An Optimized Platform for Hydrophilic Interaction Chromatography

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An Optimized Platform for HILIC-IMAC Enables Deep Coverage of the Rat Liver Phosphoproteome Francesca Zappacosta, Gilbert F Scott, Michael J Huddleston, and Roland Annan J. Proteome Res., Just Accepted Manuscript • DOI: 10.1021/pr501025e • Publication Date (Web): 09 Jan 2015 Downloaded from http://pubs.acs.org on January 13, 2015

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Journal of Proteome Research is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

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Journal of Proteome Research

An Optimized Platform for HILIC-IMAC Enables Deep Coverage of the Rat Liver Phosphoproteome

Francesca Zappacosta, Gilbert F. Scott, Michael J. Huddleston and Roland S. Annan*

Proteomics and Biological Mass Spectrometry Laboratory, GlaxoSmithKline, Collegeville PA 19426

KEYWORDS: phosphoproteome, rat liver, HILIC, IMAC, FASP

ACS Paragon Plus Environment

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Abstract: While analysis of the phosphoproteome has become an important component of understanding how cells function, it remains a non-trivial task in terms of the number of sample preparation steps and instrument time needed to achieve sufficient depth of coverage to produce meaningful results. We previously described a multi-dimensional method that uses hydrophilic interaction chromatography (HILIC) followed by Fe3+ IMAC to reduce complexity, improve selectivity and increase phosphopeptide identifications. Here we present refinements to our overall protocol that make it simpler and more efficient while providing greater coverage of the phosphoproteome. We introduce Filter-Aided Sample Prep (FASP) for cell lysis and trypsin digestion. Following HILIC separation, fractions are IMAC enriched using a 96-well filter plate. Finally, enriched samples are analyzed using an LC-MS strategy optimized for the fractionation scheme. The optimized protocol improves protein recovery, simplifies phosphopeptide enrichment, and optimizes instrument time while maintaining deep coverage of the phosphoproteome. Using the refined protocol, we identified more than 16,000 unique phosphosites from rat liver, in a single experiment using approximately one day of instrument time. All together, we present evidence for 24,485 rat liver phosphosites, representing the deepest coverage of a tissue phosphoproteome to date.


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Introduction

Much of the activity in the cellular proteome is under the control of reversible protein phosphorylation. Phosphorylation-dependent signaling regulates differentiation of cells, triggers progression of the cell cycle, controls metabolism, transcription, apoptosis and cytoskeletal rearrangements. Signaling via reversible protein phosphorylation also plays a critical role in intracellular communication and immune response. Phosphorylation can function as a positive or negative switch, activating or inactivating enzymes. It can serve as a recognition element for the addition of other posttranslational modifications, or serve as a docking site to recruit other proteins into multi-protein complexes. Phosphorylation can trigger a change in the three dimensional structure of a protein or initiate translocation of the protein to another compartment of the cell1-5. Disruption of normal cellular phosphorylation events is responsible for a large number of human diseases6-8. Given the wide range of processes that are under the control of reversible protein phosphorylation, it is not surprising that the extent of phosphorylation in higher order organisms is massive. Current phosphosite databases9 list more than 150,000 sites on over 18,000 human proteins, many more than were previously predicted10. The large majority of these sites have been identified in high throughput phosphoproteomics studies utilizing mass spectrometry. Coupled with two dimensions of peptide separation and a phosphopeptide enrichment strategy, it is now common for mass spectrometry to identify 5000-10,000 unique phosphosites from a single sample and 20,000-30,000 unique sites in a complete experiment11-20. This depth of coverage for the phosphoproteome does not come without a cost. The price to pay is measured in instrument time. While the law of diminishing returns certainly applies, generally


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speaking analyzing more samples, taking more fractions, and running longer LC-MS gradients will increase the number of phosphosites identified. However, this strategy has its limitations when considering a properly powered quantitative experiment. In a study that might investigate more than one treatment, over multiple time points or doses and uses a meaningful number of biological replicates, instrument time becomes a very real consideration when designing the experiment. Multiplexing samples via stable isotope encoded labels21-22 has eased this tension somewhat; however, the demand for proteomic studies continues to increase and the cost of the high resolution instruments needed to perform these studies is high. Thus the importance of maximizing proteome coverage while minimizing instrument time is paramount. In 2008 we introduced a phosphoproteome protocol23 that utilized hydrophilic interaction chromatography (HILIC) as a first dimension fractionation scheme in front IMAC enrichment. This combination resulted in fractions going to the final data-dependent LC-MS/MS step that were better than 95% phosphopeptides in most cases. This of course maximizes the use of the instrument, in that the mass spectrometer does not spend time sequencing irrelevant nonphosphorylated peptides. Since the publication of the original HILIC/IMAC protocol there have been advances in sample processing and instrument capability. Here we describe a revised HILIC/IMAC protocol for phosphoproteomics. Changes have been made to improve the recovery of proteins from cells and tissues, simplify the enrichment procedure, minimize the loss of phosphopeptides during the various steps and maximize the use of instrument time. We have incorporated the Filter Aided Sample Preparation (FASP)24method to improve the solubilization of proteins from cells and tissues. Sample handling, throughput and reproducibility of the IMAC protocol was improved by replacing the standard pipette tip protocol with a commercially available 96 well filter plate. To


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make maximum use of instrument time, we evaluated various combinations of HILIC fractionation and reverse phase LC gradients to achieve the largest number of phosphosite identifications in an approximately 24 hr period. Combining the newly optimized steps into a revised protocol we investigated the phosphoproteome of rat liver. Rodents are important model organisms for the study of human disease and safety. Rats are in many ways better models than mice for human biology25. The rat is the primary model for mechanistic studies of human reproduction and the primary model for human toxicology. In spite of the importance of rodents to the in vivo study of basic biological function, only recently have the phosphoproteome of various rat and mouse tissues been studied in detail13-18, with most of the work being done in the mouse. Here we report the deepest phosphoproteome coverage of any rodent tissue to date. More than 24,000 unique phosphosites were identified in rat liver with more than 16,000 identified in a single sample. This represents almost three times the number previously reported for the rat liver.

Materials and Methods Tissue Preparation All studies were conducted in accordance with the GSK Policy on the Care, Welfare and Treatment of Laboratory Animals and were reviewed by the Institutional Animal Care and Use Committee at GSK. Livers were surgically harvested from male F344/NTac rats and immediately homogenized in 4% SDS, 100 mM Tris-HCl (pH 7.6), 100 mM DTT lysis buffer (ca. 5 l buffer per mg of tissue) for a few minutes at room temperature using a blender. The homogenates were incubated for 3 min at 95°C, sonicated for 4–5 min, centrifuged at 16,000g at


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20°C for 5 min and supernatant was collected. Protein concentration in the supernatant was determined using the Pierce 660 nm Protein assay with IDCR (Pierce). FASP Digestion Aliquots of rat liver lysate (30 mg/mL) were FASP digested in 30 kDa Microcon centrifugal filters (Millipore) essentially following the protocol developed by Wisniewski and Mann19. For amounts greater than 1mg the sample was processed in multiple filters and combined after digestion. For each filter, the rat liver lysate was diluted with 200 uL of 8 M Urea, 100 mM Tris, pH 7.8. For the final set of experiments to profile the rat liver phosphoproteome, 5 mg of rat liver lysate was divided over six filters for FASP. After buffer exchange the samples were alkylated with iodoacetamide and washed twice with urea buffer. The urea buffer was replaced with 50 mM ammonium bicarbonate buffer, pH 8.5 and trypsin (Promega) was added in a ratio of 1/100 enzyme to protein. Proteins were digested in the filters at 37°C for 16 hours. Peptides were collected by centrifuging two separate 100 uL volumes of 50 mM ammonium bicarbonate through the filters. FASP-digested samples were desalted using a 6cc Sep-PAK column (Waters). Peptides were eluted in 80% ACN, 0.1% TFA. The volume was reduced to 200 uL using a SpeedVac and brought back up to 2 mL with ACN prior to injecting onto the HILIC column. Alternatively, the FASP-digested samples in ammonium bicarbonate buffer were reduced in volume and diluted to 2 mL with 98% ACN/0.1%TFA and directly injected onto the HILIC column. Peptide Fractionation by HILIC HILIC separations were performed on a 4.6 x 25 cm TSK gel Amide-80 column, 100 Å pore size, 5um particle size (TosoHaas) using ACN:TFA solvents running at 0.5 mL/min on a Beckman HPLC with a 2 mL sample loop. Buffer A was composed of 2% ACN, 0.1% TFA and


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buffer B 98% ACN, 0.1% TFA. After a 10 minute post injection hold at 80% B, peptides were eluted with a gradient going from 80% to 65% B in 35min followed by 65% to 0% B in 5 min and a 10 min hold to end for a total run time of 60 minutes. Fractions were collected every minute and stored at 4°C until use for phosphopeptide enrichment.

Phosphopeptide Enrichment using IMAC HILIC fractions from 16-56 min were pooled into 28 fractions and 20 ul of PHOS-Select- Iron Affinity Gel (Sigma) was added directly to the fractions. No preparation of the resin was required. Phosphopeptides were coupled to the beads for 30 minutes at room temperature by vortexing. Gel loader tips (Eppendorf) were built by inserting a Procise TFA-treated cartridge filter (Life Solutions). Solutions were pushed through using a 3 ml plastic syringe. After incubation with the beads, fractions were spun down at 3000 rpm for 2 min and the supernatant was discarded. Beads were washed twice with 250 mM acetic acid/30% ACN and once with water. Beads were at this point transferred into the gel loader tip and phosphopeptides were eluted with two sequential 20 ul additions of 400 mM NH4OH/30% ACN (pH 10). Samples were concentrated in a SpeedVac and made acidic for mass spectrometric analysis by the addition of 1/10 volume of 1% formic acid/ 0.5% TFA. AcroPrep Advances 96-well Filter Plates (0.45 um PTFE, PALL Corporation, PN# 8048) were used with a vacuum manifold (Waters). Prior to loading samples, plates were wetted with 200 ul ethanol per well, washed twice with 200 ul of 400 mM NH4OH/85% ACN and twice with 200 ul of 250 mM acetic acid/95% ACN. After incubation with the beads, HILIC fractions were spun down at 3000 rpm for 2 min and part of the supernatant was discarded leaving about 200 ul


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of volume containing the beads. Bead-containing solutions were transferred into the plate and vacuum was applied to get rid of the solution. Beads were washed twice with 200 ul of 250 mM acetic acid/30% ACN and once with 200 ul of water. Phosphopeptides were eluted with two sequential additions of 100 ul and 50 ul each of 400mM NH4OH/30% ACN (pH 10). Samples were collected into a 96-well plate, transferred into injection vials, concentrated in a SpeedVac and made acidic by the addition of 1/10 volume of 1% formic acid/0.5% TFA ready for mass spectrometric analysis.

Mass Spectrometric Analysis LC-MS/MS analysis was performed on a LTQ-Orbitrap Velos Pro (Thermo Scientific) mass spectrometer equipped with an Agilent 1100 Nano HPLC system and a nanospray source. Phosphopeptide fractions containing 0.1% formic acid and 0.05% TFA were loaded onto a 200µm x 5mm trap column with 0.05% HFBA at 10µl/min. After 5 min the trap is put in line with a 100µm x 5cm analytical column using mobile phases A: 0.2% formic acid in water and B: 0.2% formic acid in ACN at a flow rate of 0.5µl/min. Both trap and analytical columns are monolithic PSDVB polymer type columns (Dionex), operated at 40oC. Our default gradient program starts at 2%B and goes to 15% B in 60 min then 15% to 30% B in 30 min for a total working gradient time of 90 min. Other gradients explored had a total time of 30, 45, 60 and 120 min keeping the same two-segment gradient slope. An additional 30 minutes, that includes sample loading (5 min), end ramp from 30% B to 95% B (5 min), end hold at 95% B (5 min) and column re-equilibration at 2% B (15 min) is added to each gradient program to give the total run time. The analytical column is interfaced via high voltage liquid contact made thru a SS union to a 50µm fused silica emitter (New Objective) pulled to 15µm at the tip. MS data was acquired


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from m/z 400 to 2000 at a resolution of 30,000. Data dependent MS/MS acquisition was set to trigger up to 20 ion trap CID spectra per MS scan using a precursor isolation window of 1.5 m/z and normalized collision energy of 35. The MS/MS AGC target value was set at 1E4 with a maximum injection time of 100ms. For all gradient lengths, the dynamic exclusion was set to 30s. Data Analysis The raw MS files from the LTQ-Orbitrap were processed with MaxQuant26, version 1.5.0.25. Proteins and peptides were identified against the UniProtKB rat reference proteome database (April 2014) using the Andromeda search engine27. The following search parameters were used: mass deviation of 6 ppm on the precursor and 0.5 Da on the fragment ions; tryp/P for enzyme specificity; two missed cleavages. Carbamidomethylation on cysteine was set as a fixed modification. Oxidation on methionine, phosphorylation on serine, threonine and tyrosine and acetylation at the protein N-terminus were set as variable modifications. Thresholds for the identification of phosphopeptides were Delta Score = 6 and Andromeda score = 40. The false discovery rate was set to 1% for positive identification of proteins, peptides and phosphorylation sites. Class I phosphorylation sites are the phosphorylation sites where the phosphorylation site could be assigned with a probability of at least 0.75. Data analysis was performed in Excel. The hydropathicity and pI for phosphopeptides were calculated using tools found at http://www.gravy-calculator.de and http://proteomics.mcw.edu/tools bin/promost/promost.cgi, respectively.


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Results and Discussion Optimization of the FASP/HILIC protocol for phosphoproteomics We have previously described a phosphoproteomics protocol that uses HILIC for the enrichment and fractionation of phosphopeptides23. Revisiting this protocol recently, we set about improving some of the component methods to make the entire protocol easier and more efficient. Our first aim was to implement the FASP protocol introduced by Wisniewski and Mann24 and coupled it to the HILIC separation. We have determined several factors which appear to be crucial for the success and reproducibility of the FASP/HILIC combination. In particular, we had previously observed a lack of reproducibility in the recovery of FASP-generated peptides (as indicated by UV absorbance at 214nm). This problem became especially evident when attempting to scale up the amount of lysate to be digested in the filters for phosphoproteomics. Since the concentration of the lysate is generally fixed according to the initial lysis volume, scaling up the amount of protein for the FASP protocol means adding a greater volume of the 4% SDS lysate, which of course corresponds to adding an ever greater amount of SDS to the filter. This situation can occur when the experiment calls for large amounts of protein as is the case for many global posttranslational modification studies or when the initial lysate concentration is quite low as is often the case with difficult to access primary cells. In both cases, even though the lysis buffer is 4% SDS, the total amount of SDS added to the filters increases as the volume of the lysate added increases. To investigate whether this was in fact the source of our recovery problems, we lysed rat liver in 4% SDS and prepared nine samples with varying protein concentration in 4% SDS (see Figure 1A). Five hundred micrograms from each sample was applied to the Microcon filter and digested as per the protocol. As the protein concentration decreased, a larger volume of lysate needed to be added to the filter to make up the 500 ug. To


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balance the final volumes, a variable amount of Tris buffer was added to each sample (Figure 1A). After elution of the peptides from the FASP filter, the results were assessed by the 214nm UV response during HILIC chromatography. While this approach doesn’t provide a measure of absolute recovery, it is a convenient way to measure the relative recovery from the various conditions tested. Figure 1B shows the peptide recovery versus the amount of SDS added to the filters. A clear reduction in peptide recovery is evident as the amount of SDS added to the filter increases with the volume of lysis buffer needed to yield 500 ug of sample. Applying 200 uL of the 4% SDS lysate yields less than 5% of the amount recovered when compared to the lowest volume of SDS tested. It should be noted that at the lowest amount of SDS used in this experiment, the peptide recovery had not yet reached a plateau. We reasoned that the variable peptide recovery results from impaired trypsin digestion caused by residual SDS trapped in the filters. The critical micelle concentration for SDS in water is 0.0082M or 0.2%. Since the initial lysis solution is more than twenty fold above the CMC, unless the solution is sufficiently diluted, SDS micelles will be trapped in the filter. Subsequent urea washes will remove much of the SDS, however trypsin activity is reduced at SDS levels as low as 0.005%28. To test this, an aliquot of the material retained on the filter from the experiment described above was analyzed by SDS-PAGE (Fig 1C). The presence of large amounts of high molecular weight species suggests that residual SDS has inhibited the activity of trypsin, leaving behind undigested or partially digested proteins. Thus, it is clear from these data that the amount of SDS added to the filters should be kept as low as possible. In cases where the starting lysate is very dilute or whenever more material is required for the experiment, the lysate should be divided over several filters.


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We next investigated whether it was necessary to desalt the peptides produced by the FASP procedure prior to loading the HILIC column. The FASP eluted peptides are in an ammonium bicarbonate solution, so it should be possible to load them directly onto the HILIC column after acidification and the addition of ACN to achieve the initial HILIC loading conditions. To test the importance of desalting, we split FASP digested samples into two, each containing 2.5mg of lysate. One was desalted prior to HILIC, while the other was analyzed directly. Peptide recovery was monitored by the UV response at 214 nm during HILIC chromatography. Results from three experiments show that desalting it is not necessary prior to HILIC separation as the UV response was quite similar between the two samples (Figure S1; Supporting Information). We did find, however, that it is important to centrifuge samples at high speed prior to the HILIC column. Particularly on smaller diameter columns (1mm) we found that without prior centrifugation, column performance declined rapidly after several injections. Depending on the elution volume from the FASP filter(s) and the size of the loop on the injector, it may be necessary to reduce the volume prior to loading the sample onto the HILIC system. Previously, we had dried the eluates and resuspended the samples in the HILIC loading solution (98% ACN/0.1% TFA). While this did not seem to have an impact on our peptide-based expression proteomic studies, we observed a quite variable and sometimes severe loss of phosphopeptides when the samples were dried down. Losses are most severe for the later eluting, more hydrophilic peptides (black trace, Figure S2; Supporting Information). Losses in this area of the chromatogram would directly affect the recovery of phosphopeptides as these elute primarily in the latter part of the gradient. It is likely that these losses are due to decreased solubility of the more hydrophilic peptides in the high organic re-suspension buffer. Concentration of the peptide solution from the filters to one tenth of the injection volume and


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dilution of the sample to the final injection volume by the addition of nine volumes of ACN resulted in a better and more reproducible peptide recovery (red trace, Figure S2; Supporting Information).

Optimization of phosphopeptide IMAC enrichment using a 96-well PTFE filter plate We have previously demonstrated that performing phosphopeptide enrichment after peptide fractionation greatly improves the selectivity of the enrichment and produces a larger number of phosphopeptides identification, compared to a protocol in which single phosphopeptide enrichment is performed prior to fractionation23. While performing individual enrichments on multiple fractions has led to a dramatic improvement in phosphoproteome coverage, this strategy is quite labor-intensive. We and others have made use of 0.22 micron nylon spin filters to simplify the enrichment protocol. Unfortunately these filters are subject to contaminants leaching from the nylon that severely degrades the mass spectrometers performance (data not shown). Phosphopeptide enrichment in a gel-loader pipette tips29 have been widely and successfully adopted14, but can be prone to irreproducibility due to the many small volume manipulations involved. To improve reproducibility and reduce labor, time and sample handling of the enrichment protocol when multiple fractions or samples are being processed, we tested the performance of a 96-well PTFE filter plate (0.45 nm) for IMAC phosphopeptide enrichment. All phosphopeptide enrichments were carried out using 20 uL of Sigma Phos-Select resin. In our hands, washing and equilibrating the resin in the supplied IMAC binding buffer did not improve the performance of the resin, so all experiments were performed using the resin as supplied and adding it directly to the HILIC fractions. After 30 minutes of incubation, the beads were added to the filter plate


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which had been pre-wetted with ethanol. Phosphopeptides were eluted with a solution of 400 mM NH4OH. Preliminary results showed a very high efficiency for phosphopeptide enrichment (>90%) but a very poor recovery in terms of total number of phosphopeptides. Reasoning that a hydrophobic interaction with the filter was preventing the complete elution of the phosphopeptides, we added 30% ACN to the elution buffer. While this greatly improved the recovery of the phosphopeptides, an abundant contaminant was released from the filter, which caused a slow deterioration of the LC column performance over time. We found that washing the filter plate first in a basic solution containing high organic and then in an acidic solution containing high organic prior to the addition of the resin completely removed the contaminant, without affecting the overall phosphopeptide enrichment or the yield. We evaluated the performance of the 96-well plate IMAC enrichment protocol against our default protocol which uses gel-loader tips. Select HILIC fractions from three different rat liver preps were split in half, with one half of each sample being enriched using a gel-loader tip and the other half enriched using the 96-well plate. In each case, the filter plate performed as good or better than the gel loader tip in terms of efficiency of phosphopeptide enrichment (Table 1). The total number of phosphopeptides identified in each of the three experiments using the 96-well filter plate was at least 25% greater than the number of phosphopeptide identified using the gelloader tip. The protocol for the plate based IMAC enrichment is shown schematically in Figure 2. Performing the IMAC enrichment in the 96-well filter plate proved to be very efficient and labor-friendly, allowing the multiplexing of multiple fractions from multiple samples with minimal extra labor. After loading the samples, the entire plate can be processed in under 30 minutes. If less than the full 96 wells are needed for an experiment, the unused wells are simply


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covered and used at another time. While we routinely enrich 28 fractions per sample, these can be combined prior to MS analysis, depending on the depth of phosphoproteome coverage required for the study and the time constraints for MS analysis.

The improvements in sample

preparation yielded more peptides and proteins, efficient phosphopeptide enrichment, and less sample handling during the more sensitive steps of the protocol. Optimization of LC-MS/MS gradient length and fraction number for maximal phosphoproteome coverage The challenge for any proteomic study is to achieve deep coverage in a realistic amount of instrument running time. For our laboratory it seemed reasonable to commit approximately 24 hours of instrument time per phosphoproteome sample. This would allow us to complete a study with multiple time points, a dose curve or multiple biological replicates, in one week. Thus we set out to determine the optimal number of IMAC enriched HILIC fractions and the LC-MS/MS gradient length that would produce the largest number of phosphopeptides in our target time. To this end, five milligrams of FASP-digested rat liver lysate was separated on a 4.6 mm HILIC column. Forty of the collected one minute fractions were pooled into 28 fractions according to the diagram in Figure 3A. The pooling strategy we used was optimized following exploratory experiments to assess the elution profile of phosphopeptides in a typical lysate (data not shown). HILIC on the TSK gel Amide-80 columns using ACN:WATER/TFA gradients30 is a high resolution technique for the separation of peptides (Figure S3, Supplemental Information) with high recoveries30 and a high degree of orthogonality to reverse phase chromatography31-32. The HILIC gradient described here is optimized for phosphopeptide enrichment, such that the bulk of the unphosphorylated peptides are eluted early in the gradient while the phosphorylated peptides elute later in the second half of the gradient23. The 28 fractions were then IMAC enriched using


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the 96-well PTFE filter plate and further pooled, pair-wise into 14 fractions (Figure 3A). One tenth of each enriched fraction was analyzed by LC-MS/MS using three different gradients lengths. Each of the three gradients uses the same two-segment slope but was developed over 60, 90 or 120 minutes. A 30 minute wash and equilibration cycle at the end of each gradient contributes to the total analysis time. The results from these analyses are shown in Figure 3B. The 60 minute gradient identified 13,630 unique phosphosites in 21 hours of instrument time. It has been our practice and indeed it is a common practice in the proteomics community to analyze a technical replicate with the aim of increasing the depth of coverage; thus we repeated the 60 minute gradient and combined the two data sets. The replicate injection increased the total number of sites by nearly 15% (15,642 sites), but at a cost of double the total analysis time to 42 hours (Figure 3B). Lengthening the gradient to 90 minutes and performing a single analysis of the 14 fractions increased the number of phosphosites by 5% (16,421) over our default strategy (2x60 min ) and decreased the overall time of analysis by 30% (28 vs 42 hours). This result is consistent with Kocher33 and our overall experience with nonphosphorylated peptides, that a single analysis with an optimized longer gradient always yields more proteins in less time than two analyses with a shorter gradient. Lengthening the gradient time further to 120 min would push the total time for a single analysis (35 hours) past our target, though still less than the default 2x60 strategy. Surprisingly, the 120 min gradient yielded 12% fewer phosphosites compared to the 90 min gradient. The performance specifications for the monolithic columns used in these studies have been well documented in the literature34-36. Very high efficiency separation of complex mixtures is possible on columns as short as 5cm (Figure S4, Supporting Information). However, the peak capacity for these columns flattens out sharply with gradients of approximately 60-90 minutes, while the peak width continues to increase sharply34. This


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would be consistent with our observation of a decrease in phosphopeptide identifications at 120 minutes relative to 90 minutes due to the lack of increased peak capacity and the corresponding linear broadening of the peaks. It should be noted that LC-MS system performance was monitored throughout the experiment by analyzing a commercially available E. coli digest under a standard set of gradient and MS conditions and monitoring the number of peptides identified over the course of the experiment. We found that the number of E. coli peptides identified decreased less than 3% over the entire duration of the experiment (126 hr) suggesting that there was no significant decline in the system performance. We also evaluated the peak area counts for a group of randomly selected phosphopeptides over all four of the gradient sets and found that there was very little change, again suggesting no deterioration of instrument performance or degradation of the samples in the 4oC autosampler over the course of the 126 hr experiment. The number of phosphosites identified in the second 60 minute set which was started 55 hrs after the first 60 minute set finished, showed a decrease of approximately 5%. Having found that for our chromatography system, 14 IMAC enriched HILIC fractions (pooled as shown in Figure 3A) analyzed using a single 90 minute gradient each gave us the largest number of phosphosites in an analysis time of a little over 24 hours, we wondered whether we could improve on this by adjusting the number of fractions analyzed. We took an aliquot from each of the 28 IMAC enriched samples described above and pooled them to give a set of 14 and a set of 7 fractions. This gave us three identical samples divided into pools of 28, 14 and 7 fractions. To stay at or under our target total time goal, fraction sets were analyzed using the following fraction/gradient length combinations: 28 fractions/30min gradient; 14 fractions/90min gradient; 7 fractions/120min gradient. Gradient programs have the same two-


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segment slope but different development time. A 30 minute wash and equilibration cycle between each run adds to the total analysis time. To normalize even small effects from a decline in instrument performance or a loss of phosphopeptides in the autosampler, we analyzed the fractions in small batches, running all three combinations before moving on to the next batch. The results are shown in Figure 3C. Analyzing 14 fractions using the 90min gradient (28 hours MS time) we identified 12,774 phosphosites. In order to analyze more fractions, but stay within our target total analysis time goal we had to shorten the gradient considerably. The choice of gradient length for the 28 fraction set is limited by the between run re-equilibration time. At 30 minutes per sample, the reequilibration time contributes 50% of the total target run time for 28 fractions, limiting us to not longer than a 30 minutes gradient if we are to stay in our approximately 24 hr goal. A 28 fraction/30 min gradient combination gave us the same total analysis time as the 14 fraction/90min gradient combination (28 hr). This combination produced 15% fewer phosphosites than the 14 fraction/90 min scheme, suggesting that the fractions were still too complex for a 30 min gradient. With the sample divided into only 7 fractions we reasoned that each fraction would be sufficiently complex to benefit from a longer gradient in spite of the flattening peak capacity and concomitant peak broadening which accompanies the longer gradients on these columns. Using a 120 min gradient we were able to analyze the 7 fraction set in 17 hr, well under our target goal. However, this combination still produced fewer phosphosites relative to the 14 fraction/90 min combination. The outcome of these experiments show that for a target total run time of around 24 hr the greatest number of phosphosites identified on our system is achieved using 14 IMAC enriched


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HILIC fractions and 90 min gradients. In order to compare the performance of the various combinations of fractions and gradient lengths, we set the results from each of the 14/90 combinations to 100% and calculated the performance of the other combinations relative to them. As can be seen in Figure 3D, running the 14/60 combination twice produces the next best results, but takes twice almost twice as long. In terms of efficiency, the 7/120 combination is a good choice in that it generates 88% of the sites in 40% less time. It is important to note that this specific combination of fractions and gradient length may or may not be the optimal combination for the reader’s chromatography system. However, it certainly serves as a starting point for an evaluation and underscores the point that a chromatography platform needs to be independently evaluated to achieve the optimal configuration. Recently a very large study has been published where the phosphoproteome of fourteen different rat tissues was cataloged14. In this study sequential enrichments of unfractionated lysates were analyzed using 180 min gradients. The phosphoproteome for any individual tissue was collected in less than half a day. While this is clearly a highly efficient strategy for analyzing a large number of tissues, it yielded on average less than 50% of the number of sites identified in our optimized platform. Likewise on our system, 7 fractions and a 120 min gradient was more efficient, but yielded 10% less phosphosites. For the time being there will continue to be a tradeoff between efficiency and comprehensiveness and each researcher must, therefore, strike the balance that suits their goals.

Rat Liver Phosphoproteome We have prepared aliquots of five mg of rat liver for the above described experiments but only 1/10 of each fraction was used for MS analysis (corresponding to a 500 ug sample). Among the


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experimental conditions tested, we determined that the best results were achieved by pooling the IMAC enriched fractions into 14 samples and analyzing each sample using a 90 min gradient. Under these conditions, total MS analysis was 28 hours and led to the identification of 20,089 unique phosphopeptides (fig 4A). These corresponded to 16,421 unique phosphosites (fig 4B) of which 10,519 have a calculated localization probability >75% (class I). The number of phosphopeptides detected per fraction were as low as 604 in the earliest fractions (combination of fractions 16-24) and as high as 2773 in fractions 35-36 (fig 4A). As described in our previous paper, the majority of the phosphopeptides elute in the HILIC separation in the second half of the gradient, resulting in a partial enrichment of the phosphopeptides (fig 4B). Even more so, due to the increased hydrophilic nature of multiply phosphorylated peptides, these were found to elute in the later part of the gradient (fig. 4A). Better than 85% of all phosphopeptides were only detected in a single fraction, and on average each fraction was approximately 81% unique (Figure 4C). Efficiency of the IMAC enrichment was lower in the earlier fractions, due to the overwhelming amount of non phosphorylated peptides in these fractions (Figure S4, Supplemental Information). The lower enrichment efficiency and smaller number of sites identified in the earliest fractions suggests that this is one area for continued improvement. The data produced in this experiment compares quite favorably with other published phosphoproteomic studies in terms of the amount of starting material used, total analysis time, and number of sites identified (Table S1). While the rat is an understudied organism in terms of proteomics, the mouse has been well studied11, 16, 37. In a recent analysis of the liver from insulin treated mice, the authors identified 16,339 sites in a single biological replicate, from 10mg of tissue, in approximately 20 hours of total instrument time11. Immortalized cancer cell lines have


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been particularly well studied. Subjecting an equivalent amount of HeLa cells to phosphoproteome analysis, Zhou et.al identified 13,681 phosphosites in 48 hours38. During the course of these experiments, we prepared and analyzed three aliquots of rat liver using the 14 fraction/90 minute combination protocol (Figure 5A). This represents a typical phosphoproteomics experiment with three replicates, took 3.5 days and led to the identification of 28,157 unique phosphopeptides and 20,953 phosphosites, of which 13,737 were class I. The percentage of phosphoserine (86.2% of total), phosphothreonine (11%) and phosphotyrosine (2.6%) is in good agreement with those reported in previous large scale phosphoproteomic studies13-14. Multiply phosphorylated peptides make up ca. 24% of the total (Figure 5A). Interestingly, the distribution of intensities for multiply phosphorylated peptides is identical to the distribution of peptides which contain only a single phosphorylation site (Figure 5B). One might have expected the distribution for the multiply phosphorylated peptides to be shifted overall to lower intensities, given the common perception that these peptides ionize less well. The identified phosphopeptides map onto 4622 proteins from the UniProtKB rat reference database. Approximately 30% of these proteins have a single identified site, but nearly 20% contained between 10 and 149 sites, suggesting that hyper-phosphorylation is quite common in the proteome (Figure 5C). Reproducibility between the three replicates was remarkably good. Eighty percent of the sites identified in the second and third replicates were contained in replicate 1 and the reproducibility between the second two was 74% (Figure 5D). By running three replicates we were able to produce at least duplicate measurements on nearly 13,000 phosphosites (all yellow area, Figure 5D bottom). During the course of this evaluation, we produced five additional rat liver phosphoproteomes from the various chromatographic evaluations. Combining all eight data sets gave us a rat liver


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phosphoproteome containing 24,485 unique potential sites and 16,567 class I sites. A complete list of the sites and peptide evidence for each site is provided in Supplemental Table S2 and S3, Supporting Information. The additional sets added only 3500 new phosphorylation sites. However, since each of the additional sets contained 20-50% fewer phosphorylation sites than the core 14 fraction/90 minute phosphoproteome, it is not surprising that most of the sites from the additional sets are contained in the core set. The liver, based on the data published in both the rat14 and mouse13 tissue specific catalogs of protein phosphorylation is not one of the more abundant tissues with respect to phosphorylation. This likely reflects the fact that liver is also one of the least complex tissues in terms of the percent genome expressed39. The UniProtKB rat reference database contains 26,982 sequences (April 2014 release) while the mouse reference database contains 42,433 sequences. Reasoning that there might be significant advantage to be gained by including the mouse sequences, we searched the total rat phosphoproteome data set against a combined rat-mouse database. Surprisingly the total number of phosphosites identified in the two searches was within 0.04% of each other, suggesting there is no significant advantage to including the mouse sequences. A tissue catalog of rat phosphoproteins was recently published which contained more than 31,000 phosphosites identified across 14 rat organs and tissues14. The authors developed a highly efficient protocol that allowed them to analyze a single, unfractionated phosphopeptide enrichment using one three hour LC-MS gradient. By performing triplicate analyses, they were able to identify on average more than 8000 phosphosites per tissue in less than half a day. Collectively and per tissue this study contained the largest number of phosphosites reported to date for the rat. A similar study, containing similar numbers of phosphosites in total and per tissue has been reported for nine mouse tissues13.


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Since the rat phosphosite atlas in Lundby et al.14 contained the largest number of rat liver sites published to date, we wondered about the overlap between this set and ours and how comprehensive the combined data set might be. To do this accurately, we analyzed both data sets in a single Maxquant experiment, thus unifying the protein identifiers and filtering both sets of data to common thresholds. Based on false discovery rate data from a very large phosphopeptide library study published recently40, the most current versions of the MaxQuant software set more stringent default filters for phosphopeptide identifications than previous versions of the software. As such, the number sites identified in the liver from the Lundby data set was somewhat reduced. Nevertheless, at the phosphosite level, 84% of the sites from the Lundby liver analysis were contained in our data set (Figure 6A), with 90% of these being class I in one data set or the other. Over 3000 additional phosphosites in our liver data matched sites identified in other tissues in the rat phosphosite atlas14. At the protein level 84% of the liver phosphoproteins in Lundby were contained in our data set (Figure 6A). Taken together, these two data sets provide evidence for phosphorylation of 4858 rat liver genes. Recent evidence at the protein level has verified 8185 genes expressed in the liver41. Thus the data here suggests that at least approximately 60% of the expressed rat liver genome is phosphorylated. Perhaps the overlap in the liver sites is not too surprising considering that ours was a much larger data set. However, given that the Lundby et al. data were collected using an entirely different platform, namely TiO2 enrichment and HCD compared to our use of Fe3+ IMAC and CID, it is remarkable that only 16% (773) of the total phosphosites from Lundby are unique to the TiO2 enrichment. It has been suggested that there are substantial differences in the selectivity between the IMAC and TiO2 resins42. A recent report by Matheron et al.43 using both a large (23,000) synthetic phosphopeptide data set and a somewhat smaller (5000) cell lysate peptide


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data set, found no obvious difference in the selectivity between Ti4+ IMAC and TiO2 . Since we used the more common Fe3+ IMAC in this study, it is worth looking for potential differences in the peptides unique to our data set and the Lundby TiO2 data. We compared peptide length, hydropathicity, pI and phosphosites per peptide for the unique peptides in each set and found that the peptides unique to TiO2 were somewhat shorter, slightly more hydrophilic, and more acidic and had a slightly higher degree of multiple sites compared to the peptides unique to Fe3+ IMAC (Figure 6B). By and large, however, the differences are fairly small and more than likely reflect the fact that the TiO2 peptides were enriched as an unfractionated mixture while the Fe3+ IMAC peptides were enriched from fractions. The type of peptides unique to the TiO2 set here would be typical of any set enriched from an unfractionated mixture. While this would need to be explored further, taken together with the Matheron study, these data would seem to suggest that any substantial differences observed previously between TiO2 and IMAC are probably the result of under sampling or a lack of experience in one method or the other. For example, considering that a number of reports suggest that there is a large difference in selectivity between TiO2 and Fe3+ IMAC for singly and multiply phosphorylated peptides44-46, a comparison of the two large rat liver data sets here shows no significant difference between the two (Figure 6C), with the overall number of multiply phosphorylated peptides in each case being 27 and 25%, respectively.

Conclusions Proteomics is a rapidly evolving technology. Instrumentation and sample preparation techniques are constantly improving. The work presented here details the components of an improved and streamlined phosphoproteomic workflow whose goal is to maximize the number of identified phosphosites while maintaining a reasonable throughput. All of the components are readily


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available from commercial sources. Solubilization of proteins in 4% SDS lysis buffer provides improved coverage of the proteome, but care must be taken with the total amount of SDS in the sample prior to trypsin digestion in the FASP filter cartridges else irreproducibility and poor recoveries will be problematic. HILIC fractionation prior to phosphopeptide enrichment provides an added element of selectivity since phosphopeptides are generally more hydrophilic than nonphosphorylated peptides and elute later in the HILIC gradient. By optimization of the gradient, the bulk of the nonphosphorylated peptides elute in the first half of the gradient. Multiply phosphorylated peptides which are the most hydrophilic elute at the very end of the gradient. The very nature of HILIC chromatography generates phosphopeptide fractions where singly phosphorylated peptides are in different pools than highly acidic peptides or multiply phosphorylated peptides. This results in a highly efficient IMAC enrichment. Also because the fractions are 80-70% organic and contain no salts the IMAC resin can be added directly to the fractions with no additional cleanup or preparation. The replacement of micro pipette tips with a filter plate in the IMAC enrichment protocol has greatly improved the ease with which these steps can be performed. In our hands it has also improved the reproducibility and overall recovery. The plates are prepared during the IMAC binding step and then multiple fractions or samples are transferred to the plates and processed simultaneously. Using a multi-channel pipette, 28 fractions in the plate can be processed in the time it takes to do a single sample in a micro pipette tip. With the reversed-phase chromatography system used in our laboratory, we found that analyzing 14 fractions with 90 minute gradients provided the largest number of phosphopeptide identifications in approximately one day of instrument time. We felt that it was important to keep the analysis of a single sample within the one day time frame, as this would allow us, with


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some degree of multiplexing, to conduct a sufficiently powered quantitative experiment in one week’s time. For instance, using 6plex or 10plex isobaric tags, a full dose curve or time series could be analyzed in triplicate in just over three days. Using triple-label SILAC, a 5-point time course could still be finished in triplicate in six days. It certainly is possible to analyze the phosphoproteome in less time. However, in our system this always resulted in a reduced number of phosphosites. We used the described strategy to analyze the phosphoproteome of rat liver. From three samples, we identified more than 20,000 unique phosphorylation sites in 3.5 days. Given that the liver is not a very complex tissue in terms of gene expression and that the animals were healthy adults under normal diets, the number of phosphosites identified probably represents the minimal number of sites which exist as a baseline under homeostatic cellular conditions. Surprisingly, the rat is not a very well studied animal in terms of proteomics. Other than two other studies14,3, this is the most extensive analysis of a rat tissue and the largest single tissue phosphosite data set produced to date.

ASSOCIATED CONTENT Supporting Information. Additional information is provided as noted in the text. “This material is available free of charge via the Internet at http://pubs.acs.org.” The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium (http://proteomecentral.proteomeexchange.org) via the PRIDE partner repository47 with data set identifier PXD001372


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AUTHOR INFORMATION Corresponding Author Email: [email protected] Tel: 610-917-7165 Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT The authors wish to thank Neal Cariello and Lawrence Yoon for the rat liver tissue lysate and Alicia Lundby and Jesper Olsen for generously sharing their Rat Atlas data set. We would also like to thank Dean McNulty and Tim Sikorski for insightful discussions.

ABBREVIATIONS PTFE polytetrafluoroethylene; IDCR ionic detergent compatibility reagent; ACN acetonitrile; TFA trifluoroacetic acid; IMAC immobilized metal affinity chromatography; SDS sodium dodecyl sulfate; DTT dithiothreitol; HFBA heptafluorobutyric acid; PSDVB poly(styrene/divinyl benzene); SS stainless steel


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(37) Qi, L.; Liu, Z.; Wang, J.; Cui, Y.; Guo, Y.; Zhou, T.; Zhou, Z.; Guo, X.; Xue, Y.; Sha, J. Systematic Analysis of the Phosphoproteome and Kinase-substrate Networks in the Mouse Testis. Mol Cell Proteomics. 2014, 13, 3626-38. (38) Zhou, H.; Di Palma, S.; Preisinger, C.; Peng, M.; Polat, A.N.; Heck, A.J.; Mohammed S. Toward a comprehensive characterization of a human cancer cell phosphoproteome. J Proteome Res. 2013, 12, 260-71. (39) Ramsköld, D.; Wang, E.T.; Burge, C.B.; Sandberg, R. An abundance of ubiquitously expressed genes revealed by tissue transcriptome sequence data. PLoS Comput. Biol 2009, 5 (12): e1000598 (40) Marx, H.; Lemeer, S.; Schliep, J.E.; Matheron, L.; Mohammed, S.; Cox, J.; Mann, M.; Heck, A.J.; Kuster, B. A large synthetic peptide and phosphopeptide reference library for mass spectrometry-based proteomics. Nat. Biotechnol. 2013, 31, 557-64. (41) Low, T.Y.; van Heesch, S.; van den Toorn, H.; Giansanti, P.; Cristobal, A.; Toonen, P.; Schafer, S.; Hübner, N.; van Breukelen, B.; Mohammed, S.; Cuppen, E.; Heck, A.J.; Guryev V. Quantitative and qualitative proteome characteristics extracted from in-depth integrated genomics and proteomics analysis. Cell Rep. 2013, 5, 1469-78. (42) Bodenmiller, B.; Mueller, L.N.; Mueller, M.; Domon, B.; Aebersold, R. Reproducible isolation of distinct,overlapping segments of the phosphoproteome. Nat. Methods 2007, 4, 231−237. (43) Matheron, L.; van den Toorn, H.; Heck, A.J.; Mohammed, S. Characterization of Biases in Phosphopeptide Enrichment by Ti(4+)-Immobilized Metal Affinity Chromatography and TiO2 Using a Massive Synthetic Library and Human Cell Digests. Anal. Chem. 2014, 86, 8312-20. (44) Thingholm, T.E.; Jensen, O.N.; Robinson, P.J.; Larsen, M.R. SIMAC (sequential elution from IMAC), a phosphoproteomics strategy for the rapid separation of monophosphorylated from multiply phosphorylated peptides. Mol. Cell Proteomics 2008, 7, 661-71. (45) Zhou, H.; Ye, M.; Dong, J.; Han, G.; Jiang, X.; Wu, R.; Zou, H. Specific Phosphopeptide Enrichment with Immobilized Titanium Ion Affinity Chromatography Adsorbent for Phosphoproteome Analysis. J. Proteome Res. 2008, 7, 3957–3967. (46) Tsai, C.F.; Wang, Y.T.; Chen, Y.R.; Lai, C.Y.; Lin, P.Y.; Pan, K.T.; Chen, J.Y.; Khoo, K.H.; Chen, Y.J. Immobilized metal affinity chromatography revisited: pH/acid control toward high selectivity in phosphoproteomics. J. Proteome Res. 2008, 7, 4058-69. (47) Vizcaino, J. A.; Cote, R. G.; Csordas, A.; Dianes, J. A.; Fabregat, A.; Foster, J. M.; Griss, J.; Alpi, E.; Birim, M.; Contell, J.; O’Kelly, G.; Schoenegger, A.; Ovelleiro, D.; Perez-Riverol, Y.; Reisinger, F.; Rios, D.; Wang, R.; Hermjakob, H. The PRoteomics IDEntifications (PRIDE) database and associated tools: status in 2013. Nucleic Acids Res. 2013, 41, D1063−1069.


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96-well Plate

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Gel-loader Tip

Experiment

Phosphopeptides

Enrichment

Phosphopeptides

Enrichment

Exp1

784

67%

617

73%

Exp2

2827

92%

2240

89%

Exp3

1876

85%

1139

78%

Total

5487

85%

3996

83%

Table 1. Phosphopeptide recovery from pipette tip and 96 well plate


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A

Protein Concentration in 4% SDS buffer (ug/ul)

Lysate Required (ul)a

50 mM Tris (uL)b

Final % SDS c

Amount SDS (mg)

28.6 20.0 10.0 6.7 5.0 4.0 3.3 2.9 2.5

17.5 25 50 75 100 125 150 175 200

182.5 175 150 125 100 75 50 25 0

0.17 0.25 0.5 0.75 1.0 1.25 1.5 1.75 2

0.7 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0

a Volume of 4% SDS sample solution needed to yield 500 ug of protein. b Volume of Tris buffer added to keep the final volume of all sample equivalent. c Each sample also contain 200uL of 8M urea to give a total final volume for each sample of 400uL.

B

Relative Protein Recovery

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


250 100

C

(kDa) 191

200 80

97

150 60

64

51

100 40

39 28 19 14

50 20 0 17.5 25 50 75 100 125 150 175 200

Volume 4% SDS buffer (ul)

17.5 25 50 75 100 120 150 175 200

Input

Volume 4% SDS buffer (ul)

Figure 1. Optimization of FASP for Phosphoproteomics. (A) Experimental design to test the effect of sample loading in 4% SDS on peptide recovery. As the protein concentration of the sample decreases. an increasing volume of 4% SDS buffer is added to the filter to achieve a 500ug load. (B) Protein recovery as measured by the summed UV response from the tryptic digest HILIC chromatogram decreases dramatically with increasing initial volume of lysis buffer. (C) Digestion efficiency in filters decreases dramatically with increasing initial volume of lysis buffer.


33


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 34 of 39

PHOS-Select resin

IMAC coupling 30 min @ RT

FASP prepared trypsin digest 10

20

30 40 Time (min)

HILIC fractionation

Combine fractions and concentrate

50

Fractions moved to the filter plate

PALL AcroPrep Advance 96 Filter Plate Pre-wet with ethanol Wash with NH4 OH/ CH3 CN Wash with acetic acid/ CH3 CN

Wash with acetic acid/30%CH3 CN Wash with water Elution with NH4 OH/30% CH3 CN into a 96-well plate

Figure 2. Plate-based IMAC enrichment. After HILIC fractionation of the FASP derived tryptic peptides.; PHOS-select resin is added directly to the fractions. During this period the filter plate is wetted and washed in preparation for the samples. After transfer to the plate.; the resin is washed and the phosphopeptides eluted directly into a 96 well plate.


34

Page 35 of 39

A Absorbance 214nm (mAu)

1600 1400 1200 1000 800 600 1

400

2

27 28

200 1

0

10

14

20

30

18000

50

60

16421

17000

Phosphosites

40

Time (min)

B 15642 2X 14476 60min

90min

16000 15000

13,630

14000

120min

1X 60min

13000

D

12000 0

20

40

60

100 12,774

12500

14/90min 11500 11,345 10,799

10500

7/120min 28/30min

Relative Site count (%)

C Phosphosites

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


14 90

90

80 70

7 120 14 60 1X

14 14 60 28 120 2X 30

60 50

9500 0

10

20

30

17

Total Run Time (hr)

21

28

28

35

42

Total Run Time (hr)

Figure 3. Evaluation of gradient length and fraction number. Total run time is the sum of the gradient length plus a 30 re-equilibration times the number of fractions. (A) Pooling strategy for HILIC chromatography. One minute fractions are combined to produce 28 samples for IMAC enrichment. The enriched fractions can be further pooled as necessary for LC-MS. (B) Number of phosphosites identified in 14 HILIC fractions using various gradient lengths. (C) Number of phosphosites identified in 28 hours or less using various fraction/gradient combinations. (D) Cumulative summary of the relative performance of all fraction and gradient combinations. The various combinations are noted within each bar.


35


A - 20000

2500 - 15000

2000 1500

- 10000

1000 - 5000

1Singly 2 Phosphorylated 3 4 5 6 7

53-56

49-52

47-48

45-46

43-44

41-42

39-40

37-38

35-36

33-34

31-32

29-30

0

25-28

500 16-24

Phosphopeptides/fraction

3000

Cumulative Unique Phosphopeptides

8 Multiply 9 10 Phosphorylated 11 12 13 14

B Absorbance 214nm (mAu)

- 3500 - 3000

- 2500 - 2000 - 1500

Phosphosites

- 1000

49-52

53-56

49-52

53-56

29-30 31-32 33-34 35-36 37-38 39-40 41-42 43-44 45-46 47-48

16-24

25-28

- 500

C 3000

Phosphopeptides/fraction

2500

2000 1500 1000 500

5 6 7 Unique

47-48

4

45-46

3

43-44

2

41-42

33-34

1

39-40

31-32

37-38

29-30

35-36

25-28

0 16-24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 36 of 39

8 Common 9 10 11 12 13 14

Figure 4. Phosphosites from 14 LC-MS fractions from a typical rat liver sample following HILIC-IMAC enrichment. (A) Singly and multiply phosphorylated peptides per fraction and the cumulative total. (B) Distribution of phosphorylation sites across the HILIC phosphopeptide gradient. (C) Uniqueness of the fractions across the HILIC gradient. In addition to normal adjacent overlap, a number of phosphopeptides are found in many contiguous fractions.


36

Page 37 of 39

A

11.2%

Exp

Date

Phosphosites (all)

Phosphosites (class 1)

Proteins

2.6% pSer pThr pTyr 86.2%

1

April

16,421

10,519

4143

2

May

12,744

8,653

3612

3

June

12,519

8,284

3536

20,953

13,737

4622

total

24%

1P 2P 3P P

B

9000

Number of peptides

8000

150

7000 6000

1P >1P

100

5000

50

4000 3000

0

2000

8.0 8.2 8.2 8.6 8.6 9 9 7.9

1000

0 4.6 4.6

5.4 5.8 5.8 6.2 6.2 6.6 6.6 77 7.4 7.4 7.8 7.8 8.2 8.2 8.6 8.6 9 9 55 5.4

Log10 Intensity

C

Exp1

Exp 1

D 1600

Number of proteins

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


9,879 79%

10,003 80%

1400

1200 Exp 2

1000

Exp 3

800

Exp 2

600

7-10 11-20

400

21-30 31-155

200

Exp 1 Exp 2 9,233 74%

12,944

0 11 22 33 44 55 66

10 20 30 155

Exp 3

Sites per protein Exp 3

Figure 5. Phosphoproteome from triplicate analysis of rat liver using the optimized 14/90 fraction/gradient combination. Three samples of rat liver were processed according to the protocol described herein. (A) Distribution of phosphosites in the three samples.; individually and collectively. (B) Percentage of multiply phosphorylated peptides and their intensity distribution relative to singly phosphorylated peptides. (C) Number of proteins having between 1 and 155 sites. (D) Overlap of phosphorylation sites among the three replicates.


37


A Lundby et al. liver sites*

This study 24,485

Lundby et al. liver proteins*

This study 4,573

1,791

4,870 84%

84%

Phosphoproteins

Phosphosites 20

35 30 25 20 15 10 5 0

FZ Fe 3+ IMAC AL TiO 2

15 10

5

C

7 710101515202025253030353540404545

-3

-2

-1

0

+1

+2

GRAVY Index

Peptide Length (aa) 25

80

20

80

Percentage

0

-3.5 -3 -2.5 -2 -1.5 -1 -0.5 0 0.5 1 1.5 2

B

Percent Phosphopeptides (%)

FZ Fe 3+ IMAC AL TiO2

60

40 20

60

15

0

40

10

1P

>1P

Sites per peptide 20

5 0 1 1.5 2 2.5 3 3.5 4 4.5 5 5.5 6 6.5 7 7.5 8 8.5 9 9.5 10 10.5 11 11.5 12

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 38 of 39

2

4

6

pI

8

10

12

0 1 1 2 2 3

3 4 4 5 5

Phosphosites per peptide

Figure 6. Comparison of rat liver phosphosites from this study and Lundby et al. (14). The three MS raw files from the Lundby liver analysis were searched together with our data in a single MaxQuant experiment using MaxQuant version 1.5.0.25 against the UniprotKB rat reference proteome. The use of the Uniprot database and the tighter thresholds in this version of MaxQuant reduced the number of liver sites in the Lundby data set to 4870. (A) Overlap of phosphosites (left) and proteins (right) in the two studies. (B) Comparison of properties for peptides unique to the two studies. FZ Fe3+ IMAC are data from the present study and AL TiO2 are data from Lundby et al.14. (C) Overall percentage of multiply phosphorylated peptides in the two studies.


38

Page 39 of 39

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Exp 1 Exp 2 HILIC

FASP

12,944

Exp 3 IMAC

Figure TOC


39

An Optimized Platform for Hydrophilic Interaction Chromatography

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