From the Department of Biochemistry and Molecular Biology, University of Southern Denmark, DK-5230 Odense, Denmark
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ABSTRACT |
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A wide variety of methods are known for characterization of phosphorylated proteins. The most widely used have been peptide sequencing using Edman degradation combined with 32P labeling. This method is well established and very robust but has several limitations. For example, in Edman degradation the peptides have to be separated before the analysis using liquid chromatography. This decreases the overall sensitivity and increases analysis time, and it is therefore not well suited for analysis of complex samples.
Recently a number of MS-based strategies have been developed that are relatively sensitive and in many cases easier to perform than Edman degradation with respect to handling complex mixtures (e.g. Ref. 3). The increased sensitivity is especially needed for low stoichiometric phosphorylation. However, presently none of these MS-based methods can individually provide a complete characterization of a phosphorylated protein. For the MS-based strategies, it is common that the phosphorylated protein is enzymatically degraded to peptides, which are subsequently analyzed by MS to detect a mass increment of 80 Da per phosphate group. Because sulfonation gives the same mass shift, this strategy is often combined with phosphatase treatment to specifically cleave off the phosphate group from the peptide. This mass shift can be monitored by MS as a loss of 80 Da. This differential peptide mass mapping can be combined with purification of peptides using microcolumns packed with material of increasing hydrophobicity (4). In MALDI-TOF MS operating in reflector ion mode, the loss of phosphoric acid in the gas phase is often detected from phosphorylated peptides as a poorly resolved peak originating from metastable fragmentation (5). The exact site of phosphorylation can often be localized using tandem MS; however, the loss of phosphoric acid upon CID is frequently observed as the major fragmentation pathway, and this may interfere with the interpretation due to inadequate fragmentation of the peptide backbone.
The phosphate group is believed to have an effect on the ionization of phosphorylated peptides in MS, resulting in decreased signal intensity for phosphorylated peptides in the presence of non-phosphorylated peptides (i.e. an ion suppression phenomenon). Matrix additives like ammonium citrate (6) or phosphoric acid (7) have been shown to enhance the relative abundance of phosphorylated peptides in the presence of non-phosphorylated peptides in MALDI MS.
To reduce the suppression of phosphorylated peptides caused by the presence of non-phosphorylated peptides, it is advantageous to prepurify the phosphorylated peptides, especially from complex peptide mixtures. Enrichment of phosphorylated peptides from peptide mixtures using IMAC is widely used (814). With this approach the negatively charged phosphorylated peptides are purified by their affinity to metal ions like Fe3+ or Ga3+. However, frequently non-phosphorylated peptides, including those containing multiple acidic residues, are also enriched by this method (15). Blocking the acidic residues by O-methyl esterification has been shown to enhance the specificity of the phosphopeptide binding (15). Nonetheless in our experience the yield of this derivatization is below 100%, which compromises the sensitivity of this procedure and increases the complexity of the sample. In addition, O-methyl esterification often causes a partial deamidation and subsequent methylation of Asn and Gln residues, and these byproducts complicate the MS analysis and data interpretation further (16). Furthermore this method requires evaporation of the aqueous solvent from the peptide sample prior to the addition of the esterification reagent, and this step is known to cause adsorptive losses resulting in decreased sensitivity (17, 18).
Chemical modification by ß-elimination and concurrent Michael addition has also been widely used for affinity purification and quantitation of phosphorylated peptides (e.g. Ref. 19). However, this strategy suffers from several shortcomings including lack of reproducibility and sensitivity and introduction of unwanted side reactions (20).
Recently a promising strategy was introduced by Pinkse et al. (21) where titanium dioxide (TiO2) was used as an alternative to IMAC for the selective enrichment of phosphorylated peptides prior to ESI liquid chromatography tandem MS. They used an on-line TiO2 precolumn coupled directly to a reversed-phase capillary column, and with this setup successful analysis of various phosphorylated peptides was achieved. However, the selectivity of this method was somewhat compromised by the detection of several acidic non-phosphorylated peptides that were also retained by their TiO2 column.
Here we present a new and improved procedure for using TiO2 microcolumns that significantly enhanced the binding selectivity of TiO2 toward phosphorylated peptides, thereby enabling phosphorylated peptide characterization from low femtomole level phosphorylated proteins. Phosphorylated peptides obtained from model proteins were used to optimize and test the procedure. In addition, the method was compared with the IMAC method.
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EXPERIMENTAL PROCEDURES |
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Model Proteins and Peptide Mixtures
Serum albumin (bovine), ß-lactoglobulin (bovine), carbonic anhydrase (bovine), ß-casein (bovine), -casein (bovine), and ovalbumin (chicken) were from Sigma. Each protein was dissolved in 50 mM ammonium bicarbonate, pH 7.8 and treated with trypsin (12%, w/w) at 37 °C for 12 h.
Peptide Mixture 1
Peptide mixture 1 contained peptides originating from a tryptic digestion of 0.5 pmol of commercial -casein.
Peptide Mixture 2
Peptide mixture 2 contained peptides originating from tryptic digestions of serum albumin, ß-lactoglobulin, carbonic anhydrase, ß-casein, -casein, and ovalbumin. Peptide mixture 2 (ratios 1:1, 1:10, and 1:50) refers to a mixture of peptides originating from a tryptic digestion of 0.5 pmol of the phosphorylated proteins (ß-casein,
-casein, and ovalbumin) and 0.5, 5, and 25 pmol of the non-phosphorylated peptides (serum albumin, ß-lactoglobulin, and carbonic anhydrase), respectively.
Purification of Phosphorylated Peptides Using TiO2 Microcolumns
TiO2 microcolumns with a length of 3 mm were packed in GELoader tips. A small plug of C8 material was stamped out of a 3M Empore C8 extraction disk using an HPLC syringe needle and placed at the constricted end of the GELoader tip. The C8 disk serves only as a frit to retain the titanium dioxide beads within the GELoader tip. Note that the solvent used for either washing or loading the sample onto the TiO2 microcolumn contains organic solvent (5080% CH3CN), which abrogates adsorption of peptides to the C8 material. The TiO2 beads were suspended in 80% acetonitrile, 0.1% TFA, and an aliquot of this suspension (depending on the size of the column) was loaded onto the GELoader tip. Gentle air pressure created by a plastic syringe was used to pack the column as described previously (22, 23).
The efficacy of four different procedures for selective binding of phosphorylated peptides was investigated. The first procedure (A) was adopted from the method published previously (21). In the following optimization (procedures AD), peptide mixture 1 was used. In procedure A, peptides were loaded onto TiO2 columns in 0.1 M acetic acid. The columns were washed with 20 µl of 80% acetonitrile, 0.1 M acetic acid, and the bound peptides were eluted with 3 µl of 250 mM ammonium bicarbonate, pH 9.0. An aliquot of the eluate (0.7 µl) was mixed with 0.3 µl of 2% TFA and 0.5 µl of DHB/PA matrix solution (DHB (20 g/liter) in 50% acetonitrile, 1% phosphoric acid) directly on the MALDI target. Procedure B was the same as procedure A with the exception that the peptides were eluted with NH4OH, pH 10.5. In procedure C, the peptides were loaded onto the TiO2 columns in 0.1% TFA, and the columns were washed first with 10 µl of the DHB solution (DHB (20 mg/ml) in 50% acetonitrile) and then with 10 µl of 80% acetonitrile, 0.1% TFA before the bound peptides were eluted using 3 µl of NH4OH, pH 10.5. In procedure D, the peptides were loaded onto the TiO2 column in DHB solutions of different concentrations (1350 mg/ml in 80% acetonitrile, 0.1%TFA). The columns were washed with 10 µl of the DHB solution and 20 µl of 80% acetonitrile, 0.1% TFA. The bound peptides were eluted using 3 µl of NH4OH, pH 10.5.
To investigate the selective adsorption of phosphorylated peptides relative to non-phosphorylated peptides a series of organic acid solutions were tested as loading/washing solvents. TiO2 microcolumns were loaded with peptide mixture 2 (ratio, 1:1). The peptides were loaded onto the TiO2 microcolumns in a 0.13 M solution of one of the following acids in 1:1 (v/v) H2O/CH3CN with 0.1% TFA: phosphoric acid, benzoic acid, cyclohexane-carboxylic acid, phthalic acid, salicylic acid, and 2,5-dihydroxybenzoic acid. In the case of acetic acid, TFA was omitted from the solution. After loading the peptides onto the TiO2 columns, the columns were washed with 10 µl of 50% acetonitrile, 0.1% TFA. Each column was eluted with 3 µl of NH4OH, pH 10.5. Each eluate was purified on a Poros Oligo R3 microcolumn (see below) prior to MALDI MS analysis. For LC-ESI-MSMS analysis the eluted peptides were purified by Poros Oligo R3 reversed-phase material and eluted by 50% acetonitrile, partly dried, and diluted to 13 µl in 0.5% acetic acid.
Desalting and Concentration of Eluted Peptides
Custom made chromatographic reversed-phase microcolumns used for desalting and concentration of peptides were prepared using GELoader tips as described in detail earlier (22, 23). The eluates from the TiO2 microcolumns were diluted in formic acid to a final concentration of 5% and applied onto Poros Oligo R3 microcolumns using gentle air pressure. The columns were washed with 20 µl of 0.1% TFA. The retained peptides were eluted using 0.5 µl of DHB/PA matrix solution directly onto the MALDI target.
IMAC
IMAC purification of phosphorylated peptides was performed according to Cole et al. (24) with minor changes. Briefly 40 µl of iron-coated PHOS-selectTM metal chelate beads (Sigma) were washed two times in 100 µl of washing/loading solution (0.25 M acetic acid, 30% acetonitrile) and resuspended in 40 µl of washing/loading solution. An aliquot of this solution (20 µl) was incubated with the peptide solution in a total volume of 40 µl of washing/loading solution for 30 min with constant rotating. After incubation, the solution was loaded onto a constricted GELoader tip, and a gentle air pressure was used to pack the beads. Subsequently the beads were washed extensively with the washing/loading solution. The bound peptides were eluted using 3 µl of NH4OH, pH 10.5, and desalted using Poros R3 microcolumns prior to MALDI MS analysis (as describe above).
MALDI-TOF MS
MALDI MS was performed using a Voyager STR mass spectrometer (PerSeptive Biosystems, Framingham, MA) equipped with delayed extraction or a MALDI Q-TOF mass spectrometer (Micromass, Manchester, UK). All spectra were obtained in positive reflector mode. Mass spectrometric data analysis was performed using either the MoverZ software (www.proteometrics.com) or the software MassLynx 3.5. Sequence analysis and peptide assignment were accomplished using the GPMAW software (welcome.to/gpmaw). For analysis of phosphorylated peptides, DHB (20g/liter) in 50% acetonitrile, 1% phosphoric acid was used as the matrix. Note that inclusion of 1% phosphoric acid in the MALDI matrix solution increases the relative abundance of multiphosphorylated peptides (7). Because some of the test proteins used in this study (- and ß-casein) include several multiphosphorylated peptides the analyses were all performed using 1% phosphoric acid in the matrix solution (DHB/PA).
Nanoflow LC-ESI-MSMS
LC-ESI-MSMS analysis was performed using a Q-TOF Ultima mass spectrometer (Waters/Micromass UK Ltd., Manchester, UK) utilizing automated data-dependent acquisition. A nanoflow HPLC system (Ultimate, Switchos2, Famos, LC Packings, Amsterdam, The Netherlands) was used for chromatographic separation of the peptide mixture prior to MS detection The peptides were concentrated and desalted on a precolumn (75-µm inner diameter, 360-µm outer diameter, Zorbax® SB-C18 3.5 µm (Agilent, Wilmington, DE)) and eluted at 200 nl/min by an increasing concentration of acetonitrile (2%/min gradient) onto an analytical column (50-µm inner diameter, 360-µm outer diameter, Zorbax SB-C18 3.5 µm (Agilent)). A MS-TOF survey spectrum was recorded for 1 s. The three most abundant ions present in the survey spectrum were automatically mass-selected and fragmented by collision-induced dissociation (4 s per MSMS spectrum). The MSMS data were converted to a pkl file format using the MassLynx 3.5 ProteinLynx software, and the resulting pkl file was searched against the NCBInr protein sequence data bases using an in-house Mascot server (version 1.8) (Matrix Sciences, London, UK).
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RESULTS AND DISCUSSION |
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After elution with 250 mM ammonium bicarbonate, pH 9.0, the same microcolumn was subsequently eluted with 3 µl of NH4OH, pH 10.5, and the MALDI MS analysis of 0.7 µl of this solution yielded very abundant phosphorylated peptides (Fig. 1C), thereby demonstrating that elution with pH 9 only releases a small fraction of the adsorbed phosphorylated peptides, whereas pH 10.5 elutes most of the bound phosphorylated peptides. Subsequent elution using higher pH values did not result in further improvement in the recovery of phosphorylated peptides from the TiO2 microcolumns.
In previous purification procedures for enrichment of phosphorylated peptides using both TiO2 and IMAC 0.10.25 M acetic acid (pH 2.72.9) has been used as the loading buffer. The reason for choosing this pH value in the loading is to ensure that the acidic residues in the peptides are neutral, whereas the pKa1 value of phosphoric acid is 1.8, and therefore the phosphate group will still have a negative charge at pH 2.9. However, a significant amount of non-phosphorylated acidic peptides binds to either IMAC or TiO2 under those conditions (15, 21). The substitution of an alkyl group onto an acidic phosphate oxygen atom increases the acidity, e.g. the pKa1 value of phosphoric acid decreases to 1.1 upon methylation (i.e. CH3OPO(OH)2) (25). Thus, we anticipate that the pKa1 value of the phosphate group also decreases when it is linked to a peptide. Therefore 0.1% TFA, which has a pH value of 1.9, was used in the following buffers for the purification of phosphorylated peptides using TiO2.
A TiO2 microcolumn was loaded with peptide mixture 1 in 0.1% TFA. After washing with 80% acetonitrile, 0.1% TFA, the phosphorylated peptides were eluted from the TiO2 with 3 µl of NH4OH, pH 10.5, and the MALDI MS analysis of 0.7 µl of this solution resulted in the MALDI spectrum shown in Fig. 1D. Here the intensity of the phosphorylated peptides increased relative to the non-phosphorylated peptides, indicating a more selective enrichment of phosphorylated peptides when 0.1% TFA was used as loading buffer. However, still a significant number of non-phosphorylated peptides were observed using this procedure.
Elution of phosphorylated peptides from IMAC material using the DHB matrix solution has previously been shown to increase the recovery of some phosphorylated peptides from this chromatographic material (26). Because the binding of phosphorylated peptides to TiO2 is attributed to its ion exchange properties (21) and is similar to the binding observed in IMAC experiments, we attempted to elute the phosphorylated peptides with the DHB matrix solution.
The enrichment of phosphorylated peptides from peptide mixture 1 using TiO2 loaded in 0.1% TFA followed by washing in 80% acetonitrile, 0.1% TFA and elution of the phosphorylated peptides with pH 10.5 is shown in Fig. 2 A. Peptides from peptide mixture 1 were loaded onto a new TiO2 microcolumn as above; however, after washing with 80% acetonitrile, 0.1% TFA, the peptides were eluted directly onto the MALDI target using DHB matrix solution (20 mg/ml in 50% acetonitrile, 0.1% TFA). Phosphoric acid (1%) was added after the elution, and the eluate was subsequently analyzed by MALDI MS (Fig. 2B). Only non-phosphorylated peptides were detected as illustrated by the insets in the figure where arrows indicate the expected masses of some of the phosphorylated peptides. Thus, all phosphorylated peptides were retained by the TiO2 resin after elution with the DHB matrix solution. Subsequent elution from the same column with NH4OH, pH 10.5, recovered a larger number of phosphorylated peptides (Fig. 2C, marked with asterisks) compared with the procedure where the column was only washed with 80% acetonitrile, 0.1% TFA (Fig. 2A). In addition, a surprisingly low abundance and number of non-phosphorylated peptides were observed (four in total). In IMAC, DHB is sufficient to displace the bound phosphorylated peptides, whereas the interaction between TiO2 and the phosphate group appears to be much stronger and cannot be dissociated by DHB. However, acidic non-phosphorylated peptides can be displaced from TiO2 by competitive binding of DHB thereby increasing the selective binding of phosphorylated peptides.
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The Effect of the DHB Concentration on Phosphopeptide Isolation from Simple Mixtures
The concentration of DHB in the loading buffer was found to have a large effect on the adsorption of non-phosphorylated peptides onto TiO2. A series of experiments were performed using peptide mixture 1 and different concentrations of DHB in the loading buffer. Peptides from peptide mixture 1 were loaded onto equal length TiO2 microcolumns in 0, 1, 10, and 20 mg/ml DHB (in 80% acetonitrile, 0.1% TFA), respectively. The MALDI mass spectra covering the mass range 10002000 and 23003100 Da obtained from the elution with NH4OH, pH 10.5, are shown in Fig. 3, AD. This figure shows that the number of non-phosphorylated peptides decreases with the increasing concentration of DHB. In the experiment where the peptides were loaded in 80% acetonitrile, 0.1% TFA alone, five signals from non-phosphorylated peptides were observed (Fig. 3A, shaded areas). Inclusion of as little as 1 mg/ml DHB significantly decreased the abundance of these peptides. A further increase in the concentration of DHB to 20 mg/ml completely abrogated the adsorption of these non-phosphorylated peptides to TiO2 as is evident by their absence in the mass spectra. Interestingly the multiphosphorylated peptides in the m/z 23003100 ranges are only observed at DHB concentrations above 10 mg/ml. This indicates a pronounced suppression of the ionization of multiphosphorylated peptides in the presence of non-phosphorylated peptides.
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The absolute abundance of the phosphorylated peptides was markedly increased when using the DHB as loading buffer compared with either acetic acid or TFA despite the same amount of starting material. The same observation was made in all the other experiments performed in this study. This indicates a more efficient ionization for phosphorylated peptides in the absence of non-phosphorylated peptides.
The Effect of the DHB Concentration on Phosphopeptide Isolation from Semicomplex Mixtures
The effect of the inclusion of DHB in the loading and washing procedure for complex mixtures was investigated using peptide mixture 2 (ratio, 1:1). Peptides from peptide mixture 2 (ratio, 1:1) were applied onto TiO2 microcolumns of the same length in 0, 10, 20, 50, 100, and 200 mg/ml DHB (in 80% acetonitrile, 0.1% TFA), respectively. The resulting MALDI peptide mass maps obtained from 0.7 µl of each of the elutions (performed with 3 µl of NH4OH, pH 10.5) is shown in Fig. 5, AF. The absence of DHB caused a high number of non-phosphorylated peptides to bind to the columns. The number of non-phosphorylated peptides decreased with the increasing concentration of DHB up to 200 mg/ml where no non-phosphorylated peptides were observed. In addition, the multiphosphorylated peptides were more clearly detected when a higher concentration of DHB was used, probably due to decreased ion suppression effect. This clearly indicates that when the sample is more complex a higher concentration of DHB is needed to exclude the binding of non-phosphorylated peptides. For very complex samples a DHB concentration of 300400 mg/ml (close to a saturated solution) is highly recommended.
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Investigating the Mechanism for the Selective Enrichment of Phosphorylated Peptides by TiO2
It is clear from the results presented above that the presence of DHB in the loading buffer dramatically enhances the selective retainment of phosphorylated peptides on TiO2. We attribute this effect to a competition for binding sites on TiO2 between non-phosphorylated peptides and DHB molecules. The large molar excess of DHB thus effectively competes with non-phosphorylated peptides for adsorption to the surface of TiO2, whereas phosphorylated peptide binding is virtually unaffected. To investigate the molecular determinants of DHB that are responsible for this intriguing effect we selected a number of benzoic acid derivatives as well as other acids and determined their effect on the selective adsorption of phosphorylated peptides from complex peptide mixtures. Peptides from peptide mixture 2 (ratio, 1:1) were applied onto TiO2 microcolumns in 50% acetonitrile containing one of the following acids: trifluoroacetic acid, acetic acid, phosphoric acid, benzoic acid, salicylic acid, cyclohexane-carboxylic acid, phthalic acid, and DHB. The bound peptides were eluted using NH4OH, pH 10.5 and desalted and concentrated on Poros Oligo R3 microcolumns prior to MALDI MS analysis. The phosphorylated peptide binding selectivity was evaluated by comparing the relative abundances of these peptides with those of non-phosphorylated peptides. Fig. 7 shows MALDI mass spectra obtained from TiO2 enrichment of phosphorylated peptides using four different acids in the loading buffer (TFA, phosphoric acid, benzoic acid, and DHB). The spectra clearly show that DHB was the most efficient acid to prevent adsorption of nonphosphorylated peptides while retaining the ability of TiO2 to bind phosphorylated peptides. In contrast, phosphoric acid was not as effective as DHB to reduce binding of nonphosphorylated peptides, and it appeared to inhibit the adsorption of some of the phosphorylated peptides. For example, the relative abundances of the phosphorylated peptides at m/z 2088.9, 1660.8, and 1466.7 were dramatically reduced when the loading buffer contained phosphoric acid, whereas the relative abundances of these peptides were rather similar in the case of the other acids. Using salicylic acid or phthalic acid in the loading buffer yielded spectra very similar to those obtained from DHB, whereas cyclohexane carboxylic acid gave results very close to that of benzoic acid (data not shown). The efficacy in inhibiting adsorption of nonphosphorylated peptides follows the order DHB salicylic acid
phthalic acid > benzoic acid
cyclohexane carboxylic acid > phosphoric acid > TFA > acetic acid. Thus, the substituted aromatic carboxylic acids (i.e. DHB, salicylic acid, and phthalic acid) are markedly better competitors than the monofunctional carboxylic acid (i.e. TFA, acetic acid, cyclohexane-carboxylic acid, and benzoic acid) for preventing binding of non-phosphorylated peptides to TiO2. In line with this observation, infrared spectroscopic studies have shown that substituted aromatic carboxylic acids (including salicylic acid and phthalic acid) coordinate strongly to the surface of TiO2, whereas monofunctional carboxylic acids (including benzoic acid and acetic acid) only interact very weakly with TiO2 (27). Interestingly phosphate binds to TiO2 with affinity (KA = 4 x 104 M1) similar to substituted aromatic carboxylic acids (KA = 104105 M1) (28), but it appears to be significantly less effective in preventing binding of nonphosphorylated peptides. Clearly a high TiO2 binding affinity of the acidic loading buffer is not the decisive factor for the enhancement of phosphorylated peptide binding selectivity. In this context, it is important to note that the binding mode of phosphate to TiO2 differs from that of substituted aromatic carboxylic acids. For example, the binding mode of salicylic acid to TiO2 is a chelating bidentate salicylate species (27, 29), whereas the adsorption of phosphate anions to the surface of TiO2 is a bridging bidentate surface complex (28) (see Fig. 8 ). The coordination geometry for an optimal phosphate binding site on TiO2 is thus likely to differ from that of an optimal binding site for a substituted carboxylic acid. In this picture, phosphate will compete directly with phosphorylated peptides for binding sites on TiO2, whereas DHB targets other binding sites that appear to be similar to those preferred by non-phosphorylated peptides. Such nonequivalent binding sites on TiO2 may result from a structural heterogeneity of the TiO2 surface, but the adsorbate itself is also likely to generate nonequivalent binding sites on TiO2 by the ability of TiO2 to transmit inductive electronic effects over a few rows of atoms (30). Furthermore steric hindrance does also appear to play a role in preventing binding of non-phosphorylated peptides because bulky monofunctional carboxylic acids (e.g. cyclohexane-carboxylic acid) are more effective than less bulky acids (e.g. acetic acid). In conclusion, we attribute the selective enhancement of phosphorylated peptide binding by DHB to an effective competition predominantly with non-phosphorylated peptides for binding sites on TiO2. This effect is achieved by the existence of a heterogeneous array of adsorption sites on TiO2.
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The mechanism of the selective enrichment of phosphorylated peptides by TiO2 in combination with DHB was investigated by using other substituted aromatic acids as well as other acids. We attribute the enhancement of phosphorylated peptide binding selectivity to an effective competition between DHB and non-phosphorylated peptides for binding sites on TiO2.
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FOOTNOTES |
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Published, MCP Papers in Press, April 27, 2005, DOI 10.1074/mcp.T500007-MCP200
1 The abbreviations used are: DHB, 2,5-dihydroxybenzoic acid; PA, phosphoric acid.
2 M. R. Larsen, T. E. Thingholm, O. N. Jensen, P. Roepstorff, and T. J. D. Jørgensen, unpublished results.
* This work was possible through a grant to the Danish Biotechnology Instrument Center (DABIC) from the Danish Research Agency.
Financed by the Carlsberg Foundation.
Financed through a Steno scholarship from The Danish Natural Science Research Council. To whom correspondence should be addressed: Dept. of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej 55, DK-5230 Odense M, Denmark. Tel.: 45-65502342; Fax: 45-6593-2661; E-mail: mrl{at}bmb.sdu.dk
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REFERENCES |
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