esc.fnwi.uva.nl · Web viewIn this workflow the cells are first broken down after which the...

$: esc.fnwi.uva.nl · Web viewIn this workflow the cells are first broken down after which the protein is harvested. Afterwards, an enzymatic digestion and subsequently fractionation$
Analysis of phosphorylated proteins using MOAC and LC-ESI-MS/MS with Collision Induced Dissociation

(CID) and Electron Transfer Dissociation (ETD)

Tom Panhuise (6059074) University of AmsterdamMaster student Chemistry – Analytical SciencesInternship Master project, 2014-2016Mass Spectrometry of Bio macromolecules – SILS

Professor: prof. dr. C. G. de Koster

1Tom Panhuise (6059074) | Internship Master project 2014-2016 | SILS - Mass spectrometry of Bio macromolecules | University of Amsterdam

MOAC

CID/ ETD

Supervisor: dr. L. J. de KoningDaily Supervisor: H. L. Dekker


AbstractThe analysis of Post Translational Modification’s (PTM’s) is crucial for studies on cellular signaling and other metabolic pathways. Phosphorylation is one of the most extensively studied PTM’s and is involved with many cellular functions. Both the sequence of a peptide and the localization of phosphorylation are important for phosphoproteomic research. This is the reason why a research project was set up by the Mass Spectrometry group of SILS (UvA, Amsterdam), with the goal to propose an analysis protocol for phosphorylated peptides. An extensive literature study was performed in search of relevant protocols which might be implemented and/or adapted for an analysis setup. The resulting protocol combines trypsin digestion followed by enrichment via Metal Oxide Affinity Chromatography (MOAC).The sample is then introduced to Liquid Chromatography (LC), Electron Spray Ionization (ESI) and tandem mass spectrometry (MS/MS). The utilized nanoLC-amaZon Speed ETD/ mass spectrometer (Bruker), enabled the use of both Collision Induced Dissociation (CID) and Electron Transfer Dissociation (ETD) analysis. The fact that ETD leaves labile modifications intact during fragmentation makes it uniquely qualified to be used in phosphorylation site determination. Explorative analysis of different reaction times for ETD analysis proved that the default setting (100 ms) is optimal with regard to intensity of the ions whilst minimizing the potential Proton Transfer Reaction (PTR). Enrichment proved to be most successful when using commercially obtained TiO2 columns instead of ZrO2 or Ti/ZrO2 columns. Additionally, the use of L-glutamic acid and citric acid as additives for improved enrichment of phosphopeptides was investigated. The results indicate that the use of 70 mM L-glutamic acid yields the best enhancement of enrichment. Elution of the peptides requires a high pH, which was supplied by a buffer containing 2M, NH4OH (pH 12). Using a categorization of the “proton mobility” of a peptide, the likeliness to experience neutral loss of phosphoric acid during mass spectrometry analysis was assessed. The resulting workflow has been tested with both simple and more complex samples, regarding the amount of nonphosphorylated material in the sample. However, the resulting work flow might benefit from additional purification steps which are orthogonal to the MOAC enrichment, such as strong cation exchange chromatography (SCX). Additionally, optimization of the parameters of the ion trap system and subsequent data analysis could also greatly improve the overall analysis.

Keywords: Posttranslational modifications (PTM), Phosphorylation, Metal oxide affinity chromatography (MOAC), Reversed phase liquid chromatography (RP-HPLC), Electrospray ionization (ESI), Tandem mass spectrometry (MS/MS), Collision induced dissociation (CID), Electron transfer dissociation (ETD).


Index

1. Introduction

2. Phosphoproteomic Research

2.1 Sample preparation

2.2 Phosphorylated protein enrichment

2.3 Mass spectrometry analysis of phosphoproteins

3. Materials and Method

3.1 Reagents and materials

3.2 Methods

3.2.1 Sample preparation

3.2.2 Phosphopeptide enrichment

3.2.3 LC – MS/MS Analysis

3.2.4 Data analysis

4. Results and Discussion

4.1 Optimization of enrichment

4.1.1 Different MOAC materials

4.1.2 Additives/buffers experimentation

4.1.3 Additive optimization

4.1.4 Elution optimization

4.2 Identification techniques

4.2.1 CID/ETD fragmentation

4.2.2 Proton mobility

5. Conclusion

6. References


1. Introduction

Adaptive nature of cellular machinery is continuously required in order to react to changing conditions in the environment. In order to do so, many proteins are produced and modified in order to maintain the metabolism, structural integrity and specific functions of the cell. Amongst many different methods to cope with these changes, posttranslational modifications (PTM’s) of proteins is an often used strategy by cells.1-3 These modifications consist of either adding extra compounds or removing a specific part of the protein, due to which physical-chemical properties of the proteins change. Some examples of these PTM’s are phosphorylation, SUMOylation, acetylation, glycosylation, although many more exist in nature.1-3 Phosphorylation is one of the most extensively studied PTM’s, and is responsible for protein-signaling transduction, regulation, and many other functions.1, 2, 4-9 When phosphorylation goes wrong, and the subsequent pathways and reactions depending on this modification go awry, many kinds of pathological symptoms can be expressed, such as cancer.10 Phosphorylation of proteins can occur on the specific residues: serine, threonine and tyrosine. These three amino acids have shown to be phosphorylated to different extent in nature. Where serine is phosphorylated frequently (86.4%), threonine (11.8%) and tyrosine (1.8%) are less common phosphorylation sites.3 However, phosphorylation on basic residues (such as histidine, arginine and lysine) might also be phosphorylated to a certain extent.11

Figure 1. The possible workflows for proteomics research, adopted from Switzar et al. (2013).12 Here, three different classifications for proteomic research have been depicted: Top-down proteomics, Middle-down proteomics, and Bottom-up proteomics. The main difference between these three approaches is the size of the analyte, being the greatest in top-down (intact proteins) and the smallest in bottom-up proteomics (small peptides). This study would be classified as a “bottom-up” proteomics approach.

In order to study the effect of specific conditions on post translational modifications of proteins, a means of monitoring the localization, concentration levels, and specific classes of PTM’s is required. The most popular methods nowadays use some form of mass spectrometry (MS) in combination with isolation, purification and enhancement steps in order to investigate these modifications.1, 4, 5, 13, 14 This choice of workflow is well suited for the small quantities, dynamic range and many different isoforms of phosphorylated proteins, which pose a significant challenge to analytical analysis.


Especially orthogonal purification methods have been studied extensively.1, 4, 5, 13 Depending on the research, different approaches are defined, which are named “top-down”, “middle-down”, and “bottom-up”, see figure 1.12 These proteomic approaches use a different general set-up for the experiment, and focus either on whole protein analysis (top-down; ≤50 kDa) or peptide analysis (middle-down; 2-20 kDa, bottom-up; 0.5-3 kDa).12, 15 The different physiochemical properties of peptides can be exploited by using multiple purification methods into one workflow. Since the analytes are proteins, important parameters to separate on are properties such as: pH, hydrophobic/hydrophilic retention, adsorption of specific functional groups to affinity columns, and many more.1, 4, 5, 15-17

To this day, many different analysis approaches have been established for the analysis of phosphorylated proteins using LC-ESI-MS/MS. Thus far, no standard phosphoproteomic analysis workflow was present at the SILS Mass Spectrometry group (UvA). However, improved understanding of these posttranslational modifications is of the utmost importance for development of new cures against diseases, as well as for the understanding of cellular mechanisms. Therefore, the aim of this research project is to implement and optimize a standard workflow for the analysis of phosphorylated proteins. Additional information regarding sample processing, modifications and analysis techniques have been reviewed, but only the relevant techniques are explored in depth.


2. Phosphoproteomic research

A typical workflow (see figure 2) consists of isolation of the proteins followed by fractionation, possible enhancement, and finally LC-MS/MS. Obtained spectra are then subjected to database searching and specific software in order to identify the peptides and localize the phosphorylation sites.4, 5, 15-17 Many phosphoproteomic studies start their workflow with extraction of proteins from cells or tissue followed by digestion. This procedure is often applied before analysis, and many different methods exist for digestion.12

Figure 2. The typical workflow of a “bottom-up” phosphoproteomic research, as illustrated by C. Yang et al. (2014).15 In this workflow the cells are first broken down after which the protein is harvested. Afterwards, an enzymatic digestion and subsequently fractionation are used to obtain specific protein mixtures. These mixtures are enriched (one of many possible methods) for phosphopeptides and subjected to LC-MS/MS, after which a database search and subsequent analysis with specific software is performed.15

2.1 Sample preparation

After selection of the appropriate part of the organism under investigation (cellular membranes, cytosol, or other) the harvested cells are lysed. The choices of homogenization buffer and subsequently denaturing buffer are important, as was shown by Hoffer et al. (2008), whom reviewed the effects of different buffers.4 Here, they showed the increased identification of phosphopeptides when using a 50 mM ammonium bicarbonate buffer, compared to the use of 6 M guanidine, or 1% SDS. Interestingly, the sample had not been denatured beforehand, indicating that protease digestion does not need a denaturing agent. However, the reproducibility of these samples was less than when a solubilized sample was used.

Digestion of the isolated protein mixture can be performed in many different ways. A detailed review of the recent developments and available applications on protein digestion was performed by Switzar et al. (2013).12 Possible ways of digestion consist of enzymatic digestion, and non-enzymatic digestion, which are performed either in gel or in solution. The most common approach is enzymatic digestion of proteins using trypsin, which has been perfected in the past years, resulting in a highly efficient, low cost protease.18 This enzyme is capable of cutting peptide bonds C-terminal to the basic amino acids lysine and arginine, unless followed by a proline.19 Upon digestion with this enzyme, medium sized peptides (~14 residues) are obtained, which have a charge states of at least 2+.12 Although this result is suitable for most kind of analysis, other proteases and digestion methods have


been developed.4, 5, 12 For example, Lys-C, Arg-C, Asp-N, and Glu-C are some of the different endoproteases, which can be used under different conditions than trypsin.12 For example, Lys-C and Arg-C are able to cleave lysine and arginine N-terminally, respectively, and provide high efficiency and specificity. As mentioned by Switzar et al., Lys-C is a recent addition to the list of proteases, and is able to perform under intense denaturing conditions (high temperature, 8 M urea, or 80% acetonitrile).12 Peptides obtained with these kind of enzymes can vary in length (longer) and charge state (higher), making them ideal choices when “middle-down” analysis is performed.12

Besides using proteolytic enzymes, proteins can also be digested by using chemical cleavage or instrumental techniques, which have also been reviewed by Switzar et al.12 Usually acids are used such as formic acid (FA) or acetic acid (HAc), but more are possible. Other examples of chemicals used for protein digestion are cyanogen bromide (CNBr) and 2-nitro-5-thiocyanobenzoate (NTCB). These kinds of chemicals often have a very specific cleavage site, which differs per chemical, resulting in the same level of specificity of digestion as when enzymes are used. The obtained peptides are usually of the size that is useable in middle-down proteomics. The application of a certain chemical for the use of digestion is therefore strongly dependent on the aim of the experiments.

Besides chemicals, instrumental applications have been developed, such as electrochemical oxidation, or the acceleration of digestion (enzymatically) via ultrasound, microwave, infrared, or solvent applications.12, 15, 16, 20 Additional techniques such as immobilized enzymatic digestion, in which the digestion enzymes have been covalently linked to a support, have also been explored.4, 5, 12 The main reason for the development of these non-enzymatic digestion methods is automation. By creating a solid supporting material, or technique with the same resulting digestion as with enzymes, online applications have been realized. Additionally, these techniques require significantly less time to obtain the same results as those obtained whilst using enzymes. Another important aspect of non-enzymatic digestion is the behavior of the sample under the applied conditions (highly acidic or basic). Denaturation is readily achieved when the pH is dropped below 7.4, and might influence affinity-based purification, which is based on specific properties of the target analyte. Therefore, the use of digestion methods other than enzymatic digestion should only be used with a suitable, adapted workflow. Depending on the application and institute, the digestion of samples can be achieved in many different ways. Whereas a high-throughput workflow (non-enzymatic, online applications) might be required in settings such as hospitals or factories, research facilities might benefit more from the use of labor-intensive, but highly specific protocols.


2.2 Phosphorylated protein enrichment

After creating the peptides from the proteins, some pre-fractionation or isolation steps may be performed.4, 5, 12, 15-17, 20, 21 This is usually the case when only a small, select part of the entire sample is of interest (i.e. phosphorylated protein analysis from a whole cell lysate). Many different applications are used, depending on the specific properties of the analyte of interest, see figure 3.16 The potential ionic and Lewis base interactions that phosphate groups are capable of, are important factors for enrichment method development. Besides the charge-based methods, alternative chemical modifications, such as β-elimination with a Michael addition,15 have been developed. However, these kind of techniques are prone to introduce incomplete-, and side-reaction products, resulting in sample loss and further complication of subsequent analysis. Therefore, chemical modifications will not be reviewed here, but can be explored in extensive reviews.4, 5, 12, 14-17 Immuno-precipitation is another enrichment strategy, but is mainly applicable to phosphopeptides containing phosphotyrosine. Both phosphorylated serine and threonine are not readily enriched with this technique, due to lack of specific antibodies.16 Therefore, additional information on this enrichment route can be found elsewhere.15-17

Figure 3. Possible enrichment applications for phosphate containing molecules, by Leitner et al.16 Structural recognition can be used by applying specific antibodies to phosphate groups. Phosphate is able to retain charge, depending on the pH, and is a polar group, which can be exploited by ion exchange chromatography, antibodies and partially metal affinity techniques. Another target of enrichment is the chemical reactivity of the phosphate group.16 In this research, only the Metal Affinity (MOAC) enrichment has been explored.

Phosphorylated peptides are often enriched by means of metal-based materials.22-49 Metal-oxides or immobilized metal-oxide-matrices are able to bind the phosphate groups in a selective way, see figure 4.23 When the pH is chosen appropriately, the free oxygen atoms on the phosphate group can coordinate to the metal, whilst (most) other peptides are not able to do so.43 This gives a powerful way of selectively enriching the sample before applying it to subsequent LC chromatography and mass spectrometry. Many different adaptations of this concept have been explored, such as different kind of metals (titanium, zirconium, germanium, etc.) as well as different kind of matrices (columns, filters, porous beads, etc.). 22-49 Most of these methods show selective behavior towards the binding of phosphopeptides in varying effectiveness. Titanium (Ti), iron (Fe), zirconium (Zr), and germanium (Ge) are some of the more popular choices and have been applied in immobilized-metal-affinity chromatography (IMAC) or metal-oxide-affinity chromatography (MOAC). 15, 22, 25, 28, 36 Evaluating experiments on these different metals showed a difference in the enrichment efficiency, depending on the complexity of the sample and the preparation of the MOAC material, either ZrO2 or TiO2 can be the most effective.34, 38, 39, 42


However, many experiments do not rely solely on metal-based enrichment applications and often incorporate additional separation steps, such as strong cation exchange chromatography (SCX) or antibody-based enrichment, but these kind of approaches have not been considered for this study.7, 8, 15,

17, 21, 49, 50

Figure 4. Coordination of phosphate and chelating bidentates when adsorbed to the TiO2 surface. Other metal-based materials are prone to the same kind of coordination. This specific coordination of both phosphate and carboxylic acids is utilized during additive enhancement of MOAC enrichment. Figure adapted from Larsen et al.23

Besides the enhancement using metal-based materials, many additives to the loading and washing buffer compositions have been investigated for the further improvement of the isolation. These kinds of additives are used to flush out retained molecules which have adsorbed to the material through acidic groups that are also able to coordinate to the metal-based materials if the pH is suitable.27 Most additives are either pH regulators or inhibitors that replace unwanted, retained molecules by competitive binding, such as depicted in figure 4. For the enhancement of phosphorylated peptides using MOAC-type of applications, carboxylic acids have been studied extensively.23, 34 Of these, the substituted aromatic carboxylic acids (2,5-DHB, salicylic acid, and phthalic acid) outperformed the other types of acids tested (such as trifluoroacetic acid, acetic acid, cyclo-hexane-carboxylic acid and benzoic acid) in the prevention of non-phosphorylated peptide retention.23 From these examples, the aromatic compound 2,5-DHB has been studied in great detail and proved to be an effective way to further improve the selectivity of the enhancement of phosphorylated peptides.23 However, the use of this additive is limited to a setup using a MALDI ionization source, since it is also part of the MALDI sample solutions. Usage of this specific additive in other type of ionization source (for example ESI) might lead to severe contamination or damage. Moreover, glutamic acid (0.1 M solution) has shown to be more effective than 2,5-DHB when used in combination with TiO2 for phosphopeptide enrichment, as well as several other additives.22, 27, 31, 34, 39, 42, 48, 51


2.3 Mass spectrometry analysis of phosphoproteins

Following the enhancement step an often applied separation method is Reversed Phase High Pressure Liquid Chromatography (RP-HPLC) or a closely related system.4, 5, 13, 15-17, 20, 24, 38, 40, 41, 51, 52, 53 Liquid chromatography (LC) systems can be directed to an ElectroSpray Ionization (ESI) needle from which the liquid phase is evaporated and gas phase ions remain.41, 52, 54 Ionization using ESI depends on the Rayleigh limit principle, which is concerned with the amount of electrostatic charges per volume density. When a droplet evaporates to such an extent that the charges contained within start to repel each other, the droplet explodes into many tiny droplets. These small droplets can subsequently reach the Rayleigh limit again and explode, leaving charged gas phase ions in the end. By applying ESI to LC fractions, separated analytes are ionized in a “soft” way that keeps them intact and applies a relative high charge state. This is especially useful for low-efficiency fragmentation methods (ETD) or more robust applications (CID). The obtained ions are suitable for injection into the MS system and are subjected to a series of focusing lenses and quadrupoles, which direct the ions to the ion trap as can be seen in the example given in figure 5.55 When reaching the ion trap, fragmentation is either induced by adding an inert collision gas (usually helium) for CID or other means of fragmentation are initiated.

Figure 5. The setup of the AmaZon Speed / ETD mass spectrometer as purchased from Bruker, Daltonics.This mass spectrometer is used in combination with nano-LC-ESI inlet. In this model an optional ETD chemical ionization source is present, see enlarged section.55

The fragmentation of the peptide ions is not exclusively limited to collisional activation, since there are many other ways to introduce energy into the compounds being analyzed. Most of the techniques require a specific setup or modification to a standard mass spectrometer in order to perform the required fragmentation. The fragmentation techniques explored here are: Collision Induced Dissociation (CID), Electron Transfer Dissociation (ETD) and a hybrid mode in which CID is followed up by ETD or vice versa.14, 55-63


Whereas the CID technique is straightforward, the ETD technique requires a specific ion source that is capable of producing radical anions (usually fluoranthene 13 is used as reactant). These two fragmentation methods produce different kind of fragment ions, due to differences in the methods of introducing energy and the resulting fragmentation reactions.

The CID technique produces b- and y-ions which are the result of a peptide bond being broken (Ccarbonyl-N, cleavage activation energy required ~40 kcal/mol)64, see figure 6. However, the energy supplied during the collisions is more than enough to also break the weaker phosphate bond (O-P bond, cleavage activation energy required <20 kcal/mol)65, which results in either neutral losses of phosphate related fragments (H3PO4, 98 Da loss; HPO3 or HPO3 + H2O, 80 Da loss) or charged losses (PO2

-, 63 Da loss; PO3-, 79 Da loss; H2PO4

-, 97 Da loss). Electron transfer dissociation however, produces c- and z-ions, see figure 6.56, 57, 66-71 These ions are formed after breaking the N-Cα bond of the peptide backbone and are the result of radical reactions.67 Unlike CID, ETD fragmentation does not target the labile bonds between phosphate groups and the peptide, making it an ideal method to study the localization of the phosphorylation.57, 68, 69, 72, 73 However, ETD does require high charge densities on the peptides of interest, due to the formation of non-dissociated charge reduced radical cations.14, 57 These kinds of ions are not effective in the investigation of amino acid sequence, phosphorylation site localization or peptide identification. Multiple methods exist in order to make sure that ETD results in highly efficient ionization and fragmentation, such as the application of CID after ETD (ETcaD).56 Other methods consist of using different digestion enzymes that produce higher charge states peptides, or the use of additives which increase the charge state of the peptides in solution, as has been described previously.4, 5, 12-14 However, upon digestion with trypsin, phosphorylated peptides will usually carry a low charge state due to the deprotonated phosphogroups.

Figure 6. Nomenclature of the fragmentation ions of the peptide backbone.Traditional CID fragmentation yields b- and y-ions, whilst ETD fragmentation yields c- and z-ions. The numerals denote the amount of amino acid residues contained within the ion. The amino acid residues (the specific groups) are denoted with R.14

Many different mechanisms have been suggested for the neutral loss of phosphate related fragments upon ionization (H3PO4, 98 Da loss; HPO3 or HPO3 + H2O, 80 Da loss).14, 59, 60, 63, 74 Although it was thought that these reactions occurred via β-elimination with the charge remotely placed, Palumbo et al demonstrated that this is not the case.74 Controversially, the evidence supported a “charge-directed” mechanism in combination with a SN2 participation reaction, which led to the creation of a cyclic product ion, as can be seen in scheme 1. In this study they also promoted the relations between proton mobility and the gas-phase fragmentation reaction pathways of modified peptides, as will be discussed later on.


Scheme 1. Three potential reactions for the loss of a neutral group (H3PO4) as demonstrated from a phosposerine containing peptide.Pathway A has been widely accepted and supposedly supported by empirical data. Whereas, both pathway B and C have been obtained by molecular orbital calculations and experimental data. A: Charge-remote β-elimination reaction. B: Charge-directed E2 elimination reaction. C: Charge-directed SN2 neighboring group participation reaction.74

Since the development of ETD several different mechanisms have been proposed for the transfer of an electron to a peptide. The Cornell-mechanism, as theorized by McLafferty and colleagues, is based on the uptake of an electron at a protonated site, see scheme 2 (usually a basic amino acid residue or the N-terminally amine group).57 This results in a radical species which is typical for these kinds of mechanisms. The excited electron would then undergo relaxation upon which a proton is transferred to the oxygen located in the peptidebond. In turn, this would result in an intermediate species containing an aminoketyl radical, as can be seen in the third section of scheme 2. The last step in this mechanism is the cleavage of the N-Cα bond situated to the right of the radical site. This reaction mechanism is charge-directed in the sense that a protonated site initiates the fragmentation. The site of protonation therefore dictates which kind of ions are observed; c- or z-ions.

Scheme 2. The Cornell mechanism as described by McLafferty et al. This mechanism shows the fragmentation of the N-Cα bond upon electron transfer dissociation with a charge directed from a C-terminal amine group. Note that this could also be initiated from the N-terminus resulting in differently charged c- and z-ions.57

Charge-remote fragmentation has also been investigated and different mechanisms have been suggested, such as the Utah-Washington mechanism, see scheme 3.57 Here, the initial electron uptake is thought to happen directly on the amide group, specifically in the π* antibonding orbital. This forms the aminoketyl radical intermediate which was also found in the previous mechanism indicating the importance of the creation of this intermediate species. Depending on the specific mechanism, the anionic charge is negated with the attraction of a proton and the amide bond is broken (Washington mechanism, Utah supposes the reverse order). Depending on the source of the proton used in the neutralization of the anion, either a c- or z-ion can be observed.


Scheme 3. The Utah-Washington mechanism as proposed by Simons et al and Turecek et al. This mechanism shows the fragmentation of the N-Cα bond upon electron transfer dissociation without charge initiation . Note that this could also be initiated from the N-terminus resulting in differently charged c- and z-ions.57

Another mechanism suggested by Tsybin et al uses the “enol”-like properties and differs greatly from the previous described mechanisms.57 The first difference is the location of the fragmentation of the N-Cα bond, which occurs N-terminally rather than C-terminally, see scheme 4. Moreover, the amide bond is cleaved heterolytically in contrast to the previously used homolytical cleavage. This results in the highly basic c-fragment and the z+•-ion as seen in the last panel of scheme 4. The resulting charge and composition of the fragments are independent of the initial hydrogen donation site, which is also different in the previously mentioned mechanisms.

Scheme 4. The “enol” mechanism of ETD as suggested by Tsybin et al. Here the mechanism is shown for the fragmentation of the N-Cα bond upon electron transfer dissociation. The c-fragments often abstract a proton from the z+•-ion in order to form the c’-ion.57

Fragmentation obtained by using either CID or ETD techniques results in mass-to-charge ratios in a MS spectrum. The fragments can be directly analyzed (MS) or can be subsequently filtered for specific target ions (MS/MS, MSn), resulting in cleaner spectra of those ions.14, 56-59, 62, 75-77 Using a peak picking acquisition method for additional rounds of fragmentation of a specific ion is an often used approach. This is especially useful in combination with CID, by scanning for signals that result from the neutral or charged losses of phosphate groups. From the subsequent fragmentation of the isolated ion, the peptide on which the phosphate group was present can be identified.

Although historically the obtained spectra were analyzed by hand, nowadays large databases and extensive software packages exist to help organize and analyze the obtained results.77-83 Firstly, the obtained raw spectrum is subjected to software to clean up and prepare the data for being used in database searches. Some of these programs can also annotate sequences from the raw data, when appropriate methods have been used. Matching the peaks in the spectrum to specific peptides is performed by search engines such as MASCOT,3 which uses statistics and probability based scoring methods.81-83 After the link has been made between peptide and protein, specific parameters can be monitored in order to optimize the entire method. These include protein coverage, which states how much of the whole protein has been covered by the obtained peptides, and protein scoring. Usually a probability based method that states the expectancy values of obtaining the same scoring using random entries is used to gauge the significance of the match.83


3. Materials and Methods

3.1 Reagents and materials

Dithiothreitol (DTT), iodoacetamide (IAA), citric acid, MS PhosphoMix 2 Light (MSP2L), β-casein (from bovine milk), ammoniumbicarbonate (NH4HCO3) and L-glutamic acid were obtained from Sigma Aldrich (St. Louis, MO, USA). Water (H2O, ULC-MS grade), acetonitrile (ACN, ULC-MS grade), trifluoroacetic acid (TFA, ULC-MS grade) and formic acid (FA, ULC-MS grade) were purchased from BioSolve (Dieuze, France). Hydrochloric acid (HCl, 7.7 M) and ammonia solution (NH3, 25% solution) were procured from Merck (Kenilworth, NJ, USA). Bovine serum albumin (from bovine pancreas) was obtained from Boehringer Ingelheim (Ingelheim am Rhein, Germany). Trypsin (Trypsin Gold-Mass Spec Grade) was purchased from Promega (Madison, WI, USA). OMIX C-18 pipette tips (100 μL, 80 μg capacity) were obtained from Agilent Technologies (Santa Clara, CA, USA) and OMIX C-18 pipette tips (10 μL, 8 μg capacity) were bought from Varian (Palo Alto, CA, USA). TopTip micro-spin columns (TT1) packed with MOAC media (TiO2, ZrO2 and a mixed material of 50/50 Ti/ZrO2) were purchased from Glygen (Columbia, MD, USA).

3.2 Methods

3.2.1 Sample preparation

Samples from proteins were digested before analysis with the use of trypsin (Gold, Promega). Both bovine serum albumin and β-casein were digested using 50 μg of protein. The digestion buffer for BSA consisted of 0.1 M ammonium bicarbonate and 10% acetonitrile and water. Reduction of the protein was performed by adding 5 mM dithiothreitol to the mix and letting it react at 55˚ C for 30 minutes. Subsequent alkylation of the cysteine residues was performed using 15 mM iodoacetamide, which was left to react for 20 minutes in the dark at room temperature. After reduction and alkylation, digestion was performed by adding 1:50 (w/w) trypsin at 37˚ C overnight. Afterwards, the digestion mix was lyophilized using liquid nitrogen and a freezedryer (Thermo, HETO PowerDry LL1500 Freeze dryer). The dried sample was then resuspended in 20 μL of a solution containing 50% ACN and 2% formic acid and frozen with liquid nitrogen and stored at -26˚ C until further use. The final concentration of these aliquots was 0.04 nmol/μL. β-Casein is a phosphorylated protein and was used because of the mixture of phosphorylated and nonphosphorylated peptides obtained upon digestion. This protein was digested using a similar method as before, although it contained 20% ACN in the digestion buffer and no reduction or alkylation was necessary prior digestion. The end concentration of the β-casein digestion aliquots was 0.1 nmol/μL.A synthetic phosphopeptide kit was used (MSP2L) which is a mimic tryptic digestion of HeLa cell proteins, and no digestion was therefore necessary. This kit contained a total of 200 pmol of peptides, 20 pmol per peptide, see supplemental information. Upon arrival, the kit was dissolved in 40 μL of a solution containing 20% ACN and 0.1% FA as was suggested in the manufacturer’s notes. Aliquots were made of 2 μL, which were snap frozen using liquid nitrogen and were stored at -80˚ C until use. These aliquots contain a concentration of 5 pmol/μL of total peptides, or 0.5 pmol/μL for the separate peptides. Upon arrival a preliminary test run was performed to check the visibility and behavior of the peptides after running a 30 minute LC gradient and CID fragmentation in a MS/MS analysis. From this data it was determined that not all the peptides (especially 2.2) were readily observable, see supplemental information.


3.2.2 Phosphopeptide enrichment

Phosphopeptide enrichment was carried out using the TopTip micro-spin columns (TT1) with different packing material (TiO2, ZrO2 and Ti/ZrO2). The enrichment relies upon the coordination of the phosphorylated peptides to the material and several washing steps, as described before. A standard enrichment workflow consists of activation of the MOAC material by subjecting it to the loading buffer (without introducing the sample). Afterwards, the sample (diluted in the loading buffer) is loaded on the material and subsequently washed with more loading buffer. This is followed by additional washing with a solution containing at least 65% ACN and 2% TFA to remove non-coordinated peptides. The next step is an elution with a very basic solution which needs to be acidified quickly afterwards in order to retain the phosphategroups on the peptides. A general workflow can be seen in scheme 5. During this experiment the following samples were obtained before, and during enrichment: Negative control, Flow through plus wash fraction, and Eluted fraction. The control sample was obtained by splitting the untreated sample in equal portions and storing one of these portions as control, the other half was used for subsequent enrichment. For example, when using the kit (MSP2L), samples were prepared by adding 2 μL of the kit into 18 μL loading buffer and splitting this sample in 10 μL enriched and 10 μL negative control. Flow through and wash fraction were obtained during loading and washing of the sample and were pooled together. The eluted fraction was obtained after elution and acidification of the retained (phospho)peptides. Due to the chemicals in the loading buffer, samples should be desalted and lyophilized before analysis. The resuspension buffer which was used to dissolve the samples after desalting and freeze drying, contains 2% ACN and 0.1% TFA. It was also important to choose the right capacity for the desalting pipette tips, since some samples may contain a large amount of peptides (i.e. wash fractions should contain all the nonphosphorylated peptides). A specific loading buffer was used when additives were used during the enrichment, in which the sample was dissolved. The different loading buffers used during this study were:

o 80% ACN, 2% TFAo 65% ACN, 2% TFAo 65% ACN, 2% TFA, L-Glutamic acid (35 mM, 70 mM, and 0.14 M)o 65% ACN, 2% TFA, Citric acid (50 mM, 0.5 M, and 1 M)

From these loading buffers, only the last two contained an additive. These same loading buffers were used to activate the column and wash the loaded sample. The sample was then washed using a different washing buffer. The used washing buffers were:

o 80% ACN, 2% TFAo 65% ACN, 2% TFA

The elution buffer that was used during these experiments has been used with two different pH values:

o NH4OH (2 M (~2% solution), pH 10.5 and pH 12)

This solution was directly acidified after elution, using 1:1 (v:v) of a 10% TFA solution. This was done to prevent the loss of phosphoric acid due to the basic environment.


General workflow of phosphopeptide enrichment using MOAC columns: Dissolve the sample in the loading buffer. Keep in mind that you will split the sample in two:

one for enrichment, one for control.

Gently tap the TopTip column on a flat surface so that the material settles at the bottom.

Remove the red cap on the top and hold the column over an Eppendorf container.

Activate the material with 10 μL of the loading buffer and use a syringe to push the liquid

through (you can’t use negative air pressure as it will disturb the material). (3x) Switch the column to a new Eppendorf marked as the Flow through + wash sample.

Load half of the prepared sample on the column. Store the rest of the sample as Control.

The flow through could be loaded again, to make sure that all phosphopeptides are bound to

the material.

Wash the column with 10 μL of the loading buffer (3x). Wash the column with 10 μL of the washing buffer (3x). Switch the column to a new Eppendorf marked as the Eluted sample.

Elute the sample with 10 μL of elution buffer (3x). Quickly acidify the eluted fraction with 30 μL of 10% ACN solution.

If an additive was used, choose a tip with appropriate desalting capacity, and desalt.

Afterwards, freeze dry the sample and resuspend in such a volume that the concentration is

equal to the concentration you have in the control sample.

Store the samples at -20° C until analysis.

Scheme 5. A general workflow for the enrichment of phosphopeptides using MOAC columns. Depending on the composition of the loading-, washing-, and elution-buffers, steps may vary between experiments. Modified from Glygen manual as provided with purchase of the TopTip.

When enrichment was performed on the phosphopeptide kit (MSP2L) the amount of sample loaded onto the columns was 1 μL mixed into 9 μL loading buffer (50 fmol/ μL) the same amount was set aside as a negative control sample (untreated sample).

The same amount of sample was loaded for analysis for each separate experiment. This means that results from different experiments cannot be readily compared to each other, but comparison per experiment should be able. However, scaling of the data with regard to the mass balance between the samples was not performed due to lack of a suitable method and the small amounts of the samples. Unfortunately, all the experiments performed during this study have been done only a single time due to time constraints. This means that all the data presented represents single measurements and can only be used to give indications about certain trends. These results therefore represent the first steps into developing a phosphopeptide specific enrichment workflow for the mass spectrometry group at SILS (UvA).


3.2.3 LC – MS/MS Analysis

After enrichment of the phosphopeptides, the samples were analyzed using a nanoLC (Easy – nLC II, Bruker) coupled to an Advance Captive Spray (ESI) in combination with the amaZon Speed / ETD mass spectrometer from Bruker. The ACS electrospray source is a modification from the original Apollo II Electrospray Source which is supplied with the amaZon Speed mass spectrometer. The Advance Captive Spray source is easier to operate and tune in contrast to the other source.

The nanoLC was set to undergo a linear gradient using two solutions: solution A: water, 0.1% formic acid solution, and solution B acetonitrile, 0.1% formic acid solution. The linear gradient was completed in 25 minutes with 5 minutes delay at the start, making the LC-run 30 minutes total. The gradient starts at 5% B and 95% A, which was ramped up to 12% B and 88% A in the first 5 minutes. During the next 15 minutes the gradient was slowly increased to 30% B and 70% A, which was the highest concentration used in peptide/protein analysis. After this functional gradient the column was flushed with 95% B and 5% A over 4 minutes and reset to 5% B and 95% A during the last minute. The nanoLC setup was operated at a flow rate of 300 nL/min.

The amaZon Speed / ETD mass spectrometer used for this study has been used with the standard settings as supplied by the manufacturer, unless specifically stated otherwise (i.e. ETD reaction time experiment). Only the “positive mode” was utilized during this, the “negative mode” has not been explored. However, the “negative mode” might be worth looking into because of the negative charges phosphopeptides carry. Especially for multiple phosphorylated peptides the resulting charges after protonation might not be suitable for “positive mode” detection. When using MS/MS analysis, the first round used “enhanced resolution” whereas the second used “Xtreme resolution”. Additionally, “enhanced resolution” used the average of 5 measurements for every spectrum, whereas “Xtreme resolution” used averaging of 2 measurements. The resolution obtainable when “Xtreme Resolution” was applied, was 2+ ions with a scan speed of 52,000 u/sec. When using the “Enhanced Resolution” these parameters changed to a resolution of 4+ ions with a scan speed of 8100 u/sec.55 These rounds of fragmentation use helium as collision gas and require a fragmentation time of 28 milliseconds. The mass range analyzed with this setup ran from 300 m/z to 1400 m/z.

Figure 7. A schematic representation of the ETD reaction during ETD analysis, from the brochure of the AmaZon Speed / ETD mass spectrometer as purchased from Bruker, Daltonics.In this simplified reaction scheme the produced radical anion reagent from fluoranthene reacts with the peptide of interest after which the N-Cα bond is broken.55


For ETD measurements the specific chemical ionization cartridge had to be activated, see figure 5. This cartridge uses fluoranthene and methane gas to create an electron donating radical anion species which enables the cleavage of the N-Cα bond in a peptide, see figure 7. The reactions towards the anionic radical species start with the transfer of high energy electrons (~60-80 eV) from filaments to the mediator methane. After this interaction, two lower energy electrons are available for a reaction with the fluoranthene, which will result in C16H10

●─, as can be seen in scheme 6. However, this specific chemical is also used for the Proton Transfer Reaction (PTR), which differs from ETD in the sense that it donates a proton instead of creating a radical species. This PTR reagent is easily created when a proton is absorbed by the radical anion, creating a C16H11

─ carbanion, see scheme 7. Both reactions can be carried out using the same equipment and with only minor tweaking of chemical ionization reaction time and heating of the fluoranthene. This very flexible setup gives a choice to the user, but also poses an inherent problem. Due to the dual nature of the chemical used for PTR/ETD reactions, both reactions take place to some extent.71 Although the manufacturers have optimized the parameters for each kind of reaction, some side reactions are to be expected.71

Scheme 6. The reactions leading to the creation of the radical anion species used during ETD analysis.The reaction is started with a filament that is heated up to release high energy electrons (~60-80 eV) which can be captured by the methane. Absorbing part of the energy of this high energy electron, together with the release of an electron from methane itself, this results in two electrons of lower energy. These low energy electrons are then able to interact with fluoranthene to form the radical anion species that is used during the ETD reaction inside of the ion trap.

Scheme 7. Schematic representation of the creation of the PTR reagent from the radical anion species.After creation of the radical anion species in the chemical ionization cartridge, another reaction may take place. During this follow up reaction a proton is absorbed by the radical anion which will produce a carbanion. This compound is readily able to donate a proton to a peptide when introduced to the ion trap.

In addition to direct data dependent analyses, ETD – MS/MS analyses were also performed during which ETD was triggered by the detection of CID loss of neutral fragments with a nominal mass of 98 and 80 Da. These measurements used “enhanced resolution” for the primary CID fragmentation and used the average of 5 measurements. The subsequent round of ETD fragmentation upon neutral loss detection utilized “Xtreme resolution”, which required 100 ms of reaction time in the iontrap and used the average of 3 measurements.


3.2.4 Data analysis

The raw MS/MS data was processed with the Data Analysis program version 4.2 (Bruker Daltonics, Bremen Germany). With this program the results were deconvoluted and searched for compounds using the AutoFind (MS/MS) function. Full compound lists were extracted containing the CID and ETD data (if acquired), in order to use in subsequent database searching. The search engine used for the database searching was the Matrix Science server, MASCOT. An in house operated server (version 2.5.1) was used over the available online servers, because this allowed for larger data files to be uploaded. The used settings for the MS/MS Ions Search were dependent on the specific experiment, but the overall settings used were:

MSP2L and BSA analysis: Database: Swissprot (550,116 reviewed entries, UniProtKB). Digestion: trypsin (1 missed cleavage allowed). All taxonomy allowed. Fixed modifications: carbamidomethyl (C). Variable modifications: oxidation (M); phosphorylation on (ST) and (Y) . Peptide tolerance: ± 0.5 Da. MS/MS tolerance: ± 0.4 Da. Peptide charges: 1+, 2+, 3+ (For future analyses; 2+, 3+,4+ is recommended). Error tolerant on. Instrument: CID: ESI-Trap; ETD: ETD-Trap.

The kit (MSP2L) was analyzed using the swissprot database instead of an in house created phosphopeptide kit database, due to the ability to also detect BSA when this was spiked in.

β-casein analysis: Database: Contaminants (in house). Digestion: trypsin (1 missed cleavage allowed). All taxonomy allowed. Fixed modifications: none. Variable modifications: oxidation (M); phosphorylation on (ST). Peptide tolerance: ± 0.5 Da. MS/MS tolerance: ± 0.4 Da. Peptide charges: 1+, 2+, 3+ (For future analyses; 2+, 3+,4+ is recommended). Error tolerant on. Instrument: CID: ESI-Trap; ETD: ETD-Trap.

When measuring β-casein a contaminants database was used (matrix science, EMBL) since this protein is an often encountered contaminant. The use of the contaminant database is strictly useful when preforming optimization experiments with a compound which is listed in this database (for example β-casein). The use of this database is not recommended when analyzing other samples, because it is very limited in the amount of entries and might produce false positive results.

4. Results and Discussion


4.1 Optimization of enrichment

4.1.1 Different MOAC materials

The approach to set up a standard workflow for the analysis of phosphopeptides combines the use of the amazon Speed / ETD mass spectrometer from Bruker Daltronics 55 and commercially obtainable MOAC tips. To this end, three different MOAC tips were bought (TopTip) containing three different materials: TiO2, ZrO2 and Zr/TiO2. Due to the ambiguity in literature which material performs best, an assessment of the obtained materials was needed. The performance of these tips on phosphopeptide enrichment has been tested using a phosphopeptide kit (MSP2L, see supplemental information) and the following conditions:

Loading buffer: 80% ACN, 2% TFA Washing buffer: 80% ACN, 2% TFA Elution Buffer: 2M NH4OH, pH 10.5 Acidification: 1:1 (v/v) with 10% TFA

For each round of enrichment, the following samples were taken: control (50% of the starting mixture), flow through and wash (pooled), and the eluted fraction. The samples were freeze dried and dissolved after enrichment, after which they could be desalted. The desalting step is not absolutely necessary at this stage, since the samples contain no additives or salts and are freeze dried before analysis. However, due to the experimentation with additives later on, the workflow was kept the same for all the samples. Subsequently, these samples were analyzed upon the mass spectrometer and CID data was obtained. The obtained data was processed and presented to a database search using the in-house MASCOT server, from which the following ion scores were collected, see table 1.

Table 1. Ion score and presence of the phosphopeptides from MSP2L after MOAC enrichment using different materials.

MOAC:

Sample type

2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.92.10

TiO2

Control 49 - 31 59 32 55 10 15 - 68FT + wash - - - 69 a - - - - - -Eluted 46 - 31 79 29 61 13 16 - -

ZrO2

Control 51 - 28 64 33 65 22 12 - -FT + wash - - - - - - - - - -Eluted 46 - 27 19 - - - - - -

Ti/ZrO2

Control 51 - 29 60 42 63 15 12 - -FT + wash - - - 32 a - - - - - -Eluted 44 - 21 51 - 11 - - - -

The MASCOT ion scores of the identified phosphopeptides (MSP2L) after enrichment on different Metal Oxide Affinity Chromatography materials (TiO2 ZrO2, and Ti/ZrO2). For each enrichment experiment three samples were collected; not enriched negative control (Control), a pooled fraction of the flow through and wash fractions (FT + wash), and the eluted fraction (Eluted). Green: The assigned ion score from MASCOT. - : Not observed. a: Only observed with neutral loss of phosphoric acid.


From these results, it becomes evident that more peptides are found in the TiO2 eluted fraction than in both the eluted fraction from ZrO2 or Zr/TiO2 enrichment. However, the loss of peptide 2.10 in the titanium enrichment shows, as well as the loss of other peptides from the zirconium- and mixed-enrichment, that the enrichment is prone to sample loss. Additionally, the ion scoring of the peptides is not increased when compared to the control run as was expected, since the phosphopeptide kit was not mixed with any other peptides. This indicates the potential for enrichment, since phosphopeptides can be retained by the materials during washing and eluted from the material with little loss.After the potential for enrichment with the TopTips was confirmed, enrichment from a complex mixture was performed. This was done in order to simulate a more realistic sample, in which both phosphorylated and nonphosphorylated peptides will be present upon tryptic digestion. In this experiment the MSP2L phosphopeptide kit was mixed 1:50 (molar ratios) with tryptic Bovine Serum Albumin (BSA) digest. Again the three kinds of MOAC tips were tested for enrichment, using the same experimental conditions as used in the previous experiment although with a slight modification to the washing buffer. This change in concentration of acetonitrile in the washing buffer was done because of optimization experiments (which will follow shortly) from which it became evident that this is also a suitable concentration. Besides, this concentration was required in later experiments, due to the need for a more hydrophilic solution when solvating the additives.

Loading buffer: 80% ACN, 2% TFA Washing buffer: 65% ACN, 2% TFA Elution Buffer: 2M NH4OH, pH 10.5 Acidification: 1:1 (v/v) with 10% TFA

Again, samples were freeze dried and resuspended in (2% ACN, 0.1% TFA), after which they were desalted using OMIX tips. In the same fashion as before, three kind of samples were taken (control, flow through + wash, and eluted) which were also analyzed the same way. Again, only the CID data was collected during the mass spectrometry analysis, which was used to obtain ion scoring after database searching.

Table 2. Ion score and presence of the phosphopeptides (MSP2L) after enrichment from a complex mixture using different MOAC materials.

MOAC: Sample type 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9 2.10

TiO2

Control - - - - - - - - - -FT + wash - - - - - - - - - -Eluted 48 - 14 72 32 28 2 b 3 20 29

ZrO2

Control - - - - - - - - - -FT + wash - - - - - - - - - -Eluted 46 - - 27 - - - - - -

Ti/ZrO2

Control - - - 17 - - - - - -FT + wash - - - - - - - - - -Eluted 48 - - 46 - - - 14 - -

The obtained MASCOT ion scores of the identified phosphopeptides after enrichment from a complex 1:50 (molar ratios) mixture (containing Phosphopeptides amd BSA digest, respectively). Green: the assigned ion score from MASCOT. - : Not observed. b: Only observed with transfer of the phosphogroup to another position.


From table 2, it directly becomes clear that (almost) no phosphopeptides have been detected in the control and washing samples. However, this is to be expected from a complex sample, since the phosphopeptides will be suppressed during ionization by the overwhelming amount of nonphosphorylated peptides. Furthermore, during the washing (and loading) we expect no phosphopeptides to be detected since they should coordinate to the MOAC material. During elution we hope to find all the phosphopeptides back, as is the case with the titanium dioxide material (peptide 2.2 has never been detected, see supplemental information). Sadly, we don’t see the same kind of recovery with the zirconium containing columns. Controversially, enrichment of the complex mixture with the TiO2 column made it possible to detect the 2.10 peptide, which was not detected in the enrichment using only the MSP2L kit. This could be a result of the change in washing buffer (from 80% ACN to 65% ACN), which would indicate the necessity of appropriate washing conditions. However, it has to be noted that all of the eluted samples contained peptides originating from BSA, with very high protein scoring (ranging between 2497 – 6193) and large protein fragmentation coverage (38%-74%). This indicates incomplete enrichment of the phosphopeptides from the mixture, and might suggest additional washing or purification steps are necessary. These results also support the necessity of adding additives to the loading and washing buffers, which would compete with the nonphosphorylated peptides for binding to the MOAC material.


4.1.2 Additives/buffers experimentation

The effect of different additives and buffer conditions on the enrichment of phosphopeptides was tested using the three different MOAC materials as well as two acidic additives; L-glutamic acid and citric acid. These acids, and their concentrations, have been chosen after evaluating literature for appropriate additives and due to their ability to coordinate to the materials, see figure 8. Due to the previous results, only the trials with TiO2 are shown below. The approach of the experiments was very much similar to the previous experiments in regard to obtaining different fractions (control, flow through + wash, eluted) but with different buffer compositions. Both simple and more complex samples (β-casein tryptic digest, 1:50 molar ratios of MSP2L: BSA tryptic digest) have been analyzed using this workflow.

Figure 8. Molecular structures of the two additives used during the MOAC enrichment.Both compounds were chosen after study of recent literature as well as consideration of the suitability for the experimental setup. Left: L-glutamic acid, right: citric acid.

Comparison of the enrichment of the phosphopeptides has been performed by comparing ion scores as well as the scoring and coverage for the BSA protein or β-casein. Ideally, the eluted fraction would yield only phosphopeptides identifications with high ion scores and low protein scores for the used nonphosphorylated protein (both BSA and β-casein).

In order to test the effect of the percentage of acetonitrile in the washing buffer on the enrichment of phosphopeptides, an experiment was performed using β-casein tryptic digest, of which the peptides can be seen in the supplemental information. In this test, two different concentrations of acetonitrile were used during the washing steps, 65% and 80%. However, both the samples were subjected in a loading buffer containing 80% ACN. The following buffer compositions have been used in these experiments:

Acetonitrile buffer: Loading buffer: 80% ACN, 2% TFA Washing buffer 1: 80% ACN, 2% TFA Washing buffer 2: 65% ACN, 2% TFA Elution Buffer: 2M NH4OH, pH 10.5 Acidification: 1:1 (v/v) with 10% TFA

Before enrichment control samples were collected, during the enrichment both a wash + flow through and eluted sample were collected. Due to mechanical problems with the vacuum dryer, small aliquots (3 μL) of the samples were dried using a vacuum centrifuge (alcohol mode, 35 ˚C, 10min). The dried samples were then dissolved in six microliter of a 2% ACN, 0.1% TFA solution. These samples have been analyzed by using CID and a 30 minute LC gradient, after which protein scores and coverage as well as presence of phosphopeptides of β-casein was determined.


Control 6

5%

Wash+FT

65%

Eluate

65%

Control 8

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80%

Eluate

80 %0

250

500

750

1000

1250

1500

1750

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Figure 9. Protein scoring and coverage of β-casein after enrichment on titanium dioxide with different concentrations of acetonitrile in the washing buffer. The filled bars represent the MASCOT protein score (arbitrary units) for β-casein on the left y-axis, the faded bars represent the protein coverage in percentages on the right y-axis. The three left bars indicate the results from samples using a 65% acetonitrile washing buffer; control, wash + flow through and eluted. The three bars on the right indicate the results obtained when washing was performed with a 80% ACN washing buffer; control, wash + flow through and eluted.

Since titanium proved superior to zirconium and the mixed material, only the enrichment of phosphopeptides on TiO2 material was assessed here. Observing the protein scoring in figure 9 shows the indication that washing the column with 65% acetonitrile yields similar results as when 80% is used for the eluted sample. A decrease in the scoring and coverage can be seen when comparing the the control and the eluted samples from the 65% acetonitrile washing buffer experiment. This lowering of the protein score could be an indication that less nonphosphorylated peptides are present after washing and eluting the peptides from the MOAC material. However, it is to be noted that both eluted samples from both experiments yield a similar score and coverage after enrichment. Along with the knowledge that in both cases the same sets of phosphopeptides could be found, this gives another strong argument that changing the percentage of acetonitrile during the washing step is not influential.


4.1.3 Additive optimization

After establishing that changing the percentage of acetonitrile in the washing buffer will not benefit the enrichment much, new experiments were performed in which additives were used. According to literature, both L-glutamic acid and citric acid proved to be promising additives for enrichment of phosphopeptides. Therefore the following buffers were prepared for the enrichment of phosphopeptides from a commercially available kit in a complex mix (MSP2L:BSA digest | 1:50):

L-Glutamic acid buffer: Loading buffer: 65% ACN, 2% TFA, 0.14M L-glutamic acid Washing buffer: 65% ACN, 2% TFA Elution Buffer: 2M NH4OH, pH 10.5 Acidification: 1:1 (v/v) with 10% TFA

Citric acid buffer: Loading buffer: 65% ACN, 2% TFA, 50 mM Citric acid Washing buffer: 65% ACN, 2% TFA Elution Buffer: 2M NH4OH, pH 10.5 Acidification: 1:1 (v/v) with 10% TFA

The obtained samples (control, wash + flow through and eluted) were then analyzed using a 30 minute LC gradient prior to CID fragmentation. After processing of the data, the ion scores of the specific peptides were recorded and are shown below, see table 3.

Table 3. Ion score and presence of the phosphopeptides (MSP2L) after enrichment using different additives to the loading buffer.

Additive Sample type 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9 2.10

80% ACNControl - - - - - - - - - -Wash - - - - - - - - - -Eluted 48 - 14 72 32 28 2 b 3 20 29

L-Glutamicacid

Control - - - - - - - - - -Wash - - - - - - - - - -Eluted 46 - 30 57 33 78 12 11 - 57

Citric acidControl - - - - - - - - - -Wash - - - - - - - - - -Eluted 42 - 26 54 29 51 27 24 - 52

Enrichment of phosphopeptides from a complex sample containing (1:50 | Phosphopeptides: BSA digest), performed on titanium dioxide material with different kind of additives in the loading buffer. The three different loading buffers contained either no additive (80% acetonitrile), 0.14 M L-glutamic acid, or 50 mM citric acid. The concentrations of the additives are derived from recent publications. The scores indicate the MASCOT ion scores for the identified phosphopeptides. b: Only observed with transfer of the phosphogroup to another position.


One of the most apparent aspects of these results is the recovery of almost all the phosphopeptides in all of the enrichment experiments (peptide 2.2 has never been detected). Additionally, no phosphopeptides are detected in control or washing fractions, which is to be expected. The most intriguing difference between the different additives is the reduction in protein score and protein coverage for BSA in the eluted fraction, in which L-glutamic acid has been used as additive to the loading buffer, see figure 10. Whereas both acetonitrile and citric acid retain approximately the same scoring and coverage, the coverage is reduced about 2-fold and the protein scoring seems four times smaller than control or washing fractions. This suggests that the addition of L-glutamic acid (an amino acid) is a useful tool for eliminating unwanted coordination of nonphosphorylated peptides to the MOAC material. However, since there are still nonphosphorylated peptides present in the eluted fraction, additional washing and clean up steps might be needed. Citric acid has been used in a quantity associated with monophosphorylated peptide enrichment, and might provide different results when higher concentrations are used.

Control

FT +

Wash Elu

ted0

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5000

7500

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40%

60%

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Figure 10. The protein scoring and coverage of Bovine Serum Albumin (BSA) after titanium oxide enrichment with different additives.For each enrichment experiment three samples were collected; not enriched negative control (Control), a pooled fraction of the flow through and wash fractions (FT + wash), and the eluted fraction (Eluted). Each filled bar represents the protein score (arbitrary units) of BSA on the left y-axis, each fading bar represents the protein coverage in percentages on the right y-axis. From left to right: Enrichment with 80% acetonitrile in the loading buffer; L-Glutamic acid (0.14M) as additive to the loading buffer; Citric acid (50 mM ) additive to loading buffer.

In contrast, the loading buffers containing acetonitrile and citric acid both have increased or similar scoring and coverage of BSA after enrichment on TiO2 material. This indicates that using these additives is not as effective as using L-glutamic acid at competitively displacing nonphosphorylated peptides from titanium. However, due to limited capacity of the MOAC columns, less nonphosphorylated peptides could be co-eluting and end up in the enriched sample. This “dilution” of nonphosphorylated peptides could explain why the phosphopeptides are still observed, whereas they could not be detected in the starting mixture, see “control” table 3. Multiple rounds of enrichment with these kinds of additives could perhaps obtain the same level of enrichment obtained by a single round of L-glutamic acid assisted enrichment. Another possibility is to combine multiple additives into one washing buffer, or develop a workflow with multiple different washing steps.

In order to observe the effect of concentration of the additives on enrichment properties, a series of experiments was performed on tryptic digested β-casein with different concentrations of both L-


glutamic acid and citric acid. The benefit of using β-casein over the phosphopeptide kit is that the tryptic digestion of this protein contains both phosphorylated and non-phosphorylated peptides, as can be seen in the supplemental information. Moreover, this specific β-casein (purchased from Sigma Aldrich, St. Louis, MO, USA) also contains κ-casein, α-S1-casein and α-S2-casein. These other casein proteins also contain phosphorylation sites, which makes them eligible for phosphopeptide enrichment. However, some of the obtained peptides have a long sequence and/or contain multiple phosphorylation sites of up to five sites, see supplemental information. These two facts render some of the possible phosphopeptides unfit for analysis using our specific setup. These large peptides require a charge of 4+ or higher to reduce the m/z ratio within the detection range of our ion trap mass spectrometer. For phosphopeptides, with their negatively charged phosphate groups, higher positive charges are very unfavorable. Moreover, obtaining such a charge requires multiple basic sites along the sequence, which is often not the case. Multiple phosphorylation sites are responsible for more negative charge, which makes it difficult to observe these kinds of peptides during mass spectrometry analysis. Additionally, multiple phosphorylation sites could result in tighter binding to the MOAC material, which could result in loss during elution. Because of these reasons, only the peptides from β-casein and α-S1-casein were analyzed during this experiment.

Using L-glutamic acid as an additive, three different concentrations were tested: 35 mM, 70 mM and 140 mM. The last concentration (140 mM) lies closely to the saturation limit in water for this compound. Although the buffers also contain a high percentage of organic solvent (acetonitrile, 65%) and acid (2% TFA), this saturation limit seems to still be near this point, because the acid did not dissolve readily. Also for the citric acid additive, three different concentrations were used: 50 mM, 0.5M and 1M. These concentrations span the range as is used during Citric Acid-Assisted Two-Step Enrichment (CATSET) protocols.48 The other buffers (washing, eluting, and resuspension buffers) were kept the same as used before.

The enriched samples were analyzed with CID neutral-loss-triggered ETD mass spectrometry and the protein scoring as well as the protein coverage was determined, see figure 11. Also the presence of the specific phosphopeptides was evaluated, in combination with the charge state of the specific ion; see both tables 4 and 5.

35mM 70mM 140mM0

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Figure 11. The protein scoring and coverage of β-casein from titanium oxide enrichment with different concentrations of additives to the loading buffer. For each enrichment experiment three samples were collected; not enriched negative control (Control), a pooled fraction of the flow through and wash fractions (FT + wash), and the eluted fraction (Eluted). Each filled bar represents the protein scoring on the left y-axis, each fading bar represents the protein coverage in percentages on the right y-axis. Left: Enrichment with L-glutamic acid as additive to the loading buffer (35 mM, 70 mM, 140 mM). Right: Enrichment with citric acid as additive to the loading buffer (50 mM, 0.5 M, 1 M).

Table 4. Identification of specific phosphopeptides from both β-casein and α-S1-casein after L-glutamic acid enhanced MOAC enrichment.

Conc. L-glutamic acid (mM)

β-casein α-S1-casein2060.8

22431.0

41659.8

41926.7

61950.9

4

35CID 2+ - 2+ - 2+

NL-ETD 2+ - 2+ 2+ 2+

70CID 2+ 3+ 2+ 2+ 2+ / 3+

NL-ETD 2+ 3+ 2+ 2+ 2+

140CID 2+ 2+ / 3+ 2+ 2+ 2+ / 3+

NL-ETD - 2+ / 3+ 2+ 2+ 2+

The observed ions (either 2+ or 3+ ions) from a selection of the phosphopeptides from both β-casein and α-S1-casein after enrichment with different concentrations of L-glutamic acid in the loading buffer. The specific mass of the observed peptides is shown underneath the protein name, see supplemental information for sequence. The enrichment was performed on titanium dioxide columns. Both the CID and Neutral-Loss-triggered ETD results have been listed per concentration.

Table 5. Identification of specific phosphopeptides from both β-casein and α-S1-casein after citric acid enhanced MOAC enrichment.

Conc. citric acid (M)

β-casein α-S1-casein2060.82 2431.04 1659.84 1926.76 1950.94

0.05CID 2+ 2+ / 3+ 2+ 2+ 2+ / 3+

NL-ETD 2+ 2+ 2+ 2+ 2+

0.5CID 2+ 2+ / 3+ 2+ - 2+

NL-ETD 2+ 3+ - - -

1CID 2+ 2+ / 3+ 2+ 2+ 2+

NL-ETD 2+ - - - -


The observed ions from a selection of the phosphopeptides from both β-casein and α-S1-casein after enrichment with different concentrations of citric acid in the loading buffer. The enrichment was performed on titanium dioxide columns. The specific mass of the peptides is stated underneath the protein names, for more sequence see supplemental information. Both the CID and Neutral-Loss-triggered ETD results have been listed per concentration.

Whereas the left graph in figure 11 shows a decline in protein scoring and coverage upon increasing the concentration of the additive, the same cannot be said for the enrichment using citric acid. The lowering of the scoring is the result of less non-phosphorylated in the eluted fraction, since fewer peptides means lower confidence in protein determination. The presence of the selected phosphopeptides in the samples from enrichment with 70 mM, shows that the most peptides can be observed when using this concentration of L-glutamic acid, see table 4. This is in contrast to the hypothesis that adding more additive to the loading buffer will result in better enrichment. However, the reason might be that adding to much of the additive will result in competitive binding with the phosphopeptides. This observation, together with the fact that protein coverage is very close to the coverage obtained when using 140 mM L-glutamic acid, suggests that enrichment with L-glutamic acid as an additive should be used with the second highest concentration (70 mM).

The enrichment with citric acid shows a different trend (figure 11 and table 5), no clear increase or decline is visible from there results. Additionally, when looking to the presence of the phosphopeptides in the different eluted fractions from these runs, roughly the same peptides can be observed when increasing the concentration of citric acid when analyzed with CID fragmentation. When NL-ETD fragmentation is used less peptides can be observed when the concentration is increased. This is in accordance with the CATSET method, where a higher concentration of citric acid is used to simultaneously clear the column of non- and monophosphorylated peptides, whilst retaining multiple phosphorylated peptides. However, almost none of the possible multiple phosphorylated peptides from β-casein, α-S1-casein, α-S2-casein or κ-casein are observed, as is also the case for the enrichment with L-glutamic acid. Most of these multiphosphorylated peptides contain 4 phosphorylation sites, which results in poor ionization during mass analysis, others might have a too long sequence for the mass range of the ion trap system used.

4.1.4 Elution optimization

Eluting the phosphopeptides from the column might not be completely effective, since we have observed loss of peptides without finding them back in the wash fraction. Although some of the peptides from the phosphopeptide kit were hardly detected even without enrichment, see materials and methods and supplemental information, this indicates loss on the column (incomplete elution) or severe adsorptive loss. However, due to the fact that other peptides are observed, this last option is much less likely to be of influence. In pursue of enhancing the elution, the pH of the buffer was increased to 12, up from 10.5. This pH has been shown to improve elution in other studies and has been used in other kinds of elution buffers.4, 5, 15, 16 To test this new elution buffer, a simple enrichment without additive was performed on the tryptic digest of β-casein with both elution buffers, see figure 12.


pH 10.5 pH 120

250

500

750

1000

0%

25%

50%

75%

100%

Figure 12. The protein score and coverage of β-casein after enrichment on TiO2 using different elution basicity’s. For both experiments the eluted fractions have been compared using the protein score and coverage of β-casein. Each filled bar represents the protein scoring on the left y-axis, each fading bar represents the protein coverage in percentages on the right y-axis. From left to right: Enrichment with NH4OH (2 M (~2% solution), pH 10.5 and pH 12.0 elution buffers, respectively.

When observing the scores and coverage from both experiments, a lowering of the protein score (~39% decline) and protein coverage (~8% decline) indicates a more successful removal of nonphosphorylated peptides when using a higher pH in the elution buffer. This conclusion is rather odd, due to the fact that the elution should not have effect on the removing of nonphosphorylated peptides. However, in both experiments exactly the same amount of sample was loaded and the exact same approach was followed except for the elution. A possible explanation would be that the elution with higher pH only affects those peptides that are bound with a phosphogroup to the titanium dioxide material and not the nonphosphorylated peptides. Additionally, no difference in observed phosphopeptides was detected (this data is not shown), indicating that the increase in pH does not affect the retaining of “sticky” phosphopeptides. It might be possible that a higher concentration of the ammonia solution (~5% solution), as well as other compounds (such as bis-Tris propane), might be more effective in eluting “sticky” phosphopeptides.

4.2 Identification techniques

4.2.1 CID/ETD fragmentation

Two different methods of fragmentation have been explored, CID and ETD. Additionally, the neutral-loss triggered switching of the modes has also been explored as a hybrid approach in which both the methods are combined (CID, NL-ETD). The specific equipment used (amaZon Speed / ETD MS) contains a small cartridge which is filled with fluoranthene, see figure 5. This chemical is used to create radical species in combination with the carrier gas, methane. As was explained previously in the materials and method section, this chemical ionization compound can also be used for a different reaction; the Proton Transfer Reaction (PTR). In order to minimize this inherent side reaction, a small assessment of the reaction time for the ETD reaction has been done for the used setup.


In order to evaluate the obtained data, scoring (ion and protein) was compared to scoring from standard CID runs. Additionally, spectra from both CID and ETD fragmentation have been selected from the most abundant peptide (2.6) for analysis.

It seems that using the neutral loss triggered ETD fragmentation yields the same level of detection of the peptides, see figure 6. Additionally, ETD-obtained data tends to yield more precise information about the location of the phosphorylation site than CID obtained data. Not all phosphopeptides have the tendency to experience neutral loss of their phosphategroup upon collision induced fragmentation, which is probably due to the proton mobility regarded with these peptides (this will be explained in depth later). Adjusting the energy during CID analysis might also prove valuable in optimizing neutral loss of phosphoric acid, as this reaction only requires little cleavage activation energy (<20 kcal/mol).65 The added time in the duty cycle of the mass spectrometer due to the ETD fragmentation does not seem to affect the score of the ions. Moreover, the added localization information gives additional confirmation on the identification of the peptides.

Table 6. Comparison of scoring and identification of phosphopeptides (MSP2L) using different fragmentation techniques for mass analysis.

Analysistype 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9 2.10

ETD 31 - 26 16 64 42 53 39 19 7CID 43 - 40 59 31 77 37 26 35 31

NL-ETD 26 - - - 52 34 34 17 11 -

Using phosphopeptide kit MSP2L (see supplemental information), several tests were performed in order to demonstrate the potential signals obtained from ETD and CID with Neutral-Loss-triggered ETD mass spectrometry. The scores represent the ion scores of the phosphopeptides. These samples have not been enriched.

After observing the potential of the neutral loss triggered ETD analysis, different approaches of enrichment of complex samples were analyzed using this mode, see table 7.

Table 7. Comparison of scoring and identification of phosphopeptides (MSP2L) after enrichment from a complex mixture with- and without additive enhancement, using different fragmentation techniques.

Additive Sample type Fragmentation 2.1 2.2 2.3 2.4 2.

5 2.6 2.7 2.8 2.

9 2.10

80% Acetonitrile

ControlCID - - - - - - - - - -

CID (NL) 27 - - 58 - 37 - - - -NL-ETD 30 - - - - - - - - -

CID 48 - 14 72 32 28 2 b 3 20 29


ElutedCID (NL) 43 - 21 54 31 49 - 24 - 46NL-ETD 22 - - - 13 32 13 22 - -

L-Glutamic acid

(0.14M)

ControlCID - - - - - - - - - -

CID (NL) - - - - - 26 - - - 10NL-ETD - - - - - - - - - -

ElutedCID 46 - 30 57 33 78 12 11 - 57

CID (NL) 47 - 29 57 - 82 33 20 - 19NL-ETD 23 - - - - 35 60 17 - -

Citric acid(50mM)

ControlCID - - - - - - - - - -

CID (NL) - - - 42 - 48 - - - 9NL-ETD - - - - - - - - - -

ElutedCID 42 - 26 54 29 51 27 24 - 52

CID (NL) 51 - 41 63 35 90 - - - 39NL-ETD 31 - - - - 30 62 20 - -

Comparison between normal CID data and data obtained with CID with neutral loss triggered ETD. The samples contain a complex mixture (1:50 | Phosphopeptides : BSA digest) and have been enriched using TiO2. Per sample there are three lanes: Upper: CID obtained data; Middle: CID data from the CID NL-ETD experiment; Lower: the ETD data recorded after detection of neutral loss of a phosphoric acid during CID fragmentation. Per sample a different color has been used to visualize the groups. b: Only observed with transfer of the phosphate group to another position.

It seems that the same level of identification can be obtained when using CID with neutral-loss-triggered ETD experiments as when a normal CID experiment is performed. This shows that the increase in duty cycle does not negatively affect the ability to detect our peptides. However, not all the potential observable peptides are detected, as can be seen when comparing table 6 and 7 to each other. From table 6, it is also evident that running an ETD fragmentation experiment (without the neutral loss triggering) is comparable to a CID experiment in regard to the phosphopeptides from a commercial kit (MSP2L), although the scores are different. The results from table 6 are obtained from a simple sample, containing only the phosphopeptide kit. When analyzing a more complex mixture the results become rather different, as is visible in table 7. With this experiment, the ability to detect the phosphopeptides before (control rows in table 7) and after enrichment (eluted rows in table 7) with neutral loss triggered ETD was explored. It can be seen that the detection is less effective than when a simple sample is used. Not every identification with CID analysis yields a subsequent round of ETD analysis, and vice versa. When using the neutral loss of phosphoric acid to start a ETD analysis, it would be advantageous to optimize the conditions for this reaction to take place. The last two rows of table 7 indicate the kind of analysis that is sought, where CID analysis of the peptides 2.7 and 2.8 does not yield identification, but trigger an ETD analysis through loss of phosphoric acid. Unfortunately, this behavior is not observed for all the peptides, and indicates the need of further optimization of the applied energy to experience more neutral loss.

In pursuit of optimizing the ETD analysis an evaluation of the effect of the reaction time of the ETD was performed using tryptic digest of the phosphoprotein β-casein. Using the PeptideMass tool from ExPASy, an in silico digestion was performed on β-casein (trypsin, 1 missed cleavage allowed).85 This simulation showed that the protein contains five different phosphosites on serine residues and yields six different phosphopeptides, as can also be seen in the supplemental information. The protein scoring and coverage of the β-casein protein was determined upon fragmentation using three different ETD reaction times: 75, 100 and 125 ms, as can be seen in figure 13.


75 ms 100 ms 125 ms0

50

100

150

200

0%

25%

50%

75%

100%

ETD reaction time

Figure 13. The effect of different reaction times during ETD mass analysis on MASCOT protein score and protein coverage of β-casein.The results above are obtained after varying the reaction time in the ion trap for the ETD reaction. The standard setting for this parameter is 100 ms. On the left y-axis the protein score (arbitrary units) is shown, whilst the right y-axis indicates the protein coverage (%).

It seems clear that the scoring decreases slightly, but the coverage stays constant. This might be due to the fact that the spectra themselves did not change overly much when longer or shorter ETD fragmentation time was used. The main aspect changing was the intensity of the peaks in the spectra, which was reduced in the longer ETD experiments. However, this does mean that the relative intensities of the peaks increases, making it possible to assign more fragments from very low intensity signals previously not detected. A shorter ETD reaction time leads to incomplete ETD reactions, whereas a longer ETD reaction time probably leads to the loss of charge on the singly charged peptides. This would lead to a lowering of the intensity, as well as the quality of the fragmentation spectra. This effect was apparent when comparing the spectra obtained from the different reaction times of one of the phosphopeptides (FQpSEEQQQTEDELQDK). Mainly, the intensity of the spectra dropped, but retained the same shape. In the first two runs (75 and 100 ms) only the spectrum of the 2+ peptide were observed, but in the 125 ms run the 3+ peptide was also identified, see figure 14.


Figure 14. The mass spectra for the 2+ and 3+ ions of the phosphopeptide FQpSEEQQQTEDELQDK after Electron Transfer Dissociation with a reaction time of 125 ms. This peptide originates from tryptic digested β-casein, see supplemental information. For these experiments the reaction time in the ion trap was set to three different times: 75, 100, and 125 ms. The standard setting for the ETD reaction is 100 ms. Upper: mass spectrum of the 2+ ion; Lower: mass spectrum of the 3+ ion.

4.2.2 Proton mobility

The presented analyses also reveal which of the peptides are prone to undergo neutral loss of the phosphategroup during CID fragmentation, which is related to proton mobility. Palumbo et al (2008)74

have shown the strong inverse correlation between the abundance of product ions obtained after neutral loss of phosphoric acid and the proton mobility of the peptide. The effect of proton mobility on the fragmentation of peptides and phosphopeptides has been described by Palumbo et al (2011) as cited below:

"Fragmentation of most peptide amide backbone bonds is generally agreed to require the involvement of a proton at the cleavage site, that is, that the cleavages are "charge-directed”. However, when the ionizing proton(s) are "sequestered" at the side chains of arginine, lysine, or histidine amino acids (i.e. the most basic site(s) within the peptide), more energy may be required to "mobilize" the proton from the basic side chains to the peptide backbone to induce "charge-directed" dissociation, compared to


that required to initiate "charge remote" fragmentation pathways (i.e. where the proton is not involved in the fragmentation mechanism). As a result, the extent of proton mobility within a protonated peptide ion dictates the type and abundance of the product ions that are formed. Three subcategories of proton mobility have been defined (Kapp et al., 2003). Precursor peptides are termed "non-mobile" when the number of ionizing protons is less than or equal to the number of arginine residues; "partially mobile" when the number of ionizing protons is greater than the number of arginine’s but less than or equal to the sum of basic residues (i.e. arginine, histidine and lysine residues); or "mobile" when the number of ionizing protons is greater than the sum of basic residues. For phosphorylated peptides, the abundance of product ions corresponding to the neutral loss of 98 Da have been shown to exhibit a strong inverse correlation with the proton mobility (non-mobile >> partially mobile > mobile; Palumbo, Tepe & Reid, 2008)." 74

To demonstrate the effect of proton mobility and relate it to the observations, a classification of the peptides used in this research was performed using: the peptide sequence; the definitions of the three proton mobilities, from Kapp et al (2003); a simple logical-test in excel. In excel a “logical test” function can be used to check whether or not a certain condition holds true. For the categorization, a triple logical test was performed for some charge states (1+, 2+, and 3+) which places the peptide into a category for that specific charge state. The logical test used for the characterization of the peptides used the following formula:

=IF(“charge state” <= “number of arginine residues”, "Non-mobile", IF(“charge state” <= “sum of basic residues”, "Partially mobile", IF(“charge state” > “sum of basic residues”, "Mobile", "Error")))

The red part shows the first logical test and terms the peptide “non-mobile” if the number of ionizing protons is less than or equal to the number of arginine residues. If this is not true, the next logical test is started. The orange part shows the second logical test, which tests if the peptides is “partially mobile”. This is done by checking if the number of ionizing protons is less than or equal to the sum of the basic residues. If the second logic test also proves false, the last test is started. The green part shows this last logic test and represents the “mobile” category. Here it is tested if the number of ionizing protons is larger than the sum of basic residues, which is always the case if the last two tests failed. Just in case of human error when setting up the spreadsheet, a category “error” was set up in case this last test also proves false. By definition, peptides should always fall in one of the proton mobility categories.Table 8. Classification of the proton mobility of the peptides used in this research.

MSP2L Proton Mobility

peptide FASTA abbrev. #Phospho (M+H)+ (M+2H)2+ (M+3H)3+

LPQEpTAR 2.1 1 Non-mobile Mobile Mobile

RYpSpSRSR 2.2 2 Non-mobile Non-mobile Non-mobile

EpTQSPEQVK 2.3 1 Partially mobile Mobile Mobile

VIEDNEpYTAR 2.4 1 Non-mobile Mobile Mobile

pSRSPpSSPELNNK 2.5 2 Non-mobile Partially mobile Mobile

ADEPSSEEpSDLEIDK 2.6 1 Partially mobile Mobile Mobile

HQYSDYDpYHSSpSEK 2.7 2 Partially mobile Partially mobile Partially mobile

NTPpSQHSHpSIQHSPER 2.8 2 Non-mobile Partially mobile Partially mobile

ELpSNpSPLRENSFGSPLEFR 2.9 2 Non-mobile Non-mobile Mobile

LGPGRPLPTF 2.10 1 Non-mobile Non-mobile Mobile


PpTSE(CAM)TSDVEPDTR

β-Casein Proton Mobility

peptide [M] #Phospho (M+H)+ (M+2H)2+ (M+3H)3+

FQpSEEQQQTEDELQDK 2060.84 1 Partially mobile Mobile Mobile

IEKFQpSEEQQQTEDELQDK 2431.06 1 Partially mobile Partially mobile Mobile

ELEELNVPGEIVEpSLpSpSpSEESITR 2965.21 4 Non-mobile Partially mobile Mobile

RELEELNVPGEIVEpSLpSpSpSEESITR 3121.31 4 Non-mobile Non-mobile Mobile

ELEELNVPGEIVEpSLpSpSpSEESITRINK 3220.43 4 Non-mobile Partially mobile Mobile

FQpSEEQQQTEDELQDKIHPFAQTQSLVYPFPGPIPNSLPQNIPPLTQTPVVVPPFLQPEVMGVSK

7359.67 1 Partially mobile Partially mobile

Partially mobile

The proton mobility of the peptides from the commercial kit (MSP2L) and some of the peptides from β-casein as obtained after tryptic digestion, see supplemental information. The upper table indicates the MSP2L peptides and shows the sequence, FASTA abbreviation and the proton mobility for the 1+, 2+ and 3+ charge states of the peptide. The lower table is concerned with a selection of the tryptic peptides from β-casein. The nominal mass (in Da) of these peptides has been given instead of a FASTA abbreviation.

Indeed we see the potential NL-ETD signals mainly occurring at the partially- and non-mobile peptides, as can be seen when comparing the data from table 7 and table 8 (MSP2L data). The three categories as stated above, might give insight into why certain peptides are easily fragmented, whereas others might not be susceptible to fragmentation at all. For example, the 2.2 peptide from the kit (with sequence: RYpSpSRSR) is found to be “non-mobile” even when triply protonated and has never detected in our experiments. This might also be why the peptides 2.7 and 2.8 are found with low ion scores or not at all (both are “partially mobile” peptides). However, peptide 2.9 has shown to be difficult to detect even when no interfering peptides were added (table 1). This peptide has the classification of “non-mobile” only for the 1+ and 2+ charge state, whereas the 3+ charge state is a “mobile” peptide. Although these peptides were produced from synthetic phosphopeptides, they do represent a tryptic digest of peptides found in mammalian HeLa cells. This is probably why they carry a 2+ charge state (peptides from tryptic digest are usually charged 2+), making it unlikely for peptide 2.9 to be observed. Additionally, proton mobility also plays a role in the possible transfer of phosphate groups within a peptide (non-mobile/partially mobile have higher tendency to facilitate transfer). 74

5. Conclusion

Analysis of phosphorylated proteins is a challenging task when using mass spectrometry due to the low concentrations in samples and suppressed signals during mass spectrometric analysis. Difficulties are also encountered throughout the isolation process and purification of the protein, as well as during data processing. Therefore, the aim of this research project is to propose a standard analysis workflow for the analysis of phosphorylated proteins for the department of Mass Spectrometry of Biomolecules at SILS (UvA). Optimization and implementation of a standard workflow was realized by using the available resources as well as extensive literature study for protocol suggestions. This resulted in the exploration of combining commercially available MOAC enrichment tips with L-glutamic acid as an additive. For mass spectrometric analysis of phosphopeptides the use of the amaZon Speed / ETD (Bruker) mass spectrometer was investigated for neutral loss triggered ETD, as well as regular ETD and/or CID analysis.


A generalized workflow, as discussed in the introduction (figure 1 and 2), sums up the following important aspects of phosphoproteomic research: sample preparation, phosphopeptide enrichment and purification steps, analysis with mass spectrometry, and data analysis. The obtained conclusions regarding each of these aspects are used to propose a standard protocol, which can be used by the Mass Spectrometry group.

When addressing sample preparation, it is proposed to use the common enzymatic digestion with trypsin, since this is a well-studied and developed method. Moreover, the peptides obtained by digestion with trypsin yield a charge-state suitable (≥2+) for the MOAC enrichment and subsequent ESI-MS/MS analysis. Other proteolytic enzymes could be explored for the ability of obtaining even higher charge states, or longer peptides. However, I think this should not be the main focus of upcoming research since this would mainly be important when analyzing multiple phosphorylated peptides. The existing digestion protocols used by the SILS research group are suitable for preliminary phosphoproteomic studies.

After obtaining peptides from the phosphorylated protein, enrichment of phosphopeptides is the next step. Towards this goal, different optimization experiments have been performed from which the following conclusions can be extracted. Of the obtained MOAC columns (TopTip, Glygen) which have been used during this study, TiO2 proved to be most effective material for phosphopeptide enrichment. When reviewing recent literature and observing the found results, it is quite possible that the zirconium-based MOAC columns might prove more useful when a different supplier or protocol is used. However, for this workflow proposal, the use of TiO2 columns as obtained from Glygen (Columbia, MD, USA) is recommended. Depending on the complexity of the sample (high or low concentration of nonphosphorylated peptides) a preliminary approach of non-acid-enhanced TiO2 enrichment should be tried. If the enrichment of the phosphorylated peptides is incomplete, additives such as glutamic acid (70 mM) should be added to the loading buffer. It is important to note that not all additives can be used in combination with all systems, since they might damage the equipment. When exploring new possible additives, make sure these chemicals are completely removed before mass spectrometric analysis by either desalting steps or evaporation.

Elution of the peptides from the MOAC material should always be performed with a high pH (~12) to make sure that the retained peptides are released. Higher concentrations of the ammonia solution might prove more beneficial as well as the use of different chemicals, such as bis-Tris propane.

Taking in account all the previous statements, the following protocol is proposed for phosphopeptide enrichment by the SILS Mass Spectrometry group at the UvA, see scheme 8:

Proposed protocol for phosphopeptide analysis using TiO2 MOAC columns:1. Phosphopeptide exploration. If the sequence is known for the phosphopetide, a spreadsheet should be made to determine the

proton mobility for the expected charge states (i.e. 2+, 3+). This will provide information about the

probability for the peptide to experience neutral loss of phosphoric acid. If this probability is high

(non-mobile / partially mobile) neutral-loss-triggered ETD analysis might prove useful.


2. Phosphopeptide enrichment with MOAC (TiO 2).If the sample is not to complex and/or behavior of the peptide upon analysis is unknown, use this

non-additive-enhanced protocol, otherwise go to the next protocol. Prepare the following buffers:

Loading buffer: 80% acetonitrile (ACN), 2% trifluoroacetic acid (TFA), dissolved in water.

Washing buffer: 65% ACN, 2% TFA, in H2O.

Elution buffer: 2% solution of NH4OH in water, pH 12.

Follow these steps:

Dilute the sample in loading buffer with and end volume of ~50 μL.*1 Keep in mind that you

will split this diluted sample in two fractions: one for enrichment, one for control.

Gently tap the TopTip column on a flat surface so that the material settles at the bottom.

Remove the red cap on the top and hold the column over an Eppendorf container.

Activate the material with 10 μL of the loading buffer (without the sample) and use a syringe

to push the liquid through. (repeat 3 times) Switch the column to a new Eppendorf marked as the Flow through + wash sample.

Load half (~25 μL) of the prepared sample on the column. Store the rest of the sample as

Control. The flow through should be loaded again in order to make sure that all phosphopeptides are

bound to the material.

Wash the column with 10 μL of the loading buffer (3x). Wash the column with 10 μL of the washing buffer (3x). Switch the column to a new Eppendorf marked as the Eluted sample.

Elute the sample with 10 μL of elution buffer (3x). Quickly acidify the eluted fraction with 30 μL of 10% ACN solution.

If the sample hasn’t been desalted beforehand, choose a tip with appropriate desalting

capacity, and desalt.

Afterwards, freeze dry the sample and dissolve in a 2% ACN, 0.1% TFA solution. Choose a

volume which makes the concentration equal to the concentration you have in the control

sample.

Store the samples at -20° C until analysis (snap freeze with liquid nitrogen before storage).

3. Additive enhanced enrichment with MOAC (TiO 2).When the sample is to complex, or preliminary enrichment failed, make use of this additive

enhanced TiO2 enrichment. Prepare the following buffers:

Loading buffer: 70 mM L-glutamic acid, 65% acetonitrile (ACN), 2% trifluoroacetic acid (TFA),

dissolved in water.

Washing buffer: 65% ACN, 2% TFA, in H2O.

Elution buffer: 2% solution of NH4OH in water, pH 12.


Follow the steps as stated at protocol: 2. Phosphopeptide enrichment with MOAC (TiO2). Always remove the additives before mass analysis, either by evaporation or desalting.

4. Mass analysis of the enriched phosphopeptides. Depending on the complexity of the sample and the proton mobility, an explorative analysis with

CID should be performed. However, when it becomes evident that the peptide will experience

neutral loss of phosphoric acid, NL-ETD should be utilized. Further optimization of this analysis

technique with regard to triggering the neutral loss of phosphoric acid, will provide an even more

valuable tool.

5. Data analysis It is advised to use the in house MASCOT client, since this server can process larger data files

than the online variant. Depending on the sample, choose an appropriate database to search

against (swissprot is almost always a good database to use). Use the standard settings as given

in the Materials & methods section, or change settings after discussion with the technician.

Scheme 8. The proposed protocols for analysis and enrichment of phosphopeptides TiO2 MOAC. This collection of protocols shows the general approach for phosphoproteomic research as suggested for use by the SILS Mass Spectrometry group at the UvA. *1: Depending on the concentration (if known), prepare a concentration suitable for mass analysis (intensity ~10^8), discuss with the technician.

The experiments in this research project show that both CID and ETD can be used to identify the same peptides from an enriched sample. Electron Transfer Dissociation does give more precise knowledge about the location of the phosphorylation, due to the tendency to leave the modification intact. Using both fragmentation methods simultaneously in a neutral-loss-triggered setup (CID NL-ETD) gives the most information in a single run. The experiments performed showed the necessity to optimize the conditions for neutral loss of phosphoric acid, as this does not yet seems optimized. Using a slightly lower energy during the CID part of the CID NL-ETD analysis, might increase the occurrence of neutral loss reactions. In order to use ETD, the equipment requires the use of fluoranthene. The setup of the amaZon Speed / ETD mass spectrometer makes this possible, as well as the possibility for PTR analysis. Investigating the optimization of the reaction time of ETD, proved that the default setting of 100 ms allows the best conditions for ETD analysis (with very little PTR side reactions). For further exploration of the mass spectrometric analysis, a combination of ETD assisted with CID (Electron Transfer collision activated Dissociation, ETcaD) might prove an effective analysis method. Another possible valuable tool for studying multiple phosphorylated peptides might be the “negative-mode” analysis, which monitors negatively charged ions.

In order to predict whether or not the NL-ETD runs are suitable for a specific peptide, the proton mobility should be determined as was stated in protocol 1. In this research, the proton mobility of the peptides of a commercial synthetic phosphopeptide kit (MSP2L), and tryptic digestion of β-casein were determined. The formula used for the logical tests has been given so that other peptides may be categorized rapidly in a spreadsheet. Interestingly, the results from this study show the same inverse


correlation between proton mobility and neutral loss product ion abundance as was shown by Palumbo et al (2008).74

Both the database and the software have an extensive selection of available software packages to choose from. Therefore, the usage of the standard in-house software is recommended. Before subjecting the results to database searching, it is advisable to check if there are specific databases containing your phosphopeptide. For example, the use of a contamination database for β-casein analysis during optimization experiments, decreased calculation time drastically without compromising the scoring. However, keep in mind that reducing the amount of annotated spectra against which you search might safe time, but give rise to the possibility of introducing false positive results. Additionally, any contaminants in the sample might go unnoticed. In short, when optimizing parameters whilst using a well-known analyte, it should not matter to much if you choose a smaller database.

Further research is always needed in order to perfect the choice for an appropriate workflow design. The main focus for further optimization of this proposed workflow should be on the testing different metals for MOAC enrichment in combination with different additives. Although an extensive literature study was performed prior to the selection of additives, certain other combinations might result in significantly enhanced purification of phosphopeptides. Moreover, new discoveries of other MOAC materials or properties of possible additives might propose more candidates for optimization of the proposed enrichment workflow. Besides looking at optimizing the MOAC enrichment route, alternative enrichment mechanisms – other than metal-affinity-based techniques – could be investigated. Utilizing new enrichment methods would enable the use of two-dimensional, orthogonal purification methods, when combined with the proposed workflow. This would result in even more specific enrichment and isolation of target phosphorylated proteins, as well as the possibility to separate the phosphopeptides into different groups (i.e. mono-, VS multiple phosphorylated peptides).

Extensive database search tools and specific software have already been developed, but should not be considered a finished field. Choosing the database and software is the most impactful decision, since most operate in similar fashion. Therefore, the focus should be on optimizing data acquisition and data processing. Automating the acquisition in such a way that the obtained spectra are directly processed, saves time and reduces the chance on human error. This kind of processes can be realized using scripts. Many different applications already exist for the annotation of the spectra obtained from phosphopeptides, but these have not been utilized during this research. Although most of these applications use a specific data format, it might prove worthwhile to find a suitable application for the obtained data from the amaZon Speed / ETD mass spectrometer.

Further standardization of the analysis of phosphorylated peptides could result in a higher number of comparable results and in more convenient research methods. This would result in the evaluation of published results becoming not only less laborious but would also reduce the length of time spent on optimizing methods. Hopefully, this research project will contribute towards the development of such


an approach for the analysis of phosphopeptides for the SILS Mass Spectrometry group at the University of Amsterdam.

Acknowledgments

First, I would like to thank prof. dr. C. G. de Koster and dr. L. J. de Koning for providing me with the opportunity to perform my master research project at their group. The confidence and trust that I would be able to do this project helped me a lot along the way. Secondly, I would like to thank Henk Dekker for his immeasurable patience and wise words which pulled me through the rough bits. Of course I would like to thank the rest of the group, whom have helped me and were very kind to me during the project. Lastly, I can’t thank my girlfriend, family and friends enough for the support they’ve provided me during this period.

6. References1. Jawon Seo, and Kong-Joo Lee, Post-translational Modifications and Their Biological Functions:

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Supplemental Information


http://web.expasy.org/peptide_mass/

Figure S1. Extracted ion chromatogram (EIC) of the ten different phosphopeptides from the MSP2L kit.

In this extracted ion chromatogram (EIC) the ten chromatographic traces of the phosphopeptides of the MSP2L kit are shown. These peptides have been shown using either the 2+ or 3+ ion, depending on the intensity and the appropriate retention time. Not all of the peptides are readily detectable due to the low intensity at which they appear, see the intensities for each peptide at the left y-axis. The low intensity of the peptides 2.2, 2.9, and 2.10 proved to be indicative of the level of detection in the experiments, since detecting these peptides was difficult.


Figure S2. The chromatographic overview of the phosphopeptide kit as provided by the manufacturer (Sigma Aldrich, St. Louis, MO, USA).

Here is shown the relative retention times as well as the relative abundance of the peptides as obtained in the kit; PhosphoMix 2 (MSP2L). The 2.2 peptide was difficult to detect upon MS analysis, as becomes evident from the intensity of the peptide in this overview.

Table S1. Composition of PhosphoMix Products (Sigma Aldrich, St. Louis, MO, USA).

This is the detailed information about the synthetic phosphopeptide kit used in this research; PhosphoMix 2 (MSP2L).

* Amino acid in [brackets] denotes site of label incorporation for heavy mixes as follows:[K],13C6 15N2 [R], 13C6 15N4 [V], 13C5 15N1 [L],13C6 15N1 [I], 13C6 15N1

(CAM) denotes carbamidomethyl cysteine** A FASTA file with all of the phosphopeptide sequences in the PhosphoMix product line is

available for free download on the product display page at sigma.com/phosphomix.*** As determined using electrospray ionization following standard reverse phase

chromatography


Table S2. Accurate mass to charge (m/z) values for peptides within the PhosphoMix Products (Sigma Aldrich, St. Louis, MO, USA).

Here is shown the detailed mass to charge ratios ([M], [M+H]+1, [M+2H]+2, [M+3H]+3) of the synthetic phosphopeptide kit used in this research. Only the light isotope kit was used in this study (only the data containing “light” in the FASTA abbreviation).


Table S3. Overview of the peptides obtained after In silico digestion of β-casein and α-S1-casein as performed with the PeptideMass tool from ExPASy.

β-casein

Modifications Positions

Peptides containing the

phosphorylation on position:

Charge state

1+ 2+ 3+

Signal peptide 1-15 - - - -

Phosphorylation (s)30, 32,

33, 34, 50

30, 32-343321.44 1661.22 1107.813122.32 1561.66 1041.442966.21 1483.60 989.41

507359.67 3680.35 2453.902432.06 1216.54 811.362061.84 1031.42 687.95

α-S1-casein

Modifications Positions

Peptides containing the

phosphorylation on position:

Charge state

1+ 2+ 3+

Signal peptide 1-15 - - - -

Phosphorylation (s)

56, 61, 63, 79, 81, 82, 83, 90,

130

56769.36 385.18 257.13

1026.50 513.75 342.8456, 61, 63 2678.06 1339.53 893.36

61, 63 1927.72 964.36 643.2461, 63, 79,81-83, 90

4629.68 2315.34 1543.90

79, 81-83, 902720.98 1360.99 907.673227.28 1614.14 1076.43

1302257.11 1129.06 753.041951.96 976.48 651.331660.81 830.91 554.28

In this table only the phosphopeptides are shown as obtained after performing an in silico digestion of β-casein and α-S1-casein. For this simulation, the PeptideMass tool from ExPASy 85 has been used with the following settings:

Protein digested: o CASB_BOVIN (for β-casein)o CASA1_BOVIN (for α-S1-casein)

Enzyme used: Trypsin. Maximum number of missed cleavages: 1. All cysteine residues in reduced form. Methionine residues in oxidized form.


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