that differ between the protocols. 6a - Peptides

6b – Proteins (ungrouped)

Figure 6: Venn diagram showing the overlap of the number of Peptide Identifications (6a) and Protein Identifications (6b) between Solution Phase TMT-labelling and labelling by TMT-SPAL from the analysis of the Breast Cancer MCF7 cell lines. These figures correspond to the total identification rates in Figures 5 (left) and 5 (centre) for peptides and proteins respectively. Further analysis of Peptide Identification Rates: The difference in the numbers of peptides uniquely identified by the two protocols may be down to typical inter-sample variations in DDA based analysis but we cannot exclude the possibility of a systematic difference. We performed further analysis to assess why the TMTSPAL protocol gives improved peptide identification rates and also to determine whether there is indeed any systematic bias in the peptides identified by either protocol. We observe that there are systematic differences between the two methods consistent with the use of the more hydrophobic organic conditions of TMT-SPAL with a bias towards shorter and more hydrophilic peptides being uniquely present in the Solution Phase protocol and more hydrophobic peptides in TMT-


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SPAL (Supplementary Figures 6 and 7). We observe a similar bias in histograms showing peptide and phosphopeptide identifications versus retention time (Supplementary Figures 8a and 8b respectively) resulting in a slight reduction in the numbers of peptides and phosphopeptides identified by TMT-SPAL when compared with the Solution Phase protocol at the earliest retention times indicating a slight bias against more hydrophilic peptides by TMT-SPAL. Conversely, there is an increase in the numbers of peptides identified by the TMT-SPAL protocol at later retention times indicating a bias towards more hydrophobic species with similar biases at both the peptide and phosphopeptide level. We used 5% acetonitrile in our C18 sample loading and wash steps, which is the typical amount recommended by most manufacturers for desalting columns and in most published protocols for desalting. This percentage of organic solvent is used in both the Solution Phase and TMT-SPAL protocols and may be adversely affecting recovery of hydrophilic peptides in both protocols and lower organic solvent conditions might be preferable. The concentration of TMT at loading was adjusted so that the same concentration of TMT was used in TMT-SPAL as our Solution Phase protocol but with greatly reduced acetonitrile. Interestingly, TMTs seem to adhere to the C18; thus, the loading concentration is probably immaterial and lower acetonitrile could be used although the issue of TMT hydrolysis does need to be considered at low acetonitrile concentrations. Analysis of the effect of TMT over-labelling on Peptide Identification rates: In addition to our analysis of systematic differences between the standard Solution Phase protocol and TMT-SPAL, we tested our hypothesis that reduction of over-labelled species in the TMT-SPAL protocol is leading to increased identification rates with this protocol (see Supplementary Material). Our hypothesis is based on the fact that a finite number of MS2 spectra


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are acquired in a given analysis time on a given instrument and we see a proportionate increase in correct identifications that is of a similar magnitude to the decrease in the percentage of overlabelled spectra. In the Solution Phase data ~60% of Tyrosine over-labelled peptides are redundantly detected in the same dataset as correctly labelled while ~80% of these over-labelled peptides are detected as correctly labelled in the TMT-SPAL data. The overall figure for Tyrosine over-labelling drops dramatically in the TMT-SPAL data (~4-fold decrease in overlabelling) and of those few peptides that are detected ~90% are detected redundantly as correctly labelled in the same dataset. The reduction in redundant detections in TMT-SPAL suggests that the mass spectrometer is spending more time detecting new species. Similarly, since the overlabelled species are typically lower abundance species (Figure 4), it would be expected that most of the additional identifications in TMT-SPAL would be lower abundance species and this does appear to be the case (Supplementary Figure 9). Thus, it would appear that reduced redundancy of identifications contributes greatly to the enhanced peptide identification rate of the TMTSPAL protocol compared to the Solution Phase protocol on our high performance Orbitrap Fusion instrument and that these additional identifications are primarily lower abundance species. It has previously been reported in the literature that labelling with isobaric mass tags reduces peptide identification rates, for an equivalent amount of chromatographic separation and mass spectrometer analysis time compared to label-free analysis. Thingholm et al.20 attribute the drop in peptide identification to an increase in the number of charge states accessible to a labelled peptide compared to unlabelled peptide due to the increase in mass and size of the labelled species. Similarly, Pichler et al.21 attribute the loss of identifications to both the increase in size of the peptide (and thus accessible charge states) and also to the increase in tag-related fragment


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ions present in the MS2 fragmentation spectra of labelled peptides, which are not properly accounted for by current peptide identification software. In addition, the reporters from isobaric tags effectively act as additional pseudo-sequence ions, which split the MS2 ion signal over more peaks for a given peptide compared to the unlabelled counterpart with the effect that some sequence ions may be weakened in the labelled peptide compared to unlabelled for a given amount of peptide ion potentially reducing identification rates22. While all these mechanisms appear to be relevant, they do not explain completely the loss of peptide identifications and we now believe over-labelling reactions may also contribute significantly to loss of peptide identifications when using TMT-labelling particularly on high performance instruments such as the Orbitrap Fusion, which show improved detection of lower abundance species that might not previously have been detectable. Accuracy and Precision of Quantification of TMT-SPAL: The experiments carried out in this paper were performed using MS2 analysis of the data and all the 8-plex samples were made up in a 1:1:1:1:1:1:1:1 ratio so a full assessment of quantitative accuracy cannot be made. The only observation that can be made is that the reporter ion statistics are broadly similar (Supplementary Figures 10a and 10b) and that at this stage we believe that it is likely that TMT-SPAL protocol will not significantly distort the accuracy of the quantification relative to the Solution Phase protocol. Conclusion: By greatly reducing over-labelling, the TMT-SPAL protocol gives improved labelling compared to the previously published Solution Phase labelling protocols. TMT-SPAL is reproducible and robust and appears to give virtually complete labelling but with fewer side-reactions than the standard basic pH Solution Phase labelling protocol (Figure 3). There is a ~20% higher peptide


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identification rate, which appears to be due to a reduction in redundant sequencing of the same correctly labelled and over-labelled peptides. Indeed, over-labelling may be an overlooked mechanism contributing to the reported reduction in peptide identification rates with TMT when compared with label-free in like for like workflows20,21. This means instrument time is being spent identifying unwanted redundant over-labelled species rather than new species. An alternative factor that might contribute to improved peptide identification rates would be minimized losses during sample preparation, since the TMT-SPAL approach reduces the number of desalting steps required. Conversely, including searches for over-labelled species when using standard pH 8 solution phase labelling conditions may result in higher peptide identification rates. It is also worth emphasizing the importance of checking data for over-labelling when any kind of labelling procedure is used to ensure labelling reactions are proceeding as expected, particularly with newer high performance instrumentation, which is more capable of detecting low abundance species including products of side-reactions of labelling reagents. Our results also raise the question of whether over-labelling is an issue with other labelling steps in proteomics protocols such as the alkylation of cysteine with iodoacetamide, which is known to label amines and other reactive residues23. Similarly, carbamylation of amines by urea has been reported, with significant blocking on N-termini reported when high concentration urea is used as a denaturant24. Thus, other sources of chemical noise may also be worth investigating. There appears to be at least 80% overlap in the proteins identified by both protocols (Figure 6b) and we observe about 10% more protein identifications with the TMT-SPAL protocol overall (Figure 5 (right)). The overlap at the peptide level between the protocols is, however, lower


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(Figure 6a) indicating significant subsets of peptides that are identified uniquely by each method. Similar results are obtained when looking specifically at phosphopeptides (Figure 5). The TMT-SPAL protocol appears to show a slight systematic bias against shorter peptides and less hydrophobic peptides compared to the Solution Phase protocol and a bias in favour of slightly longer and more hydrophobic peptides (Supplementary Figures 5 and 6, 7a and 7b). Further analysis of this issue is warranted particularly in relation to the concentrations of organic solvents used, which may be causing losses of identifications particularly in the TMT-SPAL method. Organic solvent levels could undoubtedly be reduced in both methods, which would aid phosphopeptide recovery in particular. Interestingly, the different bias of the two labelling protocols also suggests that these methods could be used in parallel to get more peptide and phosphopeptide identifications overall than might be obtained by simply doing two replicate analyses with one labelling protocol. We observe very little difference in the number of unlabelled species between the protocols with high overall labelling efficiency in both protocols although care must be taken with the TMTSPAL to minimize hydrolysis due to the need to prepare the TMT reagent in aqueous buffer prior to loading of the C18 cartridge. The experiments presented here were, carried out as MS2 experiments rather than as MS3 experiments, which is known to give better quantitative accuracy25, thus we cannot categorically state that the accuracy of quantification is not changed by the TMT-SPAL protocol. Our data shows that the ion statistics for the MS2 data are, however, very similar for both protocols so we anticipate that there will be very little difference between the protocols in terms of accuracy of quantification. We will carry out more in depth analysis of this issue with MS3 experiments in due course25.


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Thus, we believe that the TMT-SPAL approach can be used as an alternative to Solution Phase labelling for complex protocols or for labelling large amounts of sample material giving the expected time saving and gain in peptide identifications. We are also investigating whether the low pH labelling works effectively as a solution phase protocol. However, since Trypsin digestion takes place at a slightly basic pH, changing the sample pH while retaining sensible sample volumes without using a desalting column is a non-trivial issue and so the TMT-SPAL process may be preferable to reduce the number of sample handling steps. This study also highlighted the need to take extreme care with solid phase sample processing steps to ensure that there is adequate binding capacity in the resins used. Usefully, the TMTSPAL protocol can clearly cope with quite large (up to 3mg) samples of peptide digest with suitable cartridges. The ability to label large samples efficiently has other useful applications, such as labelling large amounts of a sample to create ‘reference’ samples to spike into experiments to allow datasets in large-scale studies to be normalised for comparison. Similarly, with appropriate cartridges/tips, TMT-SPAL should also be suitable for labelling of very small samples, e.g. using C18 StageTips with FASP-like protocols26,27. The results presented here for labelling of peptides with N-hydroxysuccinimide ester TMTs on C18 should also be generally applicable to other active ester reagents that are moderately hydrophobic, such as biotin reagents and fluorophores and the reaction conditions developed here should be suitable for a wide range of applications beyond labelling of peptides with TMTs.

ASSOCIATED CONTENT Supporting Information.


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A Supplementary Material file is available with Supplementary Tables 1 to 5 and Supplementary Figures 1 to 10. This material is available free of charge via the Internet at http://pubs.acs.org. Raw data may be obtained from the EBI Pride Archive via the http://www.ebi.ac.uk/pride/archive/ with the PXD number PXD002072.

Internet

at

AUTHOR INFORMATION Corresponding Author Correspondence should be addressed to Andrew Thompson; Phone: +44 (0)1932 865065; Email: [email protected] Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Conflicts of Interest All authors were employees of Proteome Sciences plc at the time of writing. Funding Sources This work was funded by Proteome Sciences plc. ACKNOWLEDGMENTS We would like to acknowledge the kind assistance of Christopher Lößner for his support in identifying HSTY over-labelling and Claudia Höhle for technical support with Mass spectrometry analysis. ABBREVIATIONS


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ACN, Acetonitrile; C18, Octadecyl alkyl chain; EGFR, Epidermal Growth Factor Receptor; ER, Estrogen Receptor; FA, Formic Acid; FASP, Filter Aided Sample Preparation; FDR, False Discovery Rate; HCD, Higher energy Collisional Dissociation; HLB, Hydrophilic Lipophilic Balance; HPLC, High Performance Liquid Chromatography; HSTY, Histidine, Serine, Threonine and Tyrosine; IMAC, Immobilised Metal Affinity Chromatography; LC-MS/MS, KH2PO4,

Potassium

Dihydrogen

Phosphate;

Liquid

Chromatography

Tandem

Mass

Spectrometry; MCF7, Michigan Cancer Foundation 7; MS, Mass Spectrometry; MS2, Tandem Mass Spectrometry; MS3, 3 Stage Mass Spectrometry Analysis; MudPIT, Multi-dimensional Protein Identification Technology; NHS, N-HydroxySuccinimde; OT, OrbiTrap; PSM, Peptide Spectral Match; SCX, Strong Cation eXchange; SPAL, Solid Phase Amino Labelling; TEAB, Triethylammonium Bicarbonate; TFA, Trifluoroacetic Acid; TiO2, Titanium Dioxide; TMT, Tandem

Mass

Tag;

Tris,

tris(hydroxymethyl)aminomethane;

UHPLC,

Ultra-High

Performance/Pressure Liquid Chromatography. REFERENCES 1.

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Mohammed, S., Robust phosphoproteome enrichment using monodisperse microsphere-based immobilized titanium (IV) ion affinity chromatography. Nat Protoc 2013, 8, (3), 461-80.


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the femtomole level of phosphopeptides from proteolytic digests using 2D-NanoLC-ESI-MS/MS and titanium oxide precolumns. Anal Chem 2004, 76, (14), 3935-43. 5.

Villen, J.; Gygi, S. P., The SCX/IMAC enrichment approach for global phosphorylation

analysis by mass spectrometry. Nat Protoc 2008, 3, (10), 1630-8. 6.

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Hamon, C., Tandem mass tags: a novel quantification strategy for comparative analysis of complex protein mixtures by MS/MS. Anal Chem 2003, 75, (8), 1895-904. 7.

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S. P., Evaluating multiplexed quantitative phosphopeptide analysis on a hybrid quadrupole mass filter/linear ion trap/orbitrap mass spectrometer. Anal Chem 2015, 87, (2), 1241-9. 10. Britton, D.; Zen, Y.; Quaglia, A.; Selzer, S.; Mitra, V.; Lobetaner, C.; Jung, S.; Bohm, G.; Schmid, P.; Prefot, P.; Hoehle, C.; Koncarevic, S.; Gee, J.; Nicholson, R.; Ward, M.; Castellano, L.; Stebbing, J.; Zucht, H. D.; Sarker, D.; Heaton, N.; Pike, I., Quantification of


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Abstract Graphic: Bar Chart showing the total numbers of PSMs identified in our TMT-SPAL data (top) and in our Solution Phase data (bottom). The number of correctly labelled PSMs (dark blue) is considerably increased in the TMT-SPAL sample. Relative proportions of PSMs that were improperly labelled with labelling at histidine (red), labelling at serine (green), labelling at threonine (purple) labelling at tyrosine (light blue) are shown. 82x50mm (300 x 300 DPI)


Low-pH Solid-Phase Amino Labeling of Complex Peptide Digests with

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