w. -V... ..,x--..... ‘.,.‘..,_ ., ‘ ‘ A "inn"-- This is to certify that the dissertation entitled The Determination of One-Carbon lntennediates in Plants and the Development of a Protocol for the Identification of i Pyridoxal-I-Phosphate Modified Residues in Proteins j presented by Eric Steven Simon has been accepted towards fulfillment of the requirements for the Ph.D. degree in Chemistry W: W ' Major Professor’s Signature 5.2/09 Date MSU is an Affirmative Action/Equal Opportunity Institution ""’"‘“fl LIBRARY—— Michigan State University PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date due. MAY BE RECALLED with earlier due date if requested. DATE DUE DATE DUE DATE DUE 2/05 c‘JCfiC/DatoOmJndd-nts The Determination of One-Carbon Metabolic Intermediates in Plants and the Development of a Protocol for the Identification of Pyridoxal-l-Phosphate Modified Residues in Proteins By Eric Steven Simon A Dissertation Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Chemistry 2004 ABSTRACT The Determination of One-Carbon Metabolic Intermediates in Plants and the Development of a Protocol for the Identification of Pyridoxal-l-Phosphate Modified Residues in Proteins By Eric Steven Simon Plants produce specialized molecules that are designed to protect them from specific environmental threats. These specialized molecules fall under the category of secondary metabolites. In addition to the physiological importance of secondary metabolites to plants, many of these molecules possess pharmaceutical and industrial value. This has sparked an interest in controlling the output of specific secondary metabolites in plants. The synthetic pathways for secondary metabolites require methyl groups from one-carbon metabolism in plants. However, relatively little is known about plant one-carbon metabolism that would allow scientists to engineer plants for greater yields of secondary metabolites. This dissertation discusses some efforts that have been made to study some of these intermediates and enzymes, particularly those that are involved in a poorly understood pathway in plant one-carbon metabolism, the S-methylmethionine (SMM) cycle. Presented here is the development of some strategies used to quantify key intermediates that are involved, or related, to the SMM cycle. These protocols were applied to wild type and mutant Arabidopsis thalz’ana plants. The mutant plant lacked S- methionine methyltransferase (MMT) activity. The established function of MMT is methylating methionine which effectively starts the SMM cycle. Measurements reported here of the intermediates involved in the SMM cycle in both the wild type and mutant plants imply a function of the SMM cycle in regulating the methylation ratio in plants. The methylation ratio is the relative concentration of S-adenosylmethionine (SAM) to S- adenosylhomocysteine (SAH), (SAM/SAH). SAM is the one-carbon metabolic intermediate responsible for the donation of methyl groups to primary and secondary metabolite synthesis, and SAH is the demethylated byproduct. A protocol was also developed to study the key enzyme initiating the SMM cycle, MMT. Sequence analysis of MMT suggests a possible association with pyridoxal-l— phosphate (PLP). Before this could be assessed, a protocol had to be established that could be applied to MMT in future studies. Tandem mass spectrometric studies were performed with PLP modified peptides and the MS/MS spectra reveal that these peptides fragment to yield two distinct products corresponding to the neutral losses of phosphoric acid, H3PO4, and PLP. Although no sequence information was available, a fragmentation pattern was established that could be used to identify MS/MS spectra that correspond to PLP modified peptides (by neutral loss analysis). This strategy was applied to two known PLP binding enzymes that were digested with enzymatic proteases. The peptide products were fractionated with liquid chromatography and analyzed with tandem mass spectrometry. Neutral loss analysis of the resulting MS/MS spectra highlighted those that displayed the characteristic neutral loss patterns of PLP modified peptides. The masses of these peptides matched the theoretical peptides from the digests that contained the previously determined PLP modified residue. ACKNOWLEDGEMENTS I want to thank Professor John Allison for his guidance and support throughout my graduate career. I especially want to thank him for his patience and confidence throughout a very challenging project that certainly tested the endurance of both of us. I’d like to also thank Professor Douglas Gage for introducing me to the biological sciences which turned out to be an unanticipated, but pleasant, surprise. I should also thank the staff of the Mass Spectrometry Facility for allowing me to work there independently. I want to give thanks to my family for all of their love and support. My parents have taught me many lifelong lessons that have contributed to my successful completion of this project. I would like to thank God for blessing me with such a fine family and many good friends. He has surrounded me with great people that have always supported me throughout all of my endeavors. There’s no doubt that I have benefited from the generosity of many people. Several events and outcomes had to turn in my favor for me to be able to succeed at Michigan State University. It would not have gone my way without the help of so many people. I thank you all. iv TABLE OF CONTENTS List of Tables ....................................................................................... List of Figures ....................................................................................... Chapter 1 Introduction and Background of Plant One-Carbon Metabolism. . 1.1 Introduction to Plant One-Carbon Metabolism ............................................ .2 Secondary Metabolites 1.3 Established F acts/Characteristics of One-Carbon Metabolism in Plants ............... 1.3.1 Sources of One-Carbon Units ......................................................... 1.3.2 The One-Carbon Groups and the Inter-conversion Reactions of the Folate Pool ............................................................................................. 1.3.3 The Activated Methyl Cycle ......................................................... 1.4 Unknown Aspects of Plant One-Carbon Metabolism .................................... 1.5 Introduction to the S-Methylmethionine (SMM) Cycle ................................. 1.6 Recent Findings in Regards to the S-Methylmethionine (SMM) Cycle ............... 1.7 Continuing the SMM Investigation ......................................................... 1.8 Chapter 1 References ......................................................................... Chapter 2 THE DETERMINATION OF S-METHYLMETHIONINE (SMM), S-ADENOSYLMETHIONINE (SAM), AND s- ADENOSYLHOMOCYSTEINE (SAH) 1N PLANTS .................... 2.1 Introduction .................................................................................... 2.2 Strategy for Establishing the Function of the S-Methylmethionine Cycle ............. 2.3 The Determination of S-Methylmethionine From Plant Tissue ......................... 2.3.1 Measurements of SMM with Mass Spectrometry ................................ 2.3.2 Procedure for the Extraction / Purification of S-Methylmethionine from Plant Tissue with Ion Exchange Chromatography ....................................... 2.3.3 Procedure for the Measurement of S-Methylmethionine Levels from Arabidopsis Extracts with Mass Spectrometry ........................................... 2.3.4 Results: S-Methylmethionine Levels in Wild Type and Mutant (—MMT) Arabidopsis thaliana ......................................................................... 2.4 Determination of S-adenosylmethionine (SAM) and S-adenosylhomocysteine (SAH) from Plant Tissue ...................................................................... 2.4.1 The Molecular Structures of S-Adenosylmethionine and S- Adesnosylhomocysteine .............................................................. 2.4.2 The Stability of S-Adenosylmethionine and S-Adenosylhomocysteine ....... 2.4.3 Previous Efforts to Quantify S-Adenosylmethionine and S- Adenosylhomocysteine from Biological Tissue .................................. 2.4.4 Phenylboronate Affinity to Isolate S-Adenosylmethionine and S- Adenosylhomocysteine .............................................................. ix 24 24 25 26 28 30 32 32 34 34 37 38 39 2.4.5 Procedure for the Extraction of S-Adenosylmethionine and S- Adenosylhomocysteine from Plant Tissue ......................................... 42 2.4.6 Separation and Quantification of S-Adenosylmethionine and S- Adenosylhomocysteine with High Performance Liquid Chromatography (HPLC) ................................................................................. 43 2.4.7 Results: S-Adenosylmethionine and S-Adenosylhomocysteine Levels in Wild Type and Mutant Arabidopsis thaliana ..................................... 44 2.5 Discussion ..................................................................................... 47 2.6 Conclusions .................................................................................... 51 2.7 Chapter 2 References ......................................................................... 53 Chapter 3 Background Information on Pyridoxal-l-Phosphate Binding Enzymes ............................................................................ 55 3.1 Introduction: Questions About Methionine Methyltransferase (MMT) ............... 55 3.2 Sequence Analysis of MMT ................................................................. 57 3.3 Enzymes That Bind Pyridoxal-l-Phosphate and Typical Reactions .................... 59 3.4 Reduction of Imines ........................................................................... 67 3.5 Previously Established Techniques for the Detection of Pyridoxal-l-Phosphate in Proteins ............................................................................................... 68 3.6 Approaches for Determining Pyridoxal-l—Phosphate Binding Sites in Proteins ...... 7] 3.7 Conclusion ..................................................................................... 77 3.8 Chapter 3 References ......................................................................... 78 Chapter 4 Summarizing the Advancements in Mass Spectrometry for Identifying Proteins, Sequencing Peptides, and Studying Post- Translational Modifications .................................................... 80 4.1 Introduction .................................................................................... 80 4.2 Introduction to Mass Spectrometry ......................................................... 82 4.3 Ion Sources ..................................................................................... 84 4.3.1 Matrix-Assisted Laser Desorption/Ionization (MALDI) ........................ 84 4.3.2 Electrospray Ionization (ESI) ......................................................... 85 4.4 Mass Analyzers ................................................................................ 88 4.4.1 Time-of—Flight (TOF) ................................................................. 88 4.4.2 Quadrupole Ion-Trap Analyzers ..................................................... 89 4.4.3 Ion Cyclotron Resonance (ICR) ...................................................... 94 4.5 Proteolytic Degradation of Proteins ......................................................... 99 4.6 Mass Mapping ................................................................................. 100 4.7 Peptide Sequencing with Mass Spectrometry ............................................. 101 4.8 Tandem Mass Spectrometry and Post-Translational Modifications .................... 105 4.9 Conclusion ..................................................................................... 107 4.10 Chapter 4 References ........................................................................ 108 Chapter 5 Tandem Mass Spectrometry of Deoxy-Pyridoxal-l—Phosphate Modified Peptides ................................................................ 110 vi 5.1 Introduction: Neutral Loss Products in Mass Spectrometry ............................. 110 5.2 Background and Nomenclature for Peptides Modified by Pyridoxal-l-Phosphate. .. 112 5.3 Procedure: Synthesis of dPLP Modified Peptides ....................................... 114 5.4 MALDI Experiments ......................................................................... 116 5.5 Electrospray Experiments .................................................................... 118 5.6 Results .......................................................................................... 119 5.7 Tandem Mass Spectrometry with MALDI F TMS2 ....................................... 122 5.8 Tandem Mass Spectrometry with ESI-MS2 ................................................ 126 5.9 Results and Discussion ........................................................................ 137 5.10 Chapter 5 References ........................................................................ 143 Chapter 6 The Development of a Method for Locating Deoxy-Pyridoxal-l- Phosphate Modified Peptides Using Tandem Mass Spectrometry and Neutral Loss Analysis ...................................................... 145 6.1 Introduction .................................................................................... 145 6.2 TIP39 Peptide: The Model for Method Development .................................... 148 6.3 Procedure for the Synthesis of dPLP-modified TIP39 ................................... 148 6.4 Procedure for Tryptic Digestion of dPLP-Modified TIP39 ............................. 150 6.5 Mass Spectrometric Analysis of the dPLP—Modified TIP39 Tryptic Digest .......... 151 6.6 Results and Discussion ........................................................................ 152 6.7 Conclusions .................................................................................... 167 6.8 Chapter 6 References ......................................................................... 168 Chapter 7 Identification of Deoxy-Pyridoxal—I-Phosphate Modification Sites in Proteins with Tandem Mass Spectrometry and Neutral Loss Analysis ............................................................................. 169 7.1 Introduction: Initial Strategy for Identifying Deoxy-Pyridoxal-l-Phosphate Binding Sites in Proteins ........................................................................... 169 7.2 Procedure for Sodium Borohydride Reduction of Proteins .............................. 171 7.3 Procedure for the Reduction, Alkylation, and Tryptic Digestion of Proteins ......... 174 7.4 Procedures for HPLC and Mass Spectrometric Analysis of Protein Digests ......... 174 7.5 Results of Proteolysis, Fractionation, and Mass Spectrometric Analysis of Protein Digests ................................................................................................ 176 7.6 Discussion ...................................................................................... 179 7.7 The New Strategy ............................................................................. 181 7.7.1 Cyanogen Bromide Treatment ...................................................... 181 7.7.2 Procedure for Cyanogen Bromide Treatment ..................................... 183 7.7.3 Procedure for the HPLC Separation of Cyanogen Bromide Products and Mass Spectrometric Analysis ............................................................... 185 7.7.4 Results .................................................................................. 185 7.8 Proteolysis of the Isolated dPLP-Modified Products from Cyanogen Bromide Treatment ............................................................................................ 192 7.8.1 Trypsin and Glu-C Digestion Procedures .......................................... 195 7.8.2 Procedures for the HPLC and Mass Spectrometric Analysis of Tryptic and Glu-C Digestion Products of Alanine Racemase 36-130 and Aspartate Aminotransferase 213-287 .................................................................. 200 vii 7.9 Results: Mass Spectrometric Analysis of Tryptic and Glu-C Digestion Products of Alanine Racemase 36-130 and Aspartate Aminotransferase 213-287 ....................... 202 7.9.1 LC-MS/MS Analysis of Tryptic Digestion of Alanine Racemase 36-130. . 202 7.9.2 LC-MS/MS Analysis of Glu-C Digestion of Alanine Racemase 36-130. . 207 7.9.3 LC-MS/MS of the Tryptic Digest of Aspartate Aminotransferase 213-287... 216 7.9.4 LC-MS/MS of Glu-C Digestion of Aspartate Aminotransferase 213-287. . 219 7.10 Summary and Conclusions .................................................................. 223 7.11 Chapter 7 References ........................................................................ 229 viii Table 2.1 Table 2.2 Table 2.3 Table 2.4 Table 4.1 Table 5.1 Table 7.1 Table 7.2 LIST OF TABLES Previously reported SAM and SAH levels from plant tissue ............ Comparison of SAM, SAH determination using PBA affinity versus previously reported data from the literature ............................... Free amino acid levels in leaves of wild type and —MMT mutant Arabidopsis. Adapted from Ref. 1 ......................................... Radioactivity measurements of SMM, protein, and methionine extracts indicating the incorporation of [ 5S]methionine into each pool. Adapted from reference 1 ............................................. One and Three Letter Codes of the 20 Common Amino Acids and Corresponding Residue Masses. Modified from ref. 5 .................. One and Three Letter Codes of the 20 Common Amino Acids and Corresponding Residue Masses. Modified from ref. 5 .................. The theoretical products of cyanogen bromide treatment of Alanine Racemase ....................................................................... The theoretical products of cyanogen bromide treatment of Aspartate Aminotransferase ................................................. ix 40 48 50 52 104 115 184 184 Figure 1.1 Figure 1.2 Figure 1.3 Figure 1.4 Figure 1.5 Figure 1.6 Figure 2.1 Figure 2.2 Figure 2.3 Figure 2.4 LIST OF FIGURES Structures of some known secondary metabolites produced by certain plants ................................................................... The diagram illustrates one-carbon metabolic pathways in plants. Abbreviations: Met (methionine), Hcy (homocysteine), SAM (S- adenosylmethionine), SAH (S-adenosylhomocysteine), SMM (S- methylmethionine), and THF (tetrahydrofolate) .......................... The incorporation of one-carbon units into the tetrahydrofolate (THF) pool (reactions represented by dashed reaction arrows). Abbreviations: ATP (adenosine triphosphate), NAD+ (nicotinamide adenine diphosphate) ......................................................... (A) The structural components of tetrahydrofolate (THF). THF can bind one-carbon units of different oxidation states at positions N5 and/or N10. (B) The structures of the one-carbon units and the corresponding oxidation states are listed below the structure of THF. Reproduced from Ref. 28 ................................................... Comparison of the methyl group demand between secondary and primary metabolites (GlyBet=glycine betaine). Adapted and modified from ref.l .......................................................... Reactions involved in the S-methylmethionine (SMM) cycle ......... Structures of amino acids involved in the S-methylmethionine cycle ............................................................................. Structures of some common matrix molecules that have been used in MALDI experiments and an uncommon matrix, 2,4- dinitrophenylhydrazine, used for this study .............................. Representative MALDI MS spectra of AG 50(NH4+) ion exchange fractions from wild type and —MMT Arabidopsis thaliana. Modified from ref. 1 ...................................................................... Structures of nucleoside derivatives related to one-carbon metabolism ..................................................................... 10 15 17 27 29 33 36 Figure 2.5 Figure 2.6 Figure 2.7 Figure 3.1 Figure 3.2 Figure 3.3 Figure 3.4 Figure 3.5 Figure 3.6 Figure 3.7 Figure 3.8 Illustration of the structural characteristics of phenylboronic acid, PBA, used for purifying SAM and SAH from plant tissue. The pH dependent geometry around the central boron atom is included. At the bottom, an illustration of a typical reaction between PBA and cis, co-planar diols is displayed. In the tetrahedral geometry, the distance between two hydroxyl groups, 2.42A, corresponds to an optimal length bonding to cis, co-planar diols ............................ HPLC chromatogram of the separation of components of a PBA fraction extract from wild type (WT) Arabidopsis tissue ................ HPLC chromatogram of the separation of components of a PBA fraction extract from mutant (-MMT) Arabidopsis tissue ............... An illustration of the sequence alignment of Methionine Methyltransferase (MMT) with Aspartate Aminotransferase (AAT).. Structures of free pyridoxal-l-phosphate (PLP) and bound PLP. Conventional number designations representing positions on the pyridine ring are included .................................................... The mechanism of aldimine formation. The left side of the dotted line displays deprotonated reactants, intermediates, and products. The corresponding protonated species are on the right side. The dotted arrows on the 5 intermediate refers to the reverse reaction yielding the reactants. Adapted and modified from ref. 7 ............... The conversion of an internal aldimine linkage between PLP and an enzyme lysine residue to an external aldimine between PLP and an amino acid substrate, R-NHZ. Adapted and modified from ref. 6. . An illustration of the relative positions of the planar PLP-substrate complex and the free lysine-258 residue which deprotonates the substrate at CC and protonates C4'. Adapted and modified from ref. 6 ................................................................................. The pH dependence of the UV-visible spectroscopy of aspartate aminotansferase. Adapted and modified from reference 12 ............ This figure demonstrates the application of mass mapping to identify modifications in proteins ..................................................... Two HPLC chromatograms of tryptic digestions of the same protein: (A) unmodified and (B) PLP modified. Adapted and modified from ref. 16 ........................................................................... xi 41 45 46 58 6O 62 64 66 70 73 75 Figure 4.1 Figure 4.2 Figure 4.3 Figure 4.4 Figure 4.5 Figure 4.6 Figure 4.7 Figure 4.8 Figure 5.1 Figure 5.2 Figure 5.3 Figure 5.4 Figure 5.5 The general components of a mass spectrometer ......................... An illustation of the electospray components and the conversion ions in the liquid phase to charged droplets and eventually gas phase ions. Adapted from ref. 2 ........................................................... Diagram of a quadrupole ion-trap mass spectrometer. Modified from Ref. 21 .................................................................... Quadrupole ion trap stability diagram. Adapted and modified from ref. 22 ........................................................................... Illustration of a typical cubic ICR cell in a FTMS instrument. Modified from ref. 23 ......................................................... Illustration of the conversion of an image current to a mass spectrum with application of a fourier transform. Adapted and modified from ref. 24 ........................................................................... (A) Illustration of cleavage sites along the amide backbone of peptides and product nomenclature. (B) Examples of bn and yn structures. Adapted from ref. 5 ............................................. Tandem mass spectrum of precursor ion of m/z 421.5 (+2 charge of peptide VATVSLPR). De novo sequencing was used to identify several b and y ions that were used to sequence the peptide. Adapted from ref 5 ............................................................ The condensation reaction between the peptide F ibrinogen and pyridoxal-l-phosphate (PLP) followed by reduction with sodium borohydride (N aBH4) ......................................................... HPLC purification of unmodified and dPLP-modified Fibrinogen (FPLP) .......................................................................... A MALDI TOF spectrum of the HPLC purified mixture of Fibrinogen and dPLP-modified Fibrinogen (FPLP) ...................... MALDI FTMS spectrum of the mixture of peptides Fibrinogen and dPLP-modified Fibrinogen (FPLP) ......................................... A tandem mass spectrum of Fibrinogen obtained with MALDI FTMSZ. The sequence of F ibrinogen is displayed above the spectrum, with an illustration of the b and y ions that could result with fragmentation along the amide backbone of the peptide ........... xii 83 87 90 93 95 98 103 106 113 117 120 123 124 Figure 5.6 Figure 5.7 Figure 5.8 Figure 5.9 Figure 5.10 Figure 5.11 Figure 5.12 Figure 5.13 Figure 5.14 Figure 5.15 A tandem mass spectrum of dPLP-modified Fibrinogen (F PLP), (Mir), obtained from MALDI FTMS2 .................................... 125 A base peak ion chromatogram illustrating the separation of F ibrinogen and dPLP-modified Fibrinogen (F PLP) ...................... 127 Full scan LC-MS spectra at retention times 12.2 minutes (A) and 15.0 minutes from the separation of Fibrinogen and dPLP-modified Fibrinogen (F PLP). Refer to the chromatogram in Figure 5.7 ......... 128 CID spectra of F ibrinogen with different charge state precursors: (A) +1 charge, m/z 1189.6 Da, (B) +2 charge, m/z 595.6 Da, and (C) +3 charge, m/z 397.7 Da ......................................................... 130 CID spectra of PLP-modified Fibrinogen (FPLP) with different charge state precursors: (A) +1 charge, m/z 1420.6 Da, (B) +2 charge, m/z 711.2 Da, and (C) +3 charge, m/z 474.6 Da ............... 132 This figure describes singly and multiply charged precursors (M = phosphopeptides) undergoing CID and yielding neutral loss products corresponding to the loss of H3PO4. A11 neutral products (H3PO4) have been omitted for simplicity ............................................ 135 (A) A representation of the CID of dPLP peptides with different charge states all yielding neutral loss product ions corresponding to neutral losses of H3PO4 and dPLP. (B) The observed relative losses in m/z value from loss of H3PO4 and dPLP from dPLP peptides with charge state n .................................................................. 136 (A) Proposed six-centered transition state to explain the neutral loss of H3PO4 from a phosphopeptide precursor in tandem mass spectrometry. (B) Phosphotyrosine lacks an aliphatic hydrogen to complete a six-centered transition state. Neutral losses of H3PO4 are not observed. Adapted and modified from Ref. 5 ........................ 138 The proposed mechanism for the neutral loss of H3PO4 (231 Da) from dPLP-modified peptides in tandem mass spectrometry. The secondary amine marked a is the terminal side chain amine from the modified lysine residue of the peptide ...................................... 140 The proposed mechanism for the neutral loss of dPLP (98 Da) from dPLP-modified peptides in tandem mass spectrometry. The secondary amine marked a is the terminal side chain amine from the modified lysine residue of the peptide ...................................... 141 xiii Figure 6.1 Figure 6.2 Figure 6.3 Figure 6.4 Figure 6.5 Figure 6.6 Figure 6.7 Figure 6.8 Figure 6.9 Figure 6.10 Figure 6.11 This figure demonstrates the application of mass mapping to identify modifications in proteins ..................................................... This figure displays the amino acid sequence of TIP39 (A) along with the theoretical tryptic digestion products of TIP39 (B) ............ This figure demonstrates the application of mass mapping to identify modifications in proteins ..................................................... MALDI TOF spectrum of the tryptic digestion products of the mixture TIP39 and TIP39PLP. The peaks are annotated according to the sequence they represent. * represents products modified by dPLP ............................................................................ HPLC chromatogram displaying the separation of peptides from the tryptic digestion of TIP39 and dPLP-modified TIP39. The peaks are annotated according to the sequence they represent within TIP39. The peaks marked with * correspond to peptides modified by dPLP.. MALDI TOF spectrum corresponding to the chromatographic peak marked 24-39 from Figure 6.5 ............................................. MALDI TOF spectrum corresponding to the chromatographic peak marked 24-39 from Figure 6.5 ............................................. A comparison of the peptides that would be formed from the digestion of TIP39 (column A) versus dPLP-modified TIP39 (column B) ..................................................................... (A) A displays of the base peak ion chromatogram of the LC-MS2 experiment of the tryptic digestion products of the mixture of TIP39 and dPLP-TIP39. (B) and (C) display neutral loss chromatograms of 32.7 and 77 Da respectively ................................................. LC-MS full-scan spectra corresponding to retention times 32.6 and 34.8 minutes (A and B respectively). Spectrum A displays the +2 and +3 charge states for dPLP-modified peptide 24-39 and Spectrum B displays the same for dPLP-modified peptide 23-39 .................. MS/MS spectra of +3 precursors (A) 24-39 and (B) 23-39 of dPLP- peptides from the tryptic digestion of dPLP-modified TIP39 ........... xiv 147 149 153 155 156 157 159 160 161 163 165 Figure 6.12 Figure 7.1 Figure 7.2 Figure 7.3 Figure 7.4 Figure 7.5 Figure 7.6 Figure 7.7 Figure 7.8 Figure 7.9 Figure 7.10 Figure 7.11 Figure 7.12 (A) MS/MS spectrum of +2 precursor of dPLP peptide 24-39 (m/z 1085.5). (B) MS/MS spectrum of +2 precursor of dPLP peptide 23— 39 (m/z 1163.0) ................................................................ The initial strategy employed to locate dPLP modifications in proteins ......................................................................... The amino acid sequence in one letter designations for Alanine Racemase. The underlined lysine residue is the known PLP binding site (K-39) ...................................................................... The amino acid sequence in one letter designations for Aspartate Aminotransferase. The underlined lysine residue is the known PLP binding site (K-258) ........................................................... UV-visible spectra of Alanine Racemase before and after NaBH4 reduction ........................................................................ UV-visible spectra of Aspartate Aminotransferasese before and after NaBH4 reduction ............................................................... An illustrates of the reaction between a methionine residue in a protein and cyanogen bromide generating a product mixture of homoserine and homoserine lactone ........................................ HPLC chromatogram for the separation of CNBr products of Alanine Racemase ............................................................ HPLC chromatogram for the separation of CNBr products of Aspartate Aminotransferase ................................................. MALDI TOF spectrum of the fraction corresponding to the chromatographic peak at 27.5 minutes in the HPLC chromatogram in Figure 7.7 .................................................................... MALDI TOF spectrum of the fraction corresponding to the chromatographic peak at 27.8 minutes in the HPLC chromatogram in Figure 7.8 .................................................................... UV-visible spectrum of the HPLC fraction corresponding to the chromatographic peak at 27.5 minutes from Figure 7.7 .................. UV-visible spectrum of the HPLC fraction corresponding to the chromatographic peak at 27.8 minutes from Figure 7.8 .................. XV 166 170 172 173 177 178 182 186 188 189 191 193 194 Figure 7.13 Figure 7.14 Figure 7.15 Figure 7.16 Figure 7.17 Figure 7.18 Figure 7.19 Figure 7.20 The sequence (36-130) of the isolated CNBr product of Alanine Racemase. The predicted products of trypsin digestion are also listed ............................................................................. The sequence (36-130) of the isolated CNBr product of Alanine Racemase. The predicted products of Glu-C digestion are also listed ............................................................................. The sequence (213-287) of the isolated CNBr product of Aspartate Aminotransferase. The predicted products of tryptic digestion are also listed ....................................................................... The sequence (213-287) of the isolated CNBr product of Aspartate Aminotransferase. The predicted products of Glu-C digestion are also listed ....................................................................... (A) The base peak ion chromatogram illustrating the separation/detection of the tryptic products of AlRace 36-130. (B) The neutral loss chromatogram indicating the retention times in which the neutral loss of 32.7 Da was detected in a MS/MS spectrum. (C) The neutral loss chromatogram indicating the retention times in which the neutral loss of 77 Da was detected in an MS/MS spectrum .............................................................. A full-scan ESI spectrum at about 23.4 minutes within the LC- MS/MS run represented by the base peak ion chromatogram in Figure 7.17A. The three annotated peaks in the spectrum represent three charge states for the tryptic digestion product 36-52 of AlRace 36-130 ........................................................................... (A) MS/MS spectrum of precursor ion 994.5 Da. (B) MS/MS spectrum of precursor ion 663.0 Da. (C) MS/MS spectrum of precursor ion 497.5 Da ....................................................... (A) Base peak ion chromatogram illustrating the separation/detection of the Glu—C products of AlRace 36-130. (B) The neutral loss chromatogram indicating the retention times in which the neutral loss of 32.7 Da was detected in an MS/MS spectrum. (C) The neutral loss chromatogram indicating the retention times in which the neutral loss of 77 Da was detected in an MS/MS spectrum .............................................................. xvi 196 197 198 199 203 205 206 208 Figure 7.21 Figure 7.22 Figure 7.23 Figure 7.24 Figure 7.25 Figure 7.26 Figure 7.27 Figure 7.28 Figure 7.29 Figure 7.30 Figure 7.31 (A) Base peak ion chromatogram illustrating the separation/detection of the Glu-C products of AlRace 36-130. (B) The neutral loss chromatogram indicating the retention times in which the neutral loss of 49 Da was detected in an MS/MS spectrum. (C) The neutral loss chromatogram indicating the retention times in which the neutral loss of 115.5 Da was detected in an MS/MS spectrum .......................................................... (A) Full scan ESI spectrum at 18.9 minutes of the base peak ion chromatogram in Figures 7.20A and 21A (they are the same chromatogram). (B) Full scan ESI spectrum at 24.5 minutes of the base peak ion chromatogram in Figures 7.20A and 21A ................ (A) MS/MS of precursor ion m/z 717.0. (B) MS/MS of precursor ion m/z 478.6 .................................................................. (A) MS/MS spectrum of precursor ion m/z 1200.9. (B) MS/MS spectrum of precursor ion m/z 801.0 ....................................... An illustration of the mixture of digestion products possible by cleavage at D-47 or by the missed cleavage at D-47 ..................... (A) The base peak ion chromatogram of the LC-MS/MS analysis of the tryptic digestion of AAT. (B) The neutral loss chromatogram for loss of 49 Da a precursor ion. (C) The neutral loss chromatogram for loss of 115.5 Da a precursor ion ........................................ This figure displays a full scan spectrum at 35.1 minutes within the LC-MS/MS analysis of the tryptic digestion of AAT 213-287 ......... (A) MS/MS spectrum of precursor ion m/z 1631.7 Da. (B) MS/MS spectrum of precursor ion m/z 1642.4 Da. (C) MS/MS spectrum of precursor ion m/z 1650.9 Da ................................................ (A) Full scan spectrum (retention time 35.3 minutes) from the LC- MS/MS analysis of the Glu-C digestion of AAT 213-287. All three annotated peaks are unrelated. (B) MS/ MS spectrum of precursor ion m/z 1632.2 Da ............................................................ An illustration of the mixture of digestion products possible by cleavage at E-265 or by the missed cleavage at E-276 ................... Summary of the final approach to locating the site of dPLP modification within a protein. Arrows crossed with x symbolize steps from the initial strategy (Figure 7.1) that were ineffective and eliminated ..................................................................... xvii 209 211 212 214 215 217 218 220 222 224 227 CHAPTER 1 INTRODUCTION AND BACKGROUND OF PLANT ONE-CARBON METABOLISM The two projects described in this report stem from a collaborative effort to characterize one-carbon metabolism in plants. The first project that will be discussed is the development of techniques used to quantify key intermediates involved in one-carbon metabolic pathways. The second project, to be discussed in the later chapters, involves the development of a protocol for determining the site of pyridoxal-l-phosphate modification in proteins. The second project is related to a structural aspect of one of the enzymes involved in plant one-carbon metabolism. A protocol was developed here to address this aspect and can be applied to some enzymes that are involved in these pathways. There are many unknowns about one-carbon metabolism, especially in plants. This chapter is dedicated to summarize what is known, what is not known, and detail the impetus behind the efforts to learn more about plant one-carbon metabolism. 1.1 Introduction to Plant One-Carbon Metabolism Plants, like other eukaryotic organisms, require one-carbon units for the essential biosynthesis of primary metabolites such as nucleic acids, proteins, pantothenate, and some amino acids (1). In addition to these primary metabolic requirements for one- carbon units, plants demand methyl groups for the many synthetic products of secondary metabolism (2). Secondary metabolites are not directly involved in growth and development. They are specialized molecules that are unique to plants and are designed for protection against specific environmental stresses. These secondary metabolites are essential in allowing plants to adapt to extreme environmental conditions such as temperature, drought, salinity, and even predation. However, it is the potential value of secondary metabolites to agriculture and the pharmaceutical industry that has raised interest and accelerated research into one-carbon metabolism in the last ten years. The source(s) of one-carbon units is still under debate. However, it is known that they are stored in two main pools until needed. One pool consists of the tetrahydrofolate mediated pathways which store one—carbon units as formyl, methenyl, methylene and methyl groups (3). These units are donated from the folate pool for anabolic reactions forming primary metabolites. The other pool of one-carbon units is the activated methyl cycle (2). Methyl groups are donated from this pool for the synthesis of a number of primary metabolites. However, this pool is also responsible for supplying methyl groups to the many number of metabolites in secondary metabolism as well. Despite the obvious importance of these pathways to the plant and the potential of these secondary metabolites in industry, relatively little is known about one-carbon metabolism in plants (1). Below is a brief description of the present understanding of plant one-carbon metabolism. Included is information on knowledge and potential utility of secondary metabolites, present status of the understanding of the metabolic pathways and regulation, and a summary of the goals sought community interested in one-carbon metabolism in plants. 1.2 Secondary Metabolites All species that survive in a habitat have found ways to adapt and thrive in their environment. Some tools needed are supplied in an organism’s genetic code. Mobile organisms like mammals can learn to forage or hunt for food and to escape from predators. Without the ability to move, plants are quite vulnerable to their environment. They cannot run from predators and they must rely only on rainfall and nutrients in the soil. Plants have developed some interesting ways of dealing with these disadvantages. They produce chemicals that allow them to retain water, to tolerate extreme climactic conditions, produce toxic chemicals that ward off herbivores, and build physical barriers for protection against the environment. These chemicals fall under the category of secondary metabolites (6). They are not generated, nor required, for the standard biochemical pathways involved in grth and development. They produced only to respond to specific threats present in the environment. Alkaloids, lignin, and osmo- protectants are just a few examples of secondary metabolites that help plants deal with environmental stresses. Alkaloids are a class of compounds involved in pest resistance (5). They are invaluable in protecting plants from predation , but can also act against other plants, reducing competition for space, nutrients, and sunlight. Nicotine and caffeine are commonly known alkaloids that are potent chemical deterrents (5). In addition to their importance to plants, alkaloids are well known commodities for their medicinal value (6). Some have been observed to have physiological and antibiotic properties. Morphine and codeine are effective pain relievers and camptothecin has been used as an anti-cancer treatment (5). Lignin is a structural component of cell walls (7). The lignin content controls the rigidity of plant stems and tree trunks. It serves as a physical barrier to weather, insect infestation, and other environmental elements. In industry, lignin is an undesired component in pulp and its removal is a major step in the processing of pulp into paper (8). However, lignin is energy-rich and desirable for biofuel plants (9). It is therefore desirable to control lignification in plant breedings. Osmo-protectants induce high tolerance to drought, high salinity, and other stresses in plants (4). There has been a real interest in the agricultural industry to have plants produce larger quantities of osmo-protectants, such as glycine betaine. This would increase the growth potential and result in higher crop yields. Furthermore, transgenic plants that are engineered to accumulate osmoprotectants could potentially tolerate extreme environmental conditions. Some research groups have already introduced salt tolerance in plants that normally lacks such capability (10). Ultimately, if successful this could enable food to be grown in regions that are normally too dry and salty. The properties that secondary metabolites display have made them major targets for metabolic engineering. They have simple biosynthetic pathways and can be accumulated to high concentrations without toxic side effects. Though structurally diverse, these compounds do have a common structural component, which is the requirement for methyl groups (2). Figure 1.1 displays the structures of some secondary metabolites. Methyl groups are donated for the synthesis of these compounds from one- carbon metabolic pathways. However, before any major metabolic engineering efforts can be made, more has to be learned about the nature of one-carbon metabolism in plants. Engineering plants to produce increased levels of osmoprotectants would have a significant impact on methyl group demand. Alkaloids fl CH3 N “Tit /> o/\T N CH3 Caffeine Lfignhr Polymer '\\\ / OR CHRflOjPflflHRflUnmS H3C + H3C H30 coor—r Glycine Betaine Polymer Polymer \ \ / OCH3 H3C0/// /// OCH3 OR OR Figure 1.1. Structures of some known secondary metabolites produced by certain plants. 1.3 Established F acts/Characteristics of One-Carbon Metabolism in Plants Figure 1.2 summarizes the one-carbon metabolic pathways in plants as they are presently known. Although very similar to other eukaryotic organisms, there are some subtle differences that are unique to plants. Evidence is still being gathered about the sources of one-carbon units, folate metabolism, the activated methyl cycle, and the regulation of one-carbon metabolism as a whole. More information about these topics will be covered in the sections that follow. Secondag Metabolites Lignin Alkaloids Osmo-protectants A Methyl groups Lipids, DNA q...SAM ‘SAH Methyl ation FM/eLEMM Ho} ‘u Proteins THPF B-Methyl-THF Serine & ' Glycine . A'“ £3111)“ 1dy late ‘""' 5 , l 0-yethylene-THF Nucleic Acids 5,10—Metheny1—THF \ " 5-Formyl-THF Purines < ------ IO-Formyl-THF Figure 1.2. The diagram illustrates one-carbon metabolic pathways in plants. Abbreviations: Met (methionine), Hcy (homocysteine), SAM (S-adenosylmethionine), SAH (S-adenosylhomocysteine), SMM (S-methylmethionine), and THF (tetrahydrofolate). 1.3.1 Sources of One-Carbon Units Strong evidence now suggests that the amino acids serine and glycine are sources of one-carbon units. They can both enter the folate pool as 5,10-methylene-THF in separate enzymatic reactions (11). Serine is converted to glycine in a reversible reaction catalyzed by serine hydroxymethyltransferase (SHMT), which utilizes THF as a cofactor. THF accepts the one-carbon unit forming 5,10-methylene-THF: Serine + THF 2 Glycine + 5,10-methylene-THF + H20 (1.1) The formation of glycine is favored in Equation 1.1. Decarboxylation of glycine contributes one-carbon units to the folate pool in an oxidative reaction catalyzed by the glycine decarboxylase (GDC) complex, equation 1.2 (12). Glycine + NAD + THF —) C0; + NADH + NH3 + 5,10-methylene-THF (1.2) where NADH is nicotinamide adenine dinucleotide hydride. Studies have also shown that formate is a source of one-carbon units to the folate pool. Radiotracer experiments show that formate is readily incorporated into the folate pool as IO-formyl-THF (13). This is an adenosine triphosphate (ATP) dependent reaction catalyzed by 10-forrny1- THF-synthetase: Formate + ATP —> ADP + 10-formyl-THF (1.3) The product ADP is adenosine diphosphate. Figure 1.3 illustrates the incorporation of one-carbon units from serine, glycine, and formate into the folate pool. It has been suggested that methanol and formaldehyde could represent additional one-carbon sources, but this is not yet substantiated (1). Activated Methyl Cycle ---------- One-carbon sources T Folate one-carbon conversions ( 5-Methyl-THF Serine + THF .o‘ 5,10-Methylene-THF 1 3... Glycine + NAD“ + THF Folate Pool I 5 ,10-Methenyl-THF 5-Formyl-THF K lO-Formyl- l. Serine Hydroxymethyltransferase 2. Glycine Decarboxylase Complex ADP ‘3‘ 3 3. lO-formyl- THF-synthetase 'Q Formate + ATP + THF Figure 1.3. The incorporation of one-carbon units into the tetrahydrofolate (THF) pool (reactions represented by dashed reaction arrows). Abbreviations: ATP (adenosine triphosphate), NAD+ (nicotinamide adenine diphosphate). 1.3.2 The One-Carbon Groups and the Inter-conversion Reactions of the Folate Pool As already mentioned, one-carbon units exist, and are needed, in different chemical forms. Figure 1.4B illustrates the structure and oxidation state of each form of one-carbon unit. Tetrahydrofolate (THF) is the vehicle that inter-converts one-carbon units to various oxidation states (14). It functions to bind, transport, and donate one- carbon units for anabolic reactions. Since one-carbon units bind to THF in various oxidation states, THF exists in diverse chemical forms that are collectively called the folate, or vitamin B9, pool. THF is composed of three distinct structural components, Figure 1.4A (15, 28). Pteridine, p-aminobenzoate, and L-glutamate are incorporated into a THF molecule through a complex pathway, which predominately occurs in plant mitochondria. One-carbon units bind at the N5 and/or the N10 positions. Pteridine is formed from guanosine triphosphate (GTP) in a pathway that is slightly different from the corresponding bacterial and mammalian pathway (3, 14). P- aminobenzoate is added onto the pteridine moiety by dihydropteroate synthase, forming dihydopteroate. This reaction requires ATP, and proceeds through a pyrophosphate intermediate (3). Completion of the folate structure occurs upon addition of glutamic acid. Dihydropterate synthase adds the first glutamate residue to the carboxylic acid group of p-aminobenzoic acid yielding dihydrofolate (14). The active one-carbon unit carriers are the fully reduced folates. Dihydrofolate reductase reduces dihydrofolate to tetrahydrofolate (THF) in a nicotinamide adenine dinucleotide phosphate hydride (NADPH) dependent reaction (3). Additional glutamate residues are added on by folylpolyglutamate synthetase (14). One to eight glutamate residues can be linked, and variability is typical in plants (16). It is not clear what purpose the glutamyl chain serves, o R X 0 COOH II ,5 ,1, / \ M HN | 5 Y‘lo _ NH COOH /’\ / P-amlnobenzoate Glutamate H2N N N H Pteridine B Folate R X Redox State Tetrahydrofolate, THF H H l 0-Formyl-THF H CHO -2 S-Formyl-THF CHO H -2 5-Methyl-THF CH3 H +2 5,1 O-Methenyl-THF =CH- -2 5,1 O-Methylene-THF -CH2- 0 Figure 1.4. (A) The structural components of tetrahydrofolate (THF). THF can bind one-carbon units of different oxidation states at positions N5 and/or N10. (B) The structures of the one-carbon units and the corresponding oxidation states are listed below the structure of THF. Reproduced from Ref. 28. 10 nor the significance of the variable numbers of glutamate residues. It could be that these glutamate chains regulate folate intracellular transport across organelle membranes. It may also be possible that they facilitate binding to enzymes. It is not clear if the identity of the one-carbon unit attached has a bearing on the level of glutamylation. Further study is needed to ascertain the function of the glutamyl chain. Contrary to the rest of the pathway, which occurs in the mitochondria, glutamylation occurs in the cytosol and chloroplast (21). One-carbon units are stored in the folate pool for anabolic reactions. They can be inter-converted between oxidation states, depending on which form of one-carbon unit is needed (Figure 1.2). These one-carbon conversion reactions are mediated by THF. As described above and in Figure 1.3, one carbon units can enter the folate pool from formate forming 10-formyl-THF. The carbon atom in formyl groups carries a -2 oxidation state. 10-formyl-THF supplies primary metabolism with formyl groups for purine synthesis (18) and the formation of formylmethionyl-tRNA (19). The one-carbon group an also remain in the folate pool by conversion of lO-formyl-THF to 5,10- methenyl-THF. This is mediated by 5,10-methenyl-THF-cyclohydrolase (16). It is a reversible reaction. Methenyl groups also carry a -2 oxidation state. 5,10-methenyl- THF can be converted back to 10—formyl-THF by the same enzyme. However, it can also be converted to another formyl-THF derivative, 5-formyl-THF. Little is known about the necessity of this reaction. It may be that 5-formyl—THF serves as a storage molecule for formyl groups in the cell. Methenyl groups are converted to methylene groups in another reversible reaction that forms 5,10-methylene-THF (l6). Methylene groups carry a zero oxidation state on 11 the carbon atom. This conversion of 5,10-methenyl-THF to 5,10-methylene-THF is mediated by 5, lO-methylene—THF-dehydrogenase and requires NADPH. In plants, evidence indicates that the 5,10-methylene-THF-dehydrogenase and 5,10-methenyl-THF- cyclohydrolase activity are mediated by a bifunctional enzyme (14). Methylene groups can be donated from the folate pool for thymidylate, pantothenate, and nucleic acid synthesis (1). As discussed above, they are also used in the formation of serine and glycine which can be incorporated into proteins. 5,10-methylene-THF can then be converted to S-methyl-THF in a reversible reaction catalyzed by methylene-THF-reductase (MTHF R) (16). The conversion results in methyl groups, which carry a +2 oxidation state, from methylene. Methylene-THF- reductase represents the least understood enzyme involved in folate inter-conversion reactions (20). Despite being structurally similar to other eukaryotic MTHFR, there are some striking differences in plant MTHF R. Contrary to homologous MTHF R in mammals, Arabidopsis thaliana MTHFR uses NADH as the reductant rather than NADPH. The plant homolog is also resistant to feedback inhibition by S- adenosylmethionine. Formation of 5-methyl-THF in mammals is irreversible and feedback inhibition by S-adenosylmethionine prevents depletion of 5,10-methylene-THF. This would suggest that the MTHFR reaction in plants is reversible, as mentioned above. 1.3.3 The Activated Methyl Cycle 5-methyl-THF represents the link between the folate and the activated methyl cycle one-carbon unit pools. Like other eukaryotic organisms, plants donate methyl groups from the activated methyl cycle. As mentioned, methyl groups are required in 12 many biosynthetic reactions yielding primary metabolites. Additionally, plants need methyl groups to complete the synthesis of a diversity of specialized molecules known as secondary metabolites. The cycle begins with the transfer of a methyl group from the folate pool. The methyl group is donated from 5-methyl-THF to homocysteine, an amino acid not found in proteins, forming methionine (17). This methyl transfer is catalyzed by methionine synthase. Other than its involvement in one-carbon unit inter-conversion, methionine biosynthesis is the main role of 5-methyl-THF. In addition to methionine, the reaction results in the formation of free THF. The uniqueness of this reaction is that it is a trans- methylation reaction that is independent of S-adenosylmethionine, the principal methyl group donor in most organisms. In contrast, the corresponding reaction in animals and some bacteria requires S-adenosylmethionine and vitamin 812(3). Once formed, methionine participates in several pathways. It can be removed from the methyl cycle for the synthesis of polyamines, proteins, ethylene, or it can be further processed for methyl donation (17). The intermediate by which organisms donate methyl groups is S-adenosylmethionine (SAM) (22). Formation of this important molecule comes from the reaction of methionine with ATP. S-adenosylmethionine synthetase is the enzyme that catalyzes SAM formation (17). Phosphate and pyrophosphate are byproducts. Unlike the reversible, inter-conversion reactions of the folate pool, the methyl cycle is unidirectional. SAM is then available for methyl transfer reactions. A specific methyltransferase enzyme is responsible for mediating the methyl donation. The methyltransferase used depends on the methyl acceptor available and the primary or secondary metabolite to be 13 synthesized. S-adenosylhomocysteine (SAH) is the de-methylated byproduct. SAH is broken down by S-adenosylhomocysteine hydrolase resulting in the formation of adenosine as well as the regeneration of homocysteine, the initial methyl group acceptor from the folate pool (17). It is now available for re-methylation and another trip through the methyl cycle. It is not yet clear what happens to adenosine (17). 1.4 Unknown Aspects of Plant One-Carbon Metabolism As discussed above, although the importance of one-carbon metabolism in plants is obvious, little is known about it. The sources of one-carbon units are still not decisively established. Knowledge of the regulatory mechanisms of these pathways is scarce at both the gene and enzyme level. More is known about these pathways and regulation in bacteria, fungi, and mammals, so some generalizations can be made. However, before plants can be engineered for controlled production of targeted secondary metabolites, it is imperative that more knowledge is gained. Determination of one-carbon unit flux through these pathways would make it possible to understand some of these enigmas. In particular, how do these fluxes change when gene expression or enzyme activity is altered, or inhibited? How do these fluxes change when the plant is placed under some form of stress? The necessity of plants to produce secondary metabolites adds more confusion to understanding plant one-carbon metabolism. Studies show that demand for methyl groups in secondary metabolism can dwarf the demand for primary metabolism, Figure 1.5. Other eukaryotes are not burdened by such a demand. The question is then, how do 14 Beet Leaf Tobacco Root Birch Wood 1; 2000 dry 3° 1500 1000 500 pmol one-carbon units Figure 1.5. Comparison of the methyl group demand between secondary and primary metabolites (GlyBet=glycine betaine). Adapted and modified from ref. 1. plants meet this demand while still maintaining sufficient methyl supply for primary metabolism? 1.5 Introduction to the S-Methylmethionine (SMM) Cycle A major roadblock to answering these questions lies in the presence of the S- methylmethionine (SMM) cycle in plants (23). As mentioned above and illustrated in Figure 1.6, l-methionine can participate in protein synthesis or the methyl cycle. In a two-step cycle unique to plants however, l-methionine can acquire a methyl group forming SMM. Subsequent demethylation results in the regeneration of l-methionine. The methylation of l-methionine is carried out by the SAM dependant enzyme methionine S-methyltransferase (MMT) (24). Demethylation of SMM is mediated by homocysteine methyltransferase (HMT) (25). The methyl group from SMM is transferred to a homocysteine molecule generating two molecules of methionine. The necessity of this two-step cycle has been unclear (1). It would seem to be only capable of depleting the SAM pool and inhibiting the ability of the organism to carry out trans- methylation reactions. Furthermore, since the formation of SAM occurs with the consumption of ATP, it is a costly cycle. Without a clear function, the SMM cycle appears futile. Recent studies have found high levels of SMM in the phloem implicating SMM as a transport molecule of sulfur. More research will be needed in order to ascertain the full function and necessity of the SMM cycle. 16 O S H3C/ \/\(U\OH MMT V SAH NH2 Methionine H3C HMT s+ H3C/ \ OH NH2 0 S-methylmethionine (SMM) S H / \ OH NH2 MNIT = Methionine-S-Methyltransferase Homocysteine HIVIT = Homocysteine Methyltransferase SAM = S-adenosylmethlonine SAH = S-adenosylhomocystelnc Figure 1.6. Reactions involved in the S—methylmethionine (SMM) cycle. 17 1.6 Recent Findings in Regards to the S-Methylmethionine (SMM) Cycle Research on the SMM cycle in plants has already begun. These efforts have produced several clues that could add up to one or several possible functions of the SMM cycle. One approach that has been applied by the Hanson group was to study the machinery of the SMM cycle. They purified S-methylmethionine methyltransferase (MMT) from the leaf tissue of Wallastonia biflora (24). MMT is the methyltransferase that methylates methionine, in effect starting the SMM cycle (23). They were able to characterize MMT structurally and kinetically. Electrophoresis indicated a molecular mass estimate of 115-kDa. There was also a band at about 450-kDa which suggests a tetrameric tertiary structure. Activity studies indicated W. biflora MMT to have a pH optimum at 7.2. Kinetic analysis of MMT methyltransferase activity via substrate interaction and product inhibition indicated that SAM binds MMT before methionine, and SMM is released before the demethylated by-product, S-adenosylhomocysteine (SAH). The Hanson group extended the study of MMT to include sequence analysis and radio-tracer studies to follow the destination of SMM (26). cDNA clones of MMT from W. biflora, maize, and A. thaliana were isolated and the full length gene sequences were determined for each of these organisms. Analysis of the amino acid sequences revealed a methyltransferase domain near the N terminus, as expected. However, there were two peculiarities. One, the MMT amino acid sequence is about three times longer than other known plant methyltransferase. Two, sequence alignments indicated surprising sequence similarity between the C terminal region of MMT and other enzymes that bind pyridoxal- l-phosphate (PLP), especially aminotransferases. Modification by PLP has not yet been 18 proven nor has there been any MMT function or activity determined that would require PLP. However, these two observations suggest that MMT may serve other roles in addition to its methyltransferase activities. Radiotracer experiments were performed with 35S-methionine (26). SMM extraction found high levels of radiation in phloem exudates. This implies a role of SMM in sulfur transport. It is also possible that SMM could be demethylated by homocysteine methyltransferase forming methionine that can be used for the synthesis of grain proteins. While the reason for this transport to the phloem may not be completely clear, it certainly appears that SMM may be a transport molecule for sulfur assimilation and/or serve as a storage molecule for methionine. In an investigation unrelated to one-carbon metabolism, a previously unknown function of MMT has been discovered (27). Terry et a]. have suggested a function of plant MMT in selenium (Se) phytovolatilization. Phytovolatilization is the uptake and transpiration of contaminants, mostly volatile organic compounds, in plants. Analogous to the mammalian liver, plants have phytoremediation capabilities that allow them to remove certain toxic pollutants. Selenium can be tolerated in only small doses. It is in fact essential to most animal nutrition, but in small quantities. The authors hypothesized that MMT was involved in Se volatilization. To test this, they measured MMT gene expression in the presence and absence of selenate using northern analysis. Up- regulation of MMT was observed in roots and leaves, but not in stem tissue. This insinuated a role by MMT. To further test the hypothesis, the authors engineered a T- DNA knockout of the A. thaliana MMT gene. This produced a mutant that lacked MMT activity. They then measured the level of Se volatilization in the knockout mutant and 19 compared that to wild type Arabidopsis. Production of volatile Se was about ten times greater than that produced by mutant plants. Corroborating this was the finding that mutant tissue had greater amounts of Se than the wild type. This indicated that the inactivity of the MMT gene product rendered the mutant incapable of removing Se. Phenotypically, both the wild type and mutant could tolerate low doses of Se. However, as Se doses increased, the mutant suffered some growth deficiencies. At 10 to 20 pM Se- met, wild type roots were three times longer than mutant roots. 1.7 Continuing the SMM Investigation One focus of this study was to increase our understanding of the SMM cycle. The strategies employed required development work in some analytical and bio-analytical chemistry techniques. Approaches were developed for isolating and quantifying some key metabolic intermediates in the activated methyl cycle. The application of these approaches yielded valuable information that is helping to unlock certain mysteries of the SMM cycle. This report also summarizes progress in the location of PLP modified residues in proteins. The methods employed here have potential for future studies of MMT. For example, a positive determination of PLP modification of MMT could lead to some real insight to possible, unknown activities of this enzyme. 20 1.8 Chapter 1 References 1. 10. 11. 12. 13. Hanson, A.D., Roje, S. One-carbon metabolism in higher plants. Annu. Rev. Plant Physio]. Mol. Biol. 52: 119-137 (2001). Poulton, J .E. The Biochemistry of Plants, vol. 7. Academic Press, New York, p. 667-723 (1981). Cossins, EA. The Biochemistg of Plants, v01. 2. Academic Press, New York, p. 365-418 (1980). Bohnert, H.J., Nelson, D.E., Jensen, R.G. Adaptations to environmental stress. Plant Cell. 7: 1099-1111 (1995). Croteau, R., Kutchan, T.M., Lewis, N.G. Biochemistry and Molecular Biology of Plants. Edited by BB. Buchanan, W. Gruissem, R.L. Jones. American Society of Plant Biologists, Rockville, MD., p. 1250-1318 (2000). Kutchan, T.M. Alkaloid biosynthesis- the basis for metabolic engineering of medicinal plants. Plant Cell. 7: 1059-1070 (1995). Whetten, R., Sederoff, R. Lignin biosynthesis. Plant Cell. 7: 1001-1013 (1995). Sederoff, R. Building better trees with antisense. Nature Biotech. 17: 750-751 (1999) Sederoff, R., Campbell, M., O’malley, D., Whetten, R. Genetic regulation of lignin biosynthesis and the potential modification of wood by genetic engineering in loblolly pine. In Genetic Engineering of Plant Secondag Metabolism. Edited by BB Ellis, GW Kuroki, HA Stafford, Plenum Press, New York, p. 313-355 (1994) Jensen, R.G., Sheveleva, E., Chmara, W., Bohnert, HJ. Increase Salt tolerance by D-ononitol production in transgenic Nicotiana tabacum L. 115: 1211-1219 (1997) Prabhu, V., Chatson, K.B., Abrams, G.D., King, J. 13C Nuclear Magnetic Resonance Detection of Interaction of Serine Hydroxymethyltransferase with C1- Tetrahydrofolate Synthase and Glycine Decarboxylase Complex Activities in Arabidopsis. Plant Physiol. 112: 207-216 (1996). Douce, R., Bourguignon, J ., Neuburger, M., Rebeille., F. The glycine decarboxylase system: a fascinating complex. Trends in Plant Science. 6(4): 167-176 (2001). Li, R., Moore, M., Kinj, J. Investigating the role of one-carbon metabolism in Arabidopsis thaliana. 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John Wiley & Sons, New York. p. 457-495 (1984). Roje. S., Wang, H., McNeil, S.D., Raymond, R.K., Appling, D.R., Schachar_Hill, Y., Bohnert, H.J., Hanson, A.D. Isolation, characterization, and functional expression of cDNAs encoding NADH-dependent methylenetetrahydrofolate reductase from higher plants. J. Biol. Chem. 274(51): 36089-36096 (1999). Ravanel, S., Cherest, H., Jabrin, S., Grunwald, D., Surdin-Kerjan, Y., Douce, R., Rebeille, F. Tetrahydrofolate biosynthesis in plants: molecular and functional characterization of dihydrofolate synthetase and three isoforms of folylpolyglutamate synthetase in Arabidopsis thaliana. Proc. Nat]. Acad. Sci. USA. 98: 15360-15365 (2001). Chiang, P.K., Gordon, R.K., Tal, J ., Zeng, G.C., Doctor, B.P., Pardhasaradhi, K., McCann, P.P. S-adenosylmethionine and methylation. FASEB J. 10: 471-480. Mudd, S.H., Datko, A.H. The S-methylmethionine cycle in Lemna paucicostata. Plant Physiol. 93, 623-630 (1996). James, F ., Nolte, K.D., Hanson, A.D. Purification and properties of S- adenosylmethionine: L-methionine S-methyltransferase from Wallastonia biflora leaves. .1. Biol. Chem. 270(38): 22344-22350 (1995). Ranocha. P., Bourgis, F ., Ziemak, M.J., Rhodes, D., Gage, D.A., Hanson, A.D. Characterization and functional expression of cDNAs encoding methionine- 22 26. 27. 28. sensitive and —insensitive homocysteine S-methyltransferases from Arabidopsis. J. Biol. Chem. 275(21): 15962-15968 (2000). Bourgis, F., Roje, S., Nuccio, M.L., Fisher, D.B., Tarczynski, M.C., Li, C., Herschbach, C., Rennenberg, H., Pimenta, M.J., Shen, T., Gage, D.A., Hanson, A.D. S-methylmethionine plays a role in phloem sulfur transport and is synthesized by a novel type of methyltransferase. Plant Cell. 11: 1485-1497 (1999) Terry, N., Berken, A., Tagmount, A. An essential role of S-adenosyl-L- methionine: L-methionine S-methyltransferase in selenium volatilization by plants. Methylation of selenomethionine to selenium-methyl-L-selenium- methionine, the precursor of volatile selenium. Plant Physiol. 130: 847-856 (2002) Hanson, A.D., Gage, D.A., Schachar-Hill, Y. Plant one-carbon metabolism and its engineering. Trends Plant Sci. 5(5): 206-213 (2000). 23 CHAPTER 2 THE DETERMINATION OF S-METHYLMETHIONINE (SMM), S— ADENOSYLMETHIONINE (SAM), AND S-ADENOSYLHOMOCYSTEINE (SAH) IN PLANTS 2.1 Introduction As described in chapter l,the S-methylmethionine (SMM) cycle is a unique pathway in plants. Although organisms in the plant kingdom are unlikely to expend precious energy, in the form of adenosine triphosphate (ATP), for the formation of a useless metabolite (SMM), the cycle does appear futile. In review, SMM is formed via the methylation of a methionine molecule. This is the first step of the SMM cycle and is mediated by methionine methyltransferase (MMT). Since this is a S-adenosylmethionine (SAM) dependent methyltransferase, a molecule of ATP is sacrificed. The short cycle is completed upon the demethylation of SMM. The methyl group is transferred from SMM to a molecule of homocysteine forming two molecules of methionine. This last step of the SMM cycle is catalyzed by homocysteine methyltransferase. The following is a report of a recent effort to ascertain the function(s) of the SMM cycle. The big question then is: why does the SMM cycle exist? In chapter 1, some clues and recent findings were discussed that suggested it may not be a futile cycle. In several plant species, SMM was detected in the phloem suggesting a role of SMM as a transporter of sulfur. Studies of MMT imply that this enzyme may not function solely as a methyltransferase. It appears to be involved in selenium volatilization and its primary structure suggests an unknown activity that may involve pyridoxal-l-phosphate (PLP). If 24 there’s more to this cycle, what might it be? There are some reasonable hypotheses (1). Perhaps the SMM cycle functions to store methionine, as SMM, to prevent depletion of the methionine pool. When methyltransferase reactions are turned on, there is a high demand for SAM. Since SAM is synthesized from methionine, it is possible that SAM synthesis regulates the methionine pool available for protein synthesis. Another possibility is that the SMM cycle functions to control the methylation ratio. Briefly, this is simply the ratio of SAM to S-adenosylhomocysteine (SAH) in the plant tissue. SAH is the byproduct of a methyl group transfer between SAM and an acceptor molecule. SAH is a competitive inhibitor of methyltransferase reactions (2). Therefore, high levels of SAH, or a low methylation ratio, inhibit methyl group donations by the activated methyl cycle. 2.2 Strategy for Establishing the Function of the S-Methylmethionine Cycle To test these hypotheses, a simple strategy was employed: eliminate the SMM cycle (1). Two populations of Arabidopsis thaliana were studied. They were both grown under equivalent conditions. One population was simply the wild type, WT, Arabidopsis. The other was a mutant form of Arabidopsis. Specifically, it was an isolated insertional mutant form of the MMT gene, -MMT, of Arabidopsis that expressed an inactive MMT gene product. In other words, this mutant lacked MMT activity. Theoretically, since the mutant was unable to generate active MMT and methylate methionine, the SMM cycle should effectively be eliminated. The wild type and mutant Arabidopsis plants were grown and prepared by Dr. Andrew Hanson and his research group at the University of Florida (1). 25 The next step was to determine the effect of the missing cycle. First, it had to be proven that the SMM cycle had been eliminated. This could be done by showing that the mutant plant lacked SMM. To do this, a protocol for SMM determination in plant tissue had to be established. 1f the SMM cycle was eliminated, then there should be a negative result for SMM in —MMT, and a positive result from WT. Once established, efforts could be focused on determining the difference, if any, in the methylation ratios (SAM / SAH) between WT and —MMT. The determination of the methylation ratio would in turn require the development of a protocol for SAM and SAH measurements in plant tissue. 2.3 The Determination of S-Methylmethionine From Plant Tissue SMM is an amino acid that is not incorporated into the primary structure of proteins. Figure 2.1 illustrates the structure of SMM as well as methionine and homocysteine. Like SMM, homocysteine is an amino acid that is not used for protein synthesis. Conventional analysis of amino acids from plant tissue employs chemical derivatization, commonly by tert- butyldimethylsilylation (TBDMS), followed by GC separation and detection (3). Such an approach has been used for amino acid determination from plant tissue (4). However, the sulfonium group of SMM prohibits GC analysis due to its involatility. With that in mind, an approach was considered here that involved liquid-liquid extraction followed by selective isolation of SMM via ion- exchange. The presence, or absence, of SMM would then be determined mass spectrometry. 26 0 /‘S‘ \ HS HJC OH OH NH-fi ‘ NH2 l-methionine I-homocysteine “of 0 ‘4- b Hac/ \/\ OH H H2 S-methylmethionine (SMM) Figure 2.1. Structures of amino acids involved in the S-methylmethionine cycle. 27 2.3.1 Measurements of SMM with Mass Spectrometry As mentioned above, GC analysis of SMM is inhibited by its lack of volatility. As an alternative, SMM was isolated directly from plant tissue and analyzed with matrix assisted laser desorption ionization time of flight mass spectrometry was used (MALDI TOF MS) (5, 6). MALDI is an ionization technique that forms protonated molecular ions (7). Samples are typically mixed with an excess matrix compound and allowed to co- crystallize with it upon drying. There are primarily a select few of these matrices that are used for the majority of MALDI experiments (7). Among the favorites are 2,5- dihydroxybenzoic acid (DHB), a-cyano-4-hydroxycinnamic acid, sinapinic acid and 6- aza-2—thiothymine (ATT), Figure 2.2. Irradiation of the sample with a nitrogen laser (337nm) results in desorption of matrix and analyte molecules from a sample plate. Ionization of the analyte occurs via proton acquisition in the gas phase. Virtually all ions formed are singly charged (8). The ions are removed from the source region by a large electric field and accelerated toward a field-free flight tube. Most MALDI experiments are performed in the positive mode, in which positively charged ions are extracted and accelerated toward a much lower potential. Ions migrate down the flight tube at different speeds where they are detected with an electron multiplier. Ions with greater mass take longer to reach the detector. The time to reach the detector gets correlated to a specific mass. Normally, MALDI TOF is used for the mass measurement of large molecules like proteins, nucleotides, and polymers (7). MALDI has been used to measure such samples with molecular weights from lkDa to several hundred thousand kDa. However, this case calls for the measurement of molecules with less than 200Da molecular weights. 28 COOH \. ,IOH .-\Y\KCN lvj “UL H0 2,5-dihydroxybenzoic acid (0H3) or-cyano-4-hydroxyclnnamlc acid (CHCA) :Il:/N H2 OH 110, l \T "(LN/CH3 \ x” r HS/J\N¢N no, 2,4-dlnitrophenylhydrazlne 6-aza-thlothymine (ATT') Figure 2.2. Structures of some common matrix molecules that have been used in MALDI experiments and an uncommon matrix, 2,4-dinitrophenylhydrazine, used for this study. 29 Problems can arise with conventional matrices, like those listed above, because the low mass region can be cluttered by matrix molecules, salts, and other contaminants (9). These form low mass ions that dominate the low mass region of the mass spectrum and make analysis of low mass analytes difficult. This can be remedied to an extent if a matrix is used that contributes very modestly to the low mass region of a spectrum. Therefore, an exotic matrix was used, 2,4-dinitrophenylhydrazine (DNPH), for SMM analysis, Figure 2.2. 2.3.2 Procedure for the Extraction / Purification of S-Methylmethionine from Plant Tissue with Ion Exchange Chromatography Both WT and —MMT Arabidopsis leaves were stored at -80°C until analysis. The leaves were then powdered under liquid nitrogen. About 0.5g of tissue was used for each analysis. The powder was added to a 15mL centrifuge tube. For quantitative purposes, 50 nmol of 2H6-SMM was added to each of the samples. 2mL of 12:5:1 methanolzchloroformzwater (MCW) was added to each. Both samples were shaken continuously for five minutes continuously, stored in an ice bath for five minutes, then shaken for an additional five minutes. The samples were then centrifuged at 4800rpm for 10 minutes at 4°C. The green supernatant was saved and placed on ice. The procedure was applied a second time to the remaining residue in the centrifuge tube. The supernatant was pooled with the first repetition resulting in about 4mL of MCW extract. lmL of SmM hydrochloric acid was added to the leaf tissue that remained at the bottom of the centrifuge tube. The mixture was boiled in a hot water bath (100°C) for about 5 minutes. It was centrifuged again and the supernatant (HCl extract) was saved. 30 This procedure was repeated again and pooled with the supernatant from the first trial. There was about 2mL of HCl extract from each sample. lmL of chloroform and 1.5mL of water was added to the pooled MCW supernatant. The solution partitioned into two distinct liquid phases. The lower organic phase remained green. The top aqueous layer was clear and colorless. After centrifugation, top layer was decanted and pooled with the HCl extract. A Pasteur pipette was used to remove as much of the aqueous phase as possible without removing any of the chloroform layer. The pooled extract was lyophilized down to a minimal volume, ~O.1mL. Ion exchange was then used to purify SMM from the extract. The extract was applied to a solid phase extraction (SPE) column containing AGl affinity resin. SMM was washed from the AG] column with 15mL of water. The eluent was allowed to drip directly into a second SPE column that contained BR70 affinity resin. 15mL of 1N HCl was added to elute SMM from the BR70 column. The eluent was dried down completely and reconstituted in about 0.5mL of water. This solution was applied to a third SPE column containing AG50 (N Hf). The column was washed with 15mL of ice cold water, followed by elution of SMM, and 2H6-SMM, with 3mL of 3M NH40H. The eluent was dried down completely and reconstituted in 0.1 to 0.2mL of 0.1% fluorosilic acid. The purified SMM samples were analyzed immediately and stored at 4°C for future analysis. This protocol was performed for both the WT and —MMT samples in triplicate. 31 2.3.3 Procedure for the Measurement of S-Methylmethionine Levels from Arabidopsis Extracts with Mass Spectrometry MALDI TOF was used to measure the SMM levels in the Arabidopsis extracts. 2, 4-dinitrophenylhydrazine (DNPH) was prepared as a saturated solution in acetonitrile (0.4mg / 0.1mL). It was centrifuged for 30 seconds with a micro-centrifuge and the supernatant was used as the matrix solution. 1 pL of sample was applied to a sample well on a stainless steel MALDI plate, followed by l 11L of DNPH matrix. The spot was allowed to air dry. The measurements were made with a Voyager Elite MALDI TOF instrument (PerSeptive BioSystems) in the linear mode. The sample spot was irradiated with a nitrogen laser (337nm) under vacuum. Ions formed were accelerated toward a field free flight tube with an applied accelerating voltage of 20kV. The grid wire voltage was set at 93% and the guide wire at 0.1% of the accelerating voltage. 2.3.4 Results: S-Methylmethionine Levels in Wild Type and Mutant (—MMT) Arabidopsis thaliana Figure 2.3 illustrates two representative MALDI TOF spectra of the WT and - MMT plants respectively. In each case, ZHo-SMM was detected at m/z 170 indicating the extraction and purification protocol was successful. Furthermore, and most importantly, it appears that the elimination of the SMM cycle in the mutant population was successful (1). The WT plants were grown without any genetic manipulation and should be capable of synthesizing SMM under normal growth conditions. As Figure 2.3 shows, the extraction of SMM from the WT plant tissue resulted in a positive response for SMM at m/z 164. 32 25‘ 164 170 20- WT ' 25.4 170 20‘ -MMT Relative Intensity (x 103) Figure 2.3. Representative MALDI MS spectra of AG 50(NH4+) ion exchange fractions from wild type and —MMT Arabidopsis thaliana. Modified from ref. 1. 33 The mutant plants, -MMT, were grown under the same conditions as WT, but there was no presence of m/z 164 in the —MMT mass spectrum in Figure 2.3. The stable isotope labeled SMM, 2H6—SMM, was used to prove that the extraction protocol worked. However, was also used as an internal standard for quantifying SMM. This is unnecessary for the —MMT plants, but it would provide a baseline level for WT plants grown under optimal conditions. This value could be used to compare to future studies when SMM levels are determined for plants grown under stress. By comparing the peak ratios between the internal standard peak, m/z 170, which corresponds to 50nmol of ZHG-SMM, to the peak representing endogenous SMM, m/z 164, it was determined that WT Arabidopsis leaf tissue contains 41 i 5nmol of free SMM (1). 2.4 Determination of S-adenosylmethionine (SAM) and S-adenosylhomocysteine (SAH) from Plant Tissue This section covers some background information of SAM and SAH followed by the experimental strategy and results. Included is information about the structures of each, some stability issues, and efforts put forth by other investigators to quantify SAM and SAH from biological samples 2.4.1 The Molecular Structures of S-Adenosylmethionine and S- Adenosylhomocysteine The structures of both SAM and SAH are shown in Figure 4. As mentioned, SAM is synthesized from ATP and methionine by the catalytic activity of SAM synthetase. SAH is produced as a byproduct of methyltransferase reactions (10). 34 Structurally, they are very similar. They could most basically be described as modified adenosine nucleosides. Nucleosides are precursors to nucleotides which make up nucleic acids. A pentose sugar makes up the foundation of the structures of nucleosides. In this case, the pentose sugar is the furanose form of D-ribose. A nitrogenous base forms a glycosidic bond to the ribose ring. The identity of the base gives the nucleoside its identity and name. When the nitrogenous base is adenine, the nucleoside is called adenosine. Nucleotides are formed upon esterification of a phosphate group to a nucleoside at the 5’ position of the ribose ring. In the case of phosphorylation of adenosine, the resulting nucleotide is adenosine-5’-monophosphate (AMP), Figure 2.4. One or two more phosphates can be added onto the initial phosphate via a phosphoric anhydride linkage. These are termed the corresponding di- and triphosphate nucleotides. Adenosine -5’-triphosphate (ATP) serves as the precursor to SAM, and eventually SAH. As described, SAM is formed via the transfer of the adenosyl portion of ATP to methionine and is catalyzed by SAM synthetase. The structure of SAM includes two chiral centers, Figure 2.4. One is at the sulfonium center and the other is at the alpha carbon of the methionine residue. The biologically active form of SAM, for methyl transfer reactions, is (S,S)-SAM (11). Stereoisomerization at the sulfur position leads to the inactive for (R,S)-SAM. Although inactive, (R,S)-SAM has can bind to methyltransferases with similar affinity as (S,S)- SAM. The R,S isomer therefore represents an inhibitor of methyltransferase enzymes. Only the methyl group on SAM distinguishes the structures of SAM and SAH, Figure 2.4. In fact, they are so structurally similar, that SAH has about the same affinity to the SAM binding site of methyltransferases that SAM does (12). This makes SAH a 35 Nfi-methyladenosine NH2 N \ 1 along the a axis are ejected. From Figure 4.4, this corresponds to q = 0.908. ,8, = [a + 2:] (4.6) The process of ejecting ions from the trap for analysis of m/z has been called mass-selective axial instability ejection. Since a = 0, only q can be used to eject ions and this is done by ramping the amplitude, V, of the rf potential on the ring electrode to the [3,, = l boundary and sequentially render ions of a given m/z to an unstable trajectory in order of low to high mass. As V increases, q of a particular m/z ratio increases until it reaches 0.908 (l3Z = 1) and the ion is ejected and detected. Further development of ion traps has employed the use of a supplementary rf potential that is applied between the end cap electrodes. The frequency of the supplementary rf potential is half that of the rf applied to the ring. As ions approach q = 0.908, ion motion resonates with the supplemental rf and causes an axial excursion of the resonant ion from the rest of the ion 92 0.2 .. 0.1 ‘- _0_l .i . —0.2 ¢ - -0.3 . . -0.4 ‘ ’ _0_5 .. n —0.6 - - —0.7 Figure 4.4. Quadrupole ion trap stability diagram. Adapted and modified from ref. 22. 93 cloud at the center of the trap. This process is called axial modulation and results in greater resolution. Specific waveforms can be used to select ions to be trapped, thus ejecting all unwanted ions. Ions are most often isolated for determining structural information by tandem mass spectrometry. This technique causes fragmentation of the isolated ion by increasing its energy with an applied rf that resonates with its motion, or secular frequency, in the trap. This resonant excitation is enough to cause collision-induced dissociation (CID) when the excited ion collides with a helium atom in the trap. The fragments that remain charged are confined in the trap and analyzed sequentially using the mass-selective axial instability ejection process discussed before. 4.4.3 Ion Cyclotron Resonance (ICR) ICR has emerged in the last few years as a mass spectrometric technique (23, 24). The fundamental operation of an ICR mass spectrometer was deve10ped from knowledge of the characteristic motion of ions (cyclotron motion) in a static magnetic field, B. The force exerted on an ion of mass m with charge q and velocity v is q(v x B). The cross product of v and B indicates the force of the magnet exerted on the ion is perpendicular to the plane of the velocity and magnetic field vectors. Therefore v can be represented as yxy, if the axial direction is represented by the z axis, Figure 4.5. This outward force is balanced by an inward, centrifugal force given as m V.,}..Z/r. Equating the magnetic force to the centrifugal force, equation 4.7, reveals a relationship between a mass to charge ratio, m/q, and a characteristic radius in the ICR cell, equation 4.8. 94 Receiver Plates (Detection) Transmitter Plates (Excitation) Trapping Plates Figure 4.5. Illustration of a typical cubic ICR cell in a FTMS instrument. Modified from ref. 23. 95 ”W” = (15,..73' (4.7) I” m5... r = _: (4.8) (18 This indicates that ions of a particular m/q ratio have characteristic, circular orbits of radius r in a static magnetic field. More importantly, the ions have characteristic frequencies, called cyclotron frequencies ((1),), which is dependent upon the m/z ratio of the ions. According to equation 4.9 below, the cyclotron frequency decreases with increasing mass—to-charge ratio. a) = l: = —E— (4.9) m m / q The cell itself typically has a cubic or cylindrical geometry. For descriptive purposes, a cubic cell will be considered here but the theory is the same. Figure 4.5 offers a visual example of a typical cubic cell. Ions can be generated inside the cell, but for our experiments, ions were generated externally and then transferred to the cell. Ions enter the cell along the z-axis and are confined axially with high potentials (~1-5V) on the trapping plates (see Figure 4.5). Detection and measurement of the ions present in the cell necessitates excitation of ions of a particular m/z to higher radial orbits. Cyclotron motion of ions have characteristic frequencies, 00,. The magnitude of the frequency depends upon the m/z value, but typically lie in the radiofrequency, rf, region (kHz-MHz). If the frequency of an applied rf voltage is applied to the ICR cell, to the transmitter plates, is resonant to the frequency of the ions cyclotron motion, then those ions will be excited to higher orbital radii. The amplitude of the rf is adjusted such that the ions pass close enough to a pair of 96 receiver plates as to induce a current of electrons attracted to the positive charges of the excited ions. As the ions continue to move to the other side of the cell in their excited cyclotron motion, they approach in close vicinity to the other receiver plate. To counter this charge, electrons move from the one receiver plate to the other. The frequency of the electron motion between the two plates reflects the frequency of the ion “excited” cyclotron motion. These frequencies are measured and recorded as an “image current”, Figure 4.6. This experiment will not work if the ion motion is incoherent. Ions with the same m/z values enter the cell, adopt a particular orbital trajectory of radius r, but have random phases. Ions of a given m/z value must be bunched into an ion packet. Otherwise, some ions will be close to one receiver plate while other ions of the same m/z value will be closer to the other. The result is no net frequency of motion. Therefore, ions in the cell are subjected to an azimuthal electric field (two-dimensional electric field) prior to resonant excitation which creates ion packets with coherent cyclotron motions (same phases). Frequency sweeps can be used to excite all m/z values sequentially to higher orbitals for detection. The result is an image current that contains all of the frequencies that were detected superimposed onto each other. The frequencies detected are characteristic of particular m/z values. The amplitude of each frequency detected reflects the abundance of the ions in the cell. Fast Fourier transform deconvolutes the frequency components, and amplitude information, into mass (m/z) and ion abundance information. A plot of ion abundance versus m/z constitutes the corresponding mass spectrum. The 97 Acquired Transient Signal fli—p Mass Spectrum (Image Current) Transform Relative Intensity Relative Intensity 0 0.2 0:4 0:6 0.8 1.0 700 900 1100 1300 1500 1700 1900 Time (s) m/z Figure 4.6. Illustration of the conversion of an image current to a mass spectrum with application of a fourier transform. Adapted and modified from ref. 24. 98 application of a Fourier transform has prompted ICR MS to be named Fourier transform mass spectrometry (F TMS). Like ion traps, ions can be isolated in ICR cells for further experimentation. Ions can be ejected from the cell if its characteristic frequency is applied at high enough amplitude such that its orbital radius exceeds the dimensions of the cell. Frequencies can be applied, such as in sweeps, such that all unwanted ions are ejected. Ions that remain can be studied structurally by tandem mass spectrometry. This requires resonant excitation to increase the ions energy, and leaking into the cell an inert collision gas. Collisions cause fragmentation and the charged fragments can be analyzed with frequency sweeps as described before. 4.5 Proteolytic Degradation of Proteins Many biochemical techniques have been used in tandem with mass spectrometry. Electrophoresis, for example, has been used to separate cellular proteins for analysis by mass spectrometry. Another technique that has received extensive utility with mass spectrometry is the degradation of proteins to smaller peptides with proteolytic enzymes and other reagents. This is an important tool for determining the identity of proteins as well as studying post-translational modifications. A brief description of these reagents will be discussed here with emphasis on those used in this project. Further discussion will be dedicated to the analysis of protein digestion products with mass spectrometry. Trypsin is the most commonly used protease used in the fields of protein science and proteomics. This proteolytic enzyme cleaves proteins at the amide backbone on the C-terminal side of arginine (R) and lysine (K) residues. R and K residues are fairly 99 prevalent in protein amine acid compositions and often result in peptides with 1000 to 2000 Da masses, although larger masses can be observed. Glu-C (from Staphylococcus aureus) is another protease that cleaves at the amide backbone, but on the C-terminal side of glutamate (E) and aspartate (D) residues. These are also fairly prevalent residues and result in peptides similar in size to those formed in tryptic digests. Cyanogen bromide (CNBr) is a non-enzymatic reagent that cleaves the amide backbone on the C-terminal side of methionine (M) residues. M residues are not common so, CNBr treatment often results in larger peptides as compared to trypsin and Glu-C. For most applications, this is a limiting property of CNBr. However, this characteristic proved invaluable to our method development. 4.6 Mass Mapping Mass mapping has been developed as a simple, yet effective, technique to identify proteins. Proteolytic digestion of a proteins yields peptide products that are specific to that protein. The presence of peaks representing these specific products in mass spectra provide adequate information to identify the protein, if the protein is known and its sequence exists in a databank. The protein is digested with trypsin and the peptide products are analyzed with mass spectrometry. The data is entered into a computer database to search for a protein sequence that would yield the same theoretical products as those observed in the mass spectrum. The protein that theoretically fits the data is assigned as the identity of the protein. 100 4.7 Peptide Sequencing with Mass Spectrometry Another approach to identifying proteins involves accurate amino acid sequencing of proteolytic digestion products (peptides). Often, protein amino acid sequences are so unique that complete sequencing of one peptide can identify a protein. If the single peptide sequence matches a sequence somewhere within the complete protein sequence of a protein in a database, then identification can be achieved. Success increases when more peptides from the digest are sequenced. Peptides are sequenced with mass spectrometry via CID experiments (or tandem mass spectrometry), which were discussed above. Collisions between peptide ions, MC, and reagent gases, N, (inert gases like argon or helium) activate the ions by converting some of its translational energy into internal energy (forming the excited species [Ml']+). Over a short period of time, the energy of the peptide ion is redistributed and the weakest bond breaks (Equations 4.10 and 4.11). M1++N -—>[Ml*]++N (4.10) [Mi"‘]+ —> M2++M3 (4.11) The fragmentation results in a product ion, Mf, and a neutral product M3. Since molecules and ions have Boltzmann energy distributions, several fragmentation products can be observed. By convention, the peptide being collisionally activated is termed the precursor ion and the products termed product ions. Product ions can contribute valuable structural information about the precursor ion. CID experiments with peptides have contributed to a readily employed technique for peptide sequencing. Emperical observations of peptide CID experiments have shown that the product ions most commonly observed are those that correspond to bond 101 breaking along the amide backbone. Three common sites of bond cleavage along the amide backbone can be observed and Figure 4.7 illustrates these possibilities. Bond cleavage between the or-carbon of a residue and a carbonyl group represents one possibility. Assuming the precursor ion was singly charged, then one of the fragments will retain a single charge, while the other fragment will be neutral (Scheme 1). If the charge remains with the N-terminal fragment, then the product ion is labeled an. If the charge remains with the product ion on the C-terminal side of the cleavage site, then the product ion is labeled xn. Likewise, if the cleavage site is between a carbonyl and amine group of the amide backbone then there are two fragmentation products. If the charge remains with the N-terminal fragment, then this product ion is labeled bn. If the charge remains with the C-terminal fragment, then the product ion is labeled yn. A similar logic is applied to cn and zn ions which result when the amide backbone is cleaved between an amine group and an or-carbon. When amino acids are condensed to form amide bonds and peptide sequences, the amino acids are called “residues”. The reaction results in loss of water from the amino acid mass. Therefore, residue masses are calculated as the amino acid mass minus the mass of water (-18 Da). Table 4.] lists the masses of the amino acid residues common in protein sequences. The only residues that don’t apply to those listed in Table 4.1 are those that reside on the N and C-termini (see Figure 4.7). The mass of a residue on the N-terminus is one mass unit larger than that listed in Table 4.1 due to a hydrogen atom attached to the or-carbon amino group of that residue. The C-terrninal residue will have a mass that is 18 mass units larger than those listed in Table 4.1. This is due to a hydroxyl group attached to the carboxyl of the or-carbon on the C-terminal residue. 102 NH ,NH | H,N NH H ‘OH 0 R 2 4 ‘ a2 \ b2 02 R1 //or R, 0 H2 H FRN‘ \\W/ 7//\\OH 0 R2 0 R, b2 Y2 Figure 4.7. (A) Illustration of cleavage sites along the amide backbone of peptides and product nomenclature. (B) Examples of bn and yn structures. Adapted from ref. 5. 103 Table 4.1. One and Three Letter Codes of the 20 Common Amino Acids and Corresponding Residue Masses. Modified from ref. 5. amino acid nominal (3/1 letter codes) residue mass alanine (Ala IA) 71 arginine (Arg/R) 156 aspartic acid (Asp/ D 115 asparagine (Asn/N) 1.14 - cysteine (Cys/C) 103 glutamic acid (Gin/E) 129 glutamine (Gln/Q) 128 glycine (Gly/G) 57 histldine (His/H) 137 isoleucine (Ile/I) 113 leucine (Lou/L) 113 lysine (Lys/K) 128 methionine (Met/M) 131 phenylalanine (Phe/F) 147 proline (Pro/P) 97 serine (Ser/S) 87 threonine (Thr/T) lOl tryptophan (T NV) 186 tyrosine (T yr 163 valine (V al/V) 99 104 Tandem mass spectra (MS/MS spectra) can yield sequence information of the peptide in question. This is done by piecing a series of ions together that differ in mass by corresponding residue masses in the sequence. Figure 4.8 illustrates an example of this process. Manual interpretation of MS/MS spectra in this manner has been called de novo sequencing. Problems arise when a peptide preferentially fragments in one or only a few sites. When this happens, no sequence information can be gained. This has been shown to be more common when the peptide sequence include proline or aspartic acid residues. Common fragmentation products observed are preferential cleavage of the peptide on the N-terminal side of the proline residues and the C-terrninal side of aspartic acid residues. This indicates that not all peptide bonds have the same propensity to break in low-energy CID experiments. 4.8 Tandem Mass Spectrometry and Post-Translational Modifications While certain residues can inhibit the usefulness of an MS/MS spectrum by limiting the sequence information available, the problem is more profound when the peptide is modified. Post-translational modifications (PTMs) occur on proteins and gives them filnctional capability. The most commonly studied PTM has been phosphorylation. Protein phosphorylation is a reversible modification and believed to be involved in the regulation of protein activity and cellular processes. In order to assess this, it is necessary to determine where these phosphorylation sites are. Phosphorylation sites are most commonly phosphate esters formed on the side chain hydroxyl group of threonine, serine and tyrosine residues. 105 100 Relative Abundance '5 8 8 8 ‘6 8 5‘ 8 8 0 Internal b ion ProArg+ / y8 y7 Y6 y5 Y4 y3 y2 y1 842 743 672 571 472 385 272 175 VATVSLPR 100 171 272 371 458 571 668 824 bl b2 b3 b4 b5 b6 b7 b8 y2/b3 CID on [M + 2H]2+ = 421.5 m/z Figure 4.8. Tandem mass spectrum of precursor ion of m/z 421.5 (+2 charge of peptide VATVSLPR). De novo sequencing was used to identify several b and y ions that were used to sequence the peptide. Adapted from ref. 5. 106 Tandem mass spectrometry has been used to determine sites of protein phosphorylation. Previous studies have shown that phosphorylated peptides preferentially lose the phosphate group in low-energy CID experiments. Corresponding MS/MS spectra display a dominant peak that represents the neutral loss of the phosphate group as phosphoric acid. An MS/MS spectrum of a phosphorylated peptide displays the dominant neutral loss peak, but little other information can be gathered. Although these peptides may be almost impossible to sequence, the characteristic neutral loss of phosphoric acid can be used to locate phosphopeptides from a mixture of peptides (from a digest for example). Neutral losses from protonated molecules, MH+ correspond to a decrease in mass by 98 Da. For multiply protonated molecules, such as [MHz]2+ and [MH3]3+, losses of phosphoric acid correspond to an m/z decrease of 49 and 32.7 Da relative to the parent ion mass respectively. This information can be used to sort out of a list of MS/MS spectra those that may correspond to phosphopeptides. 4.9 Conclusion This completes the background discussion of instrumental background and biochemical principles that will be applied to the development of a protocol for identifying sites of pyridoxal-l-phosphate modification of proteins. The discussion was a simple review of pre-existing techniques that have been used for similar investigations in the past. The following section introduces a new application of these techniques for studying more exotic PTMs like PLP. 107 4.10 Chapter 4 References l. 10. ll. 12. Zenobi, R., Knochenmuss, R. Ion formation in MALDI mass spectrometry. Mass Spec. Reviews. 1998, 17, 337-366. Kebarle, P. A brief overview of the present status of the mechanisms involved in electrospray mass spectrometry. J. Mass Spectrom. 2000, 35, 804-817. Gaskell, S.J. Electrospray: principles and practice. J. Mass Spectrom. 1997, 32, 677-688. Zhu, H., Bilgin, M., Snyder, M. Proteomics. Annu. Rev. Biochem. 2003, 72, 783- 812. Aebersold, R., Goodlett, D.R. Mass spectrometry and proteomics. Chem. Rev. 2001, 101, 269-295. Wilkins, M.R., Gasteiger, E., Gooley, A.A., Herbert, B.R., Molloy, M.P., Binz, P.A., Cu, K., Sanchez, J .C., Bairoch, A., Williams, K.L., Hochstrasser, D.F. High- throughput mass spectrometric discovery of protein post-translational modifications. J. Mol. Biol. 1999, 289, 645-657. Ficarro, S.B., McCleland, M.L., Stukenberg, P.T., Burke, D.J., Ross, M.M., Shabanowitz, J ., Hunt, D.F., White, RM. Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nature Biotech. 2002, 20, 301-305. Posewitz, M.C., Tempst, P. Immobilized gallium(III) affinity chromatography of phosphopeptides. Anal. Chem. 1999, 71, 2883-2892. Schlosser, A., Bodem, J ., Bossemeyer, D., Grummt, I., Lehmann, W.D. Identification of protein phosphorylation sites by combination of elastase digestion, immobilized metal affinity chromatography, and quadrupole-time of flight tandem mass spectrometry. Proteomics. 2002, 2, 911-918. Tholey, A., Reed, J ., Lehmann, W.D. Electrospray tandem mass spectrometric studies of phosphopeptides and phosphopeptide analogues. J. Mass Spectrom. 1999, 34, 117-123. Adamczyk, M., Gebler, J .C., Wu, J. Selective analysis of phosphopeptides within a protein mixture by chemical modification, reversible biotinylation and mass spectrometry. Rapid Commun. Mass Spectrom. 2001, 15, 1481-1488. Hayashi, H., Wada, H., Yoshimura, T., Esaki, N., Soda, K. Recent topics in pyridoxal-5’-phosphate enzyme studies. Annu. Rev. Biochem. 1990, 59, 87-1 10. 108 l3. 14. 15. l6. l7. l8. 19. 20. 21. 22. 23. Hoffman, E.D., Charette, J., Stroobant, V. Mass Spectrometry: Principles and Applications. John Wiley & Sons, New York, 1996. Posthumus, M.A., Kistemaker, P.G., Meuzelaar. Laser-desorption-mass spectrometry of polar nonvolatile bio-organic molecules. Anal. Chem. 1978, 50(7), 985-991. Karas, M., Gluckmann, M., Schafer, J. Ionization in matrix-assisted laser desorption/ionization: singly charged molecular ions are the lucky survivors. J. Mass Spectrom. 2000, 35, l-12. Wiley, W.C., McLaren, I.H. Time-of—flight mass spectrometer with improved resolution. Rev. Sci. Instr. 1955, 26(12), 1150-1157. Cotter, R.J. The new time-of-flight mass spectrometry. Anal. Chem. 1999, 71, 445A-451A. Chemushevich, I.V., Ens, W., Standing, K.G. Orthogonal-injection TOF MS for analyzing biomolecules. Anal. Chem. 1999, 71, 452A-461A. Whittal, R.M., Li, L. Time-lag focusing MALDI-TOF mass spectrometry. American Laboratory. 1997, 29(24), 30-36. Guilhaus, M., Mlynski, V., Selby, D. Perfect timing: time-of—flight mass spectrometry. Rapid Commun. Mass Spectrom. 1997, 11, 951-962. March, R.E. Quadrupole ion trap mass spectrometry: theory, simulation, recent developments and applications. Rapid Commun. Mass Spectrom. 1998, 12, 1543- 1554. March, R.E. An introduction to quadrupole ion trap mass spectrometry. J. Mass Spectrom. 1997, 32, 351-369. Marshall, A.G., Hendrickson, C.L., Kackson, G.S. Fourier transform ion cyclotron resonance mass spectrometry: a primer. Mass Spec. Rev. 17: 1-35 (1998). 24. Amster, J .I. Fourier transform mass spectrometry. J. Mass Spectrom. 31: 1325-1337 (1996). 109 CHAPTER 5 TANDEM MASS SPECTROMETRY OF DEOXY-PYRIDOXAL-L-PHOSPHATE MODIFIED PEPTIDES 5.1 Introduction: Neutral Loss Products in Mass Spectrometry Mass spectrometry (MS) has been used not only for molecular mass calculations, but also structural determinations of molecules (1). Electron impact (EI) ionization is a well-established example of a technique that can provide both molecular weight and structural attributes of molecules (2). The interaction of neutral molecules with 70eV electrons imparts enough energy into the molecule to not only ionize it, but cause subsequent fragmentation. The fragmentation process can be so extensive that little if any of the molecular ion is left to be detected. However, the wide distribution of fragment ions that form provide extensive information about the structure of the molecule. The behavior of different classes of molecules in an El MS experiment has been well classified. For example, alcohols have been shown to fragment in such a way that they lose a neutral molecule of water (3). The neutral loss of 18 mass units from a molecule in E1 MS has therefore been used as an indicator of the presence of a hydroxyl group in organic compounds. Neutral loss determinations with mass spectrometry have been extended not only for determining substituents in organic molecules, but determining prosthetic groups and other modifications in peptides (4, 5). Peptides are too large and nonvolatile to be studied with E1 MS, so tandem mass spectrometry with ESI and MALDI ion sources are used. However, El MS serves as an effective analogy to tandem mass spectrometry in 110 that we can get both molecular weight and structural information. Ions formed from the source first get sorted by a mass analyzer and selected ions of a given m/z value can be isolated (or trapped) and induced to fragment via collisions with an inert gas, as opposed to an electron (6, 7). Unmodified peptides fragment in such a way that the dominant products correspond to fragmentation along the amide backbone (4). The result is a series of ions in a mass spectrum that differ by the mass of an adjacent amino acid residue. The series can be used to determine the amino acid sequence of the peptide. Not all amide bonds have the same propensity to fragment so the products have different levels of abundance (which is reflected in the corresponding mass spectrum). In the case of modified peptides, such as phosphorylation of serine, threonine, and, to a lesser extent, tyrosine, the labile bond between the phosphate and the residue is more susceptible to fragmentation than amide bonds so the product representing the loss of the modification (as phosphoric acid) is usually the major peak in the corresponding mass spectrum (5, 8). Tandem mass spectra of phosphopeptides are dominated by a peak representing the neutral loss of phosphoric acid. Often, few other peaks are present. This makes sequencing these peptides difficult, if not impossible. However, a dominant peak representing the neutral loss of 98 Da provides a solid indicator that the peptide ion precursor was a phosphopeptide. Similarly, the neutral loss of 18 Da from an organic molecular ion in E1 is a strong indicator of an alcohol. Tandem mass spectrometry has become a major contributor to locating sites of phosphorylation in proteins (5, 8). The behavior of peptides modified by pyridoxal-l-phosphate (PLP) in tandem mass spectrometry has not yet been studied. In order to determine whether a peptide is modified with PLP, it is important to know if the loss of the modification represents the 111 dominant product or whether fragmentation along the amide backbone is observed. This chapter summarizes our efforts to characterize the fragmentation behavior of peptides modified by PLP. 5.2 Background and Nomenclature for Peptides Modified by Pyridoxal-l-Phosphate PLP forms Schiff bases, or aldimine bonds, with primary amines (9). Figure 5.1 illustrates the formation of a Schiff base upon the reaction of a peptide (F ibrinogen, for example) with PLP. It is a dehydration reaction in resulting in loss of water (9). This occurs simply by mixing PLP and the peptide together. The peptide must have a primary amine available, so a lysine residue must be part of the peptide amino acid sequence. The reaction is reversible and ideal conditions call for a pH environment buffered at about 8 (9). Furthermore, a buffer lacking primary amines is necessary to prevent side reactions. Acidic conditions favor the dissociation of the Schiff base and this presents a problem when separating peptides with liquid chromatography and analyzing with MALDI or ESI mass spectrometry since both require the use of acidic reagents. However, reduction of the aldimine bond by sodium borohydride, NaBH4, is an irreversible reaction that yields a secondary amine linkage between PLP and the peptide (10). This prevents the loss of PLP under any pH conditions. The ultimate result of the reduction of the aldimine bond is peptide modification by PLP minus oxygen. Therefore, we will call reduced, PLP modified peptides: deoxy-pyridoxal-l-phosphate peptides (dPLP peptides). The peptide Fibrinogen was used as the model for comparing the MS/MS spectral characteristics of dPLP modified versus unmodified peptides. The amino acid sequence of Fibrinogen is HHLGGAKQAGDV. A listing of amino acid letter designations is 112 H-H-L-G-G-A-K-Q-A-G-D-V H-H-L-G-G-A-K-Q-A-G-D-V Fibrinogen (MW = 1188 g/mol) ' H20 I (MW = 18 glmol) /N o / OH OH H204P \ H20,P \ l i \N/ CH 3 \N/ CH3 Pyridoxal-I-Phosphate (PLP) NaBH4 (MW I 247 gimol) H-H-L-G-G-A-IT-Q-A-G-D-V /NH OH H20,P ' \ \N/ CH3 dPLP-Modified Fibrinogen (FPLP) (MW = 1419 amu) Figure 5.1. The condensation reaction between the peptide Fibrinogen and pyridoxal-l- phosphate (PLP) followed by reduction with sodium borohydride (N aBH4). 113 included in Table 5.1 along with residue masses of each. The sequence of F ibrinogen includes a lysine (K) residue which can be modified by dPLP. For simplicity, dPLP modified F ibrinogen will be written as FPLP. 5.3 Procedure: Synthesis of dPLP Modified Peptides F PLP was synthesized by a modification of the procedure used by Morino et a] (l 1). A stock solution of F ibrinogen was prepared by dissolving 0.5mg of the peptide in 0.3mL of SOmM (N -[2-Hydroxyethyl]piperazine-N’-[2-ethanesulfonic acid]), (HEPES), buffered to pH 7.8. A saturated solution of PLP was prepared in the same buffer. 1M sodium borohydride, NaBH4, was also prepared in HEPES buffer. Only the Fibrinogen stock solution was saved for future experiments (stored at 4°C). All other solutions were prepared fresh daily. 5 pL of Fibrinogen stock was added to 300p.L of HEPES buffer in a 1.5mL micro-centrifuge vial. 511L of 1M NaBH4 was added to the mixture. The vial was placed in darkness and allowed to react for 45 minutes at room temperature. Two drops of concentrated acetic acid was added to stop the reaction. The pH at this point was about3. The solution was dried down to about IOOuL. F PLP and unmodified F ibrinogen were then purified with High Performance Liquid Chromatography, HPLC. A Reliasil C18 reversed phase column with the dimensions 2.0mm x 150mm was used. The column contained 300A particles with a pore size of 5 pm. Two mobile phases were used for gradient elution. Mobile phase A consisted of 95% water, 5% acetonitrile, and O. 1% trifluoroacetic acid (TFA). Mobile phase B consisted of 5% water, 95% acetonitrile, and 0.1% TFA. The mobile phase gradient began at 5% B and increased to 65% B at 30 114 Table 5.1. One and Three Letter Codes of the 20 Common Amino Acids and Corresponding Residue Masses. Modified from ref. 5. amino acid nominal (3/1 letter codes) residue mass alanine (Ala IA) 71 arginine (Arg/R) 156 aspartic acid (Asp! D 115 asparagine (Asn/N) 1.14 - cysteine (Cys/C) 103 glutamic acid (Glu/E) l 29 glutamine (Gln/Q) 128 glycine (Gly/G) 57 histidine (His/H) 137 isoleucine (Ile/I) 113 leucine (Leu/L) 113 lysine (Lys/K) 128 methionine (Met/M) 131 phenylalanine (Phe/F) 147 proline (Pro/P) 97 serine (Set/S) 87 threonlne (Thr/T) 101 tryptophan (T NV) 186 tyrosine (T yr 163 valine (Val/V) 99 115 minutes. A steeper gradient increased mobile phase B to 95% at 32 minutes where it remained for 3 minutes. The composition of B then decreased to 5% (at 38 minutes into the run) where it remained though the remainder of the run. Total time of the run was 45 minutes. The flow rate throughout was 65 pL/minute. Peptides were detected with a UV- visible detector set at 214nm. 5.4 MALDI Experiments Figure 5.2 displays the HPLC chromatogram illustrating the separation of F ibrinogen and FPLP. Fractions corresponding to both HPLC chromatographic peaks were collected together (or pooled together) as a mixture for analysis with mass spectrometry. Initially, MALDI TOF was used to analyze the HPLC purified mixture of F ibrinogen and FPLP. MALDI TOF allows for quick analysis and could be used for assessment of the NaBH4 reaction to determine if dPLP modification of Fibrinogen was achieved. Figure 5 .1 displays the molecular masses of the products and reactants of the modification reaction. Measurements were made with a PerSeptive Biosystems Voyager- DE STR MALDI TOF instrument in linear mode. Typical settings were applied including 20000V accelerating voltage, grid voltage at 95% of the accelerating voltage, guide wire was set to 0.05% of the accelerating voltage, and the delay time was 100 nsec. A nitrogen laser (wavelength 337 nm) was used for desorption/ionization of the sample. luL of sample was co-deposited onto a designated spot of a stainless steel sample plate with 1 11L of a saturated solution of 2,5-dihydroxybenzoic acid (DHB) as the matrix. DHB was prepared by saturation in a 50:50 mixture of water and acetonitrile. 116 1001 80- co 2‘ o g, 8 E = 3', 40- N *5 8 ( = 20- cc .8 -___ .9. .1: < o I l V 10 13 16 19 Time (min.) Figure 5.2. HPLC purification of unmodified and dPLP-modified Fibrinogen (FPLP). 117 MALDI Fourier Transform Mass Spectrometry (F TMS) was used for high mass accuracy and resolution of the F ibrinogen and FPLP mixture as well as MALDI tandem mass spectrometry experiments. These experiments were performed with a Bruker Daltonics Apex II MALDI FTMS instrument. The instrument was equipped with a ScouthO MALDI source (337 nm nitrogen laser). Ions were transferred from the source to an ion-cyclotron resonance cell enclosed within a 7T superconducting magnet for detection. Tandem mass spectrometry was performed by isolation of selected ions in the cell and reaction (C1D) with argon gas. 5.5 Electrospray Experiments All ESI experiments were performed with a ThermoFinnigan LCQDECA. Fibrinogen and FPLP were separated with HPLC which was interfaced directly to the LCQDECA. HPLC was performed with a Ultrafast Microprotein Analyzer (Michrom BioResources Inc.). About 20uL of sample was injected into the HPLC. The same column (described above) was used for the LC-MSn experiments as the one used for fraction collection and MALDI TOF analysis. The mobile phase and gradient were changed. Mobile phase A was composed of 5% acetonitrile, 95% water, and 0.1% formic acid while mobile phase B was composed of 95% acetonitrile, 5% water, and 0.1% formic acid. The mobile phase composition began as 5% B at the start. It was maintained that way for the first two minutes, then it was increased to 80% B by the 40 minute mark. It was maintained there for five minutes and then reduced to 5% B again over another five minutes. It was maintained at 5% B for the duration of the run. The 118 total time of the run was 60 minutes. The flow rate was maintained at 75 pL/minute throughout. Peptides eluting from the HPLC were immediately analyzed by the LCQDECA. Peptides were converted to gas phase ions by electrospray and transferred to an ion trap mass analyzer. Mass-to-charge ratios of ions were determined over time as peptides eluted from the HPLC. MS/MS experiments were carried out according to data- dependent settings. A low-limit abundance threshold was set. Any ion more abundant than the threshold could be isolated and induced to fragmentation by CID. The most intense peak was selected first and induced to fragmentation. It was set to perform MS/MS on the same m/z value three times in a row. After that, the instrument isolated the second most intense peak for MS/MS experimentation and so on. 5.6 Results Fibrinogen was reacted with PLP in the presence of NaBH4 resulting in a mixture of dPLP modified F ibrinogen (FPLP) and unmodified Fibrinogen. Figure 5.3 displays a MALDI TOF spectrum of the HPLC purified mixture. The protonated, monoisotopic mass (MH+) of Fibrinogen is 1189.61 Da while FPLP is 1420.46 Da. The mass of FPLP corresponds to the peptide mass plus 231 Da from the addition of dPLP. The limited mass resolution of MALDI TOF in this spectrum is apparent. The masses detected are actually average masses. Nonetheless, the information provided from the spectrum indicates that the reduction reaction with NaBH4 was successful in modifying some of the peptide. The large peak at m/z 1190.86 Da corresponds to the unmodified F ibrinogen peptide. The peak at m/z 1420.80 Da is about 231 Da larger than m/z 1190.86 Da and corresponds to 119 45000 - 1190.86 >5 #5 30000 - fl .8 E. a 3 § 15000 - § 1211.92 r 1326.00 1420.80 T l l l I I 1100 1150 1200 1250 1300 1350 1400 1450 1500 m/z Figure 5.3. A MALDI TOF spectrum of the HPLC purified mixture of F ibrinogen and dPLP-modified Fibrinogen (FPLP). 120 the mass of F ibrinogen plus the addition of dPLP (FPLP). The relative intensity of FPLP versus Fibrinogen is small. There are a few possible explanations for this. One, the peak intensities reflect the true populations of the peptides. This would mean that the reduction reaction resulted in a low yield of FPLP. It could also mean that the signal for FPLP is suppressed. Studies have shown that phosphorylated peptides can be very difficult to detect with MALDI (12). Since PLP includes a phosphate group in its structure, it’s not unreasonable to assume similar behavior from dPLP-peptides with MALDI. The two other peaks in Figure 5.3 can also be accounted for. The peak at m/z 1211.92 is 21 Da larger than the assigned Fibrinogen peak and most likely a represents a sodium adduct of F ibrinogen. This should result in a mass difference of 22 Da (+23 Da for Na, - 1 Da for the proton replaced), but the error could be attributed to the technique. Normally, if one peak representing a protonated ion has a complementary sodium adduct peak at 22 m/z values higher, then it would be expected that all of the peaks representing protonated ions would have a complementary peak. In this case, they may be below the detection limit. The peak at m/z 1326.00 Da could be attributed to the attachment of pyridoxal, a metabolic precursor in the pathway to PLP formation in which the phosphate group is absent on the molecule as phosphoric acid (H3PO4). The mass is 94 Da smaller than the modified peptide FPLP. A mass reduction of 98 Da could be attributed to loss of phosphoric acid. It should be noted that the formation of this side-product did not interfere with any of the experiments. For MS/MS experiments, all unwanted ions were ejected prior to excitation and fragmentation of the isolated, target ion (FPLP). Therefore, the side-product was not given any filrther attention. 121 Figure 5.4 is an FTMS spectrum of the same sample as in Figure 5.3 (Fibrinogen and FPLP). The same peaks are observed. The peak at m/z 1189.38 can be assigned to Fibrinogen as before. The peak at m/z 1420.79 is 231.41 mass units larger than Fibrinogen and represents dPLP-modified Fibrinogen (FPLP). With FTMS, more certainty is gained in the identity of the ion represented by m/z 1322.79. This peak is exactly 98 mass units below FPLP which would come from pyridoxal. 5.7 Tandem Mass Spectrometry with MALDI FTMS2 The ions represented by the peak at 1420.79 (Figure 5 .4) correspond to dPLP- modified F ibrinogen (FPLP). Before examining the nature of the fragmentation of FPLP, a control experiment was performed with unmodified F ibrinogen. Figure 5.5 is an MS/MS spectrum of F ibrinogen (m/z 1 189.3 8) obtained with MALDI FTMSZ. A tabulated list of peaks and peak assignments is included. The dominant product ions are b, y, and internal ions (b and y ions were described in chapter 4). Internal ions result when the amide backbone is broken at two different locations resulting in a peptide ion that lacks both the C and N-terminal residues of the precursor peptide (4). The point of this experiment was to show that Fibrinogen can be easily sequenced using tandem mass spectrometry (Figure 5.5). Figure 5.6 is a MALDI FTMS2 spectrum of FPLP (m/z 1420.8). In contrast to unmodified Fibrinogen, the MS/MS of dPLP-modified Fibrinogen results in no interpretable sequence information. The spectrum only shows the m/z range1055 to 1355 to highlight the important area where peaks were observed. No peaks were observed below this range. The two most intense peaks in the spectrum are m/z 1189.7 and 1322.8 122 16000000- 1189.4 12000000 ~ 2' '0 5 8000000 4 E 4000000 _ Amass = 231 Da PI 1211.7 13223 1420.8 0 -L J“ .L Iil . .11. I I Al I I 1 1160 1210 1260 1310 1360 1410 1460 mlz Figure 5.4. MALDI FTMS spectrum of the mixture of peptides Fibrinogen and dPLP- modified F ibrinogen (FPLP). 123 yll 3'10 3'9 3’0 3’7 Yo Y5 3'4 3'3 3’2 3'1 HHLGGAKQAGDV b, b, b b, b, b, b, b b b", 11,, bll 3’7 3’11 Intensity y 10 y, y‘ y. 1 b3) b8 200300400600600700800900100011001200 m/z Figure 5.5. A tandem mass spectrum of F ibrinogen obtained with MALDI FTMSZ. The sequence of Fibrinogen is displayed above the spectrum, with an illustration of the b and y ions that could result with fragmentation along the amide backbone of the peptide. 124 900000 - 1189.7, [MH — dPLP]+ 600000 - 1322.8, [MH — H3PO‘]+ 300000 - Absolute Intensity , 1.11111“ I I . Alli . . J, 1205 1255 1305 mlz : lhmll‘lllmr 1055 1105 1155 Figure 5.6. A tandem mass spectrum of dPLP-modified F ibrinogen (FPLP), (MH’), obtained from MALDI FTMSZ. 125 respectively. The peak at m/z 1322.8 is 98 mass units smaller than the precursor ion (m/z 1420.8). This was attributed to the neutral loss of phosphoric acid (H3PO4) from the dPLP group of F PLP. The base peak of the spectrum was 1189.7, which is about 231 mass units smaller than the precursor peptide ion. This is the same mass as the unmodified Fibrinogen peptide indicating the loss of the entire dPLP group. According to the MALDI data discussed above, MS/MS spectra of dPLP- modified peptides are dominated by two characteristic neutral loss peaks. One peak represents a neutral loss of 98 mass units corresponding to H3PO4. The other common neutral loss was 231 mass units corresponding to the neutral loss of dPLP. MALDI is limited in that only singly charged precursors are generated from the ion source (13). Ultimately, for this project, proteins would be digested, separated with liquid chromatography, and directly analyzed by ESI mass spectrometry. Therefore, it was necessary to assess the fragmentation patterns of multiply-charged precursor ions of FPLP as well. 5.8 Tandem Mass Spectrometry with ESI-MS2 The sample containing F ibrinogen and FPLP was injected onto a reversed phase HPLC column. Figure 5.7 is a full scan base peak ion chromatogram illustrating the separation of the two peptides. A base peak chromatogram is simply a time-dependent plot of the intensity of the most abundant peak (base peak) in each full scan spectrum. Figure 5.8 displays a full scan spectrum from each of the chromatographic peaks at about 12.2 and 15.0 minutes respectively. In each of the spectra, the charge states of the ions representing the peptides Fibrinogen (Figure 5.8A) and FPLP (Figure 5.88) are present. 126 ul 0! a N co to 9 e e o o e g .h 6 IllllllllllllllllllllllllllLiJllJllllllllllLLlllllI Relative Abundance ‘10 to 6° 0 O FPLP (15.0 min.) Fibrinogen (12.2 min.) [IILITltirAf‘11IIT] 10 15 20 25 30 Time (min) Figure 5.7. A base peak ion chromatogram illustrating the separation of F ibrinogen and dPLP-modified F ibrinogen (FPLP). 127 595.6, [MH2]2" 100 E 397.7. [MI-1313+ 4 80j 1189.6. MH“ N O a 0 11111111411111 - 1 ll “ Al” ll I l n 11.1.11 A unnumnnnm 11 m 111 111‘1 ll Llflll 1 we, {may h e llllJlllllllllllllll 711.2. [MH,]2+ Relative Abundance 8 8: N O 4421; 882.1 1420.6, MH” "1 O A ‘ 1 ‘L ‘l' 11‘ .l-_-A A‘A-A-Au- mn- - A A AA AAA L AAA-mm AAALA- .- A n- AAIMA l'rTrU'IIIIII [fitillli'l"fiI't'v[IIr'l'I'IIIIUIIIT'IIIITrIrUIIIIUUIIIIIIIFIIIIIFIYIIUUII Figure 5.8. Full scan LC-MS spectra at retention times 12.2 minutes (A) and 15.0 minutes from the separation of F ibrinogen and dPLP-modified F ibrinogen (FPLP). Refer to the chromatogram in Figure 5.7. 128 These charge states served as the precursors for isolation and CID experiments. As before with the MALDI FTMS/MS experiments, it was necessary to gather data from the CID of F ibrinogen (unmodified) as a control. This would help assess the significance of the modification and charge state on the fragmentation patterns, relative to an unmodified peptide. Figure 5 .9 displays stacked MS/MS spectra of the +1, +2, and +3 charge states of Fibrinogen respectively. Figure 5.9A is the MS/MS spectrum of the singly charged precursor [MH+], (m/z 1189.6). Only b and y ions were annotated since they represent the dominant products. Almost all of the remaining peak assignments have been accounted for as internal ions. Figure 5.9B displays the MS/MS spectrum of the +2 precursor of Fibrinogen, [MH2]2+, (m/z 595.6). It should be pointed out that the interpretation of MS/MS spectra can get even more complicated when multiply charged precursors are involved. This is due to the fact that many of the product ions can be multiply charged as well. With this in mind, we can still get solid sequence information from this MS/MS spectrum (Figure 5.98). The annotated peaks again correspond to b and y ions since they represent the most abundant products. The peaks at m/z 527.00 and 536.93 are doubly charged products that represent y“ and b“ respectively. The singly charged analogues of these products were observed in Figure 5 .9A. The peak at m/z 578.02 in Figure 5.98 has not been assigned but appears to be a doubly charged analogue of m/z 1154.59 of Figure 5.9A, likewise unassigned. Figure 5.9C displays the MS/MS spectrum of the triply charged precursor of Fibrinogen, [MH3]3+, (m/z 397.37). Unlike the first two spectra, there is little sequence information available. Still, the dominant peaks can be rationalized if the charge states of the products are assumed. The peak at 129 bu 100 111724 Y” so he 10524 60 A 137 Y9 9155 7014 902 3 9 40 900-5 11,, 11716 1 1117-5. 1 .11 Yu b u no 5270 571mm) y, 9155 37113 (+2) / 5730 802.3 Y; b‘ 3'3 20 3701 489.2\ 573.3 1 745.5 1.... “11111.4.1..511 J1] 113 393.3 1, / 4591 2 (+2) b, Y6 7014 Yu L”517.2 / 1052.4 Relative Abundance O O 100 bu 537 0 (+2) b2 so 2751 I’ll bio 40 353-3 479.4 20 y“ \ (+31 (*2) 11L .1 J l 2331 "'l""l""l""l‘ .-,.4..,....,fi..,. . .. 1 . .. 200 400 600 800 1000 1200 ml: . vavv [vffi'fit Figure 5.9. CID spectra of Fibrinogen with different charge state precursors: (A) +1 charge, m/z 1 189.6 Da, (B) +2 charge, m/z 595.6 Da, and (C) +3 charge, m/z 397.7 Da. 130 m/z 537.01 could be assigned as a doubly charged b” ion while the peak at m/z 358.31 could be assigned as the triply charged b” analogue. The peak at m/z 479.41 is most likely the doubly charged blo ion and so on. The control CID control experiments of F ibrinogen served two purposes. One, it illustrated the ability of LC-MS2 for sequencing unmodified peptides. The MS/MS spectra in Figures 5.9A and 5.93 provided a lot of sequence information for Fibrinogen. Two, it also illustrated the role of the precursor ion charge state in the fragmentation patterns of peptides (4). The +3 charge state does not yield much information for sequencing the peptide even though the few peaks present can be accounted for. Sequencing the peptide would be impossible if only the +3 precursor MS/MS spectrum was available in this case. Peak assignments for the +1 and +2 precursor MS/MS spectra were challenging even though the sequence was already known. It should also be pointed out that the amino acid composition can play a role in the products formed in CID experiments. Aspartic acid (D) residues have been observed empirically to be preferential cleavage sites in CID experiments (4). Specifically, internal cleavage of the amide backbone on the C-terminal side of D typically results in a prominent bn product. The base peaks in both Figures 5.9A and 5.9C have been identified as b” typify this observation since this is the b ion on the C-terminal side of the aspartic acid residue. The b” product is also represented as one of the larger peaks in Figure 5.9B. Therefore, charge states and amino acid compositions are two things to keep in mind when trying to assess the effect of dPLP modification on the MS/MS fragmentation of peptides. The MS/MS spectra of the corresponding charge states of FPLP are displayed in Figure 5.10. Figure 5.10A is the MS/MS spectrum of the singly charged precursor, MH+, 131 100 Relative Abundance A 0 1322.5. [MH - H,Po,1+ 1189.8. [Ml-l - dPLPr A - .. .. .- ”-1“ -11 .u. 2 662.1, [MM2 - 11,130,}2+ B 695.4.[MI-l2-dPLPP‘ \ 4 A Illulllllllll‘ J ‘A All 1 ILL 11 44 441.8, [14111, - 1-1,1=o,13+ C 5/95.6. [MH,3+ - dPLPH*F* 1 1.1 ..1 ll- JIJJ 1 11 [rillifilllillllrllll 200 400 600 800 1000 1200 1400 ml: Figure 5.10. CID spectra of PLP-modified F ibrinogen (F PLP) with different charge state precursors: (A) +1 charge, m/z 1420.6 Da, (B) +2 charge, m/z 711.2 Da, and (C) +3 charge, m/z 474.6 Da. 132 while 5.108 and 5.10C are the doubly ([MH2]2+) and triply ([MH3]3+) charged precursor MS/MS spectra respectively. Contrary to the unmodified charge state precursors of Fibrinogen, few product ions are formed from the CID of all three charge states of F PLP. All three spectra are dominated by two peaks implying that the neutral losses of H3PO4 and dPLP are the dominant products. In Figure 5.10A, (MS/MS of m/z 1420.6, MH+) the two dominant peaks are m/z 1322.5 and 1189.6 Da. Relative to the precursor mass, 1420.6 Da, m/z 1322.5 Da is 98 Da smaller. This corresponds to the neutral loss of phosphoric acid ([MH — H3PO4]+). The peak at m/z 1189.6 Da is 231 Da smaller than the precursor ion mass. This mass difference corresponds to the neutral loss of dPLP ([MH — dPLP]+. This result agrees with the interpretation of the MS/MS spectrum observed for the CID experiment of F PLP with MALDI FTMS/MS (Figure 5 .6). Before discussing Figure 5.108 and 5.10C which are spectra of the multiply charged precursors of FPLP, it is helpful to first discuss some observations and nomenclature of tandem mass spectrometry of multiply charged phosphopeptides. As discussed, MS/MS spectra of phosphopeptides are characterized by a dominant peak that corresponds to the neutral loss of H3PO4. For a singly charged precursor, MH+, this corresponds to a mass difference of 98 Da. However, if a doubly charged precursor, [MH2]2+, is subjected to CID, loses H3PO4, and both charges are retained on the product ion, then the mass difference between the product and precursor ion masses will be 98/2 Da (49 Da). Similarly, if a triply charged, phosphorylated precursor is subjected to CID, loses H3PO4, and all three charges are retained on the product ion, then the charge on the product ion will be 98/3 Da (32.7 Da). When searching for MS/MS spectra that may represent phosphopeptides, it is common to search for neutral losses of 98, 49, and/or 133 32.7 Da as these are common for phosphopeptides, depending on the charge state of the precursor. Figure 5.11 illustrates these calculations. Since we now know that dPLP peptides yield two dominant neutral loss products in CID experiments, we can use the same logic to assign peaks in MS/MS spectra from multiply charged precursors. Since the loss of H3PO4 is a common product of the CID of dPLP peptides, we expect to see neutral losses of 98, 49, and 32.7 Da from +1, +2, and +3 precursors of dPLP peptides as well. The loss of dPLP is the other common neutral loss product of the CID of dPLP peptides. From a singly charged precursor, MH+, the neutral loss of dPLP corresponds to a mass of 231 Da. If the doubly charged dPLP peptide ion is subjected to CID, loses dPLP, and both charges are retained on the product ion, then the neutral loss mass is 231/2 Da (115.5 Da). Likewise, the neutral loss mass from the +3 precursor would be 231/3 Da (77 Da). Figure 5.12 illustrates these calculations. Figure 5.10 B displays the MS/MS spectrum of the doubly charged precursor of F PLP (m/z 711.2 D). The peak at m/z 662.1 Da is 49.1 Da below the precursor mass and corresponds to the neutral loss of H3PO4 ([MH; - H3PO4]2+) according the calculations described in Figure 5.11. The peak at m/z 595.4 Da is 115.8 Da below the precursor mass and corresponds to the neutral loss of dPLP ([MH; - dPLP]2+) as also described in Figure 5.11. Figure 10C displays the MS/MS spectrum of the triply charged precursor of FPLP (m/z 474.6 Da). The peak at m/z 441.8 Da is 32.8 Da below the precursor mass and corresponds to the neutral loss of H3PO4 ([MH3 - H3PO4]3+) (refer to Figure 5.11). If the triply charged precursor lost dPLP, and the product ion retained all three charges, then a neutral loss product 77 Da below the precursor mass should be observed. No ion was 134 (1) Relative m/z (2) Relative m/z (3) Relative m/z (4) Relative m/z MH+ —£D—> [MH — H3PO4]+ (M + 1)/1 {((M + 1)/1) —— (98/1)} [MH2]2+ -—w’—+ [MH2 — H3PO4]2+ (M + 2)/2 {((M + 2)/2) — (98/2)} [MH,]3+ —°2—> [MH3 — H3PO4]3+ (M + 3)/3 {((M + 3)/3) — (98/3)} [MHn]n+ i, [MHn— H3PO4]n+ (M + mm {((M + n)/n) — (98/n)} Figure 5.11 This figure describes singly and multiply charged precursors (M = phosphopeptides) undergoing CID and yielding neutral loss products corresponding to the loss of H3PO4. All neutral products (H3PO4) have been omitted for simplicity. 135 (1) war —9-I-D—> [MH — H3PO4]+ or [MH — dPLP]+ Relative m/z (M + 1)/1 {((M + 1)/1) — (98/1)} {((M + 1)/1) — (231/1)} (2) [M12]2+-—Cm—-> [MH2 - H,P 0,]2+ or [MH2 — dPLPP“ Relative m/z (M + 2)/2 {((M + 2)/2) — (98/2)} {((M + 2)/2) - (231/2)} (3) [MH,]3+—CID—> [MH3 — H3P 033+ or [MH, — dPLP]3+ Relative m/z (M + 3)/3 {((M + 3)/3) — (98/3)} {((M + 3)/3) — (231/3)} (4) [mm-55L» [MEL-Hapotl’” or [MEL—dPLPR Relative m/z (M + n)/n {((M + n)/n) - (98/n)} {((M + n)/n) — (23l/n)} r_1 1m.;H3P_0. 1* [MH.. — dPLPE 1 [(M +‘n)/n] — 98 [(M +' n)/n] — 231 2 [(M + n)/n] — 49 [(M + n)/n] — 115.5 3 [(M + n)/n] — 32.7 [(M + n)/n] — 77 Figure 5.12. (A) A representation of the CID of dPLP peptides with different charge states all yielding neutral loss product ions corresponding to neutral losses of H3PO4 and dPLP. (B) The observed relative losses in m/z value from loss of H3PO4 and dPLP from dPLP peptides with charge state n. 136 present according to the spectrum (Figure 5.10C). However, the next most abundant peak, m/z 595.6 Da, represented the neutral loss of dPLP from a doubly charged precursor in Figure 5.108. This could be a result if not all of the charges were retained. Therefore, the peak has been assigned as the doubly charged product ion corresponding to the neutral loss of dPLP ([MH3 — dPLPH+]2+). 5.9 Discussion and Conclusions Although tandem mass spectra of dPLP peptides provide no sequence information, they do contain characteristic spectral features that can be used to identify dPLP peptides. The two dominant features of MS/MS spectra of dPLP peptides represent the neutral losses of H3PO4 and PLP. This behavior has been observed with other post- translational modifications, most notably phosphorylation. Phosphoester bonds are labile relative to amide bonds and thus break easier. This explains the observation that CID experiments of phosphopeptides most often result in a base peak representing the neutral loss of H3PO4. Lehmann et al. proposed a mechanism for the loss of H3PO4 from phosphopeptides which is displayed in Figure 5.13A (phosphoserine) (5). They suggest that loss of H3PO4 proceeds through a six—centered transition state and the cyclic rearrangement of three electron pairs. Transfer of an aliphatic hydrogen atom to the phosphate group occurs during this cyclic electron pair rearrangement. Support for this mechanism comes form the observation that phosphotyrosine (pY) often does not lose a phosphate group. As seen in Figure 5138, pY lacks an aliphatic hydrogen atom to allow the proposed six-centered transition state. More fragmentation along the amide 137 Protein \ Hr H+ Protein Protein 7... A O Phosphoserlne Protein \NH H+ Protenn 7. O Pho sphotyro sine Figure 5.13. (A) Proposed six-centered transition state to explain the neutral loss of H3PO4 from a phosphopeptide precursor in tandem mass spectrometry. (B) Phosphotyrosine lacks an aliphatic hydrogen to complete a six-centered transition state. Neutral losses of H3PO4 are not observed. Adapted and modified from Ref. 5. 138 phosphate group. As seen in Figure 5138, pY lacks an aliphatic hydrogen atom to backbone has been observed for pY peptides than for phosphoserine and phosphothreonine, both of which have the aliphatic hydrogen in a favorable position for the six-centered transition state to be formed. We propose two mechanisms to rationalize the observation of the two characteristic neutral losses from dPLP peptides in tandem mass spectrometry. Figure 5.14 illustrates our proposal for the neutral loss of dPLP (loss of 231 Da) from a dPLP peptide precursor ion. Six atoms have been labeled 1 through 6 counterclockwise to illustrate a six-centered transition state. The electron pair from the O-H bond moves to the amine nitrogen atom at position 1 forming a hydride bond. Electron pairs then shift according to the illustration in Figure 5.14 resulting in double bonds between positions 2- 3 and 4-5. The double bond on the pyridine ring between positions 3-4 must break and form a ketone if this is indeed the mechanism of the neutral loss of dPLP. The structure of the product of the reaction in Figure 5.14 does account for the observed neutral loss mass of 231 Da. It would be a reasonable assumption that if the mechanism proposed by Lehmann et al. (5) for the neutral loss of H3PO4 (loss of 98 Da) is correct, then the loss of H3PO4 from a dPLP peptide might proceed through a similar mechanism. The Lehmann proposal suggests that an aliphatic hydrogen atom gets transferred to the phosphate group and subsequent electron pair rearrangement (through a six-centered transition state) results in the cleavage of the phosphoester bond (Figure 5.13A) releasing H3PO4. In this case, the structure of the dPLP group prohibits a six-centered transition state. However, if an eight-centered transition state is the true mechanism, then a transfer of an aliphatic hydrogen to the phosphate group is possible as illustrated in Figure 5.15. The 139 H9 231 Da Figure 5.14. The proposed mechanism for the neutral loss of H3PO4 (231 Da) from dPLP-modified peptides in tandem mass spectrometry. The secondary amine marked 8 is the terminal side chain amine from the modified lysine residue of the peptide. 140 H—CH K, CH3 HC CH3 TH o) \ \ \ P [Edi /N _’ + H07 \0 N How / 0 V lac/V HO HO 980a Figure 5.15. The proposed mechanism for the neutral loss of dPLP (98 Da) from dPLP- modified peptides in tandem mass spectrometry. The secondary amine marked a is the terminal side chain amine from the modified lysine residue of the peptide. 141 atoms that are proposed to participate in the eight-centered transition state are labeled 1- 8. The aliphatic hydrogen atom, labeled 8, belongs to the methylene group involved in the reduced linkage between dPLP and lysine. After the hydrogen atom is transferred to the phosphate group, electron pair rearrangement results in cleavage of the phosphophoester bond between positions 3 and 4. The product H3PO4 would account for the neutral loss of 98 Da from the dPLP peptide precursor. While these proposed fragmentation mechanisms offer logical explanations for the observed products, more experimentation is needed to deduce the true mechanisms of the neutral losses of H3PO4 and dPLP from dPLP peptides. Even mechanisms for the loss of H3PO4 from phosphopeptides are hypothetical at this point (5). Nonetheless, these experiments have established a CID pattern that can be used for identifying dPLP- peptides. 142 5.10 Chapter 5 References l. 10. ll. 12. Johnstone, R.A.W., Rose, M.E. Mass Spectrometry for Chemists and Biochemists. Cambridge University Press, New York, 1996, p. 325—396. De Hoffman, E., Charette, J ., Stroobant, V. Mass Spectrometry Principles and Applications. John Wiley & Sons, New York, 1996, p. 7-11. . Smith, R.M. Understanding Mass Spectra. Ed. Kenneth L. Busch. Jonh Wiley & Sons, New York, 1999, p. 137-138. Aebersold, R., Goodlett, D.R. Mass spectrometry in proteomics. Chem. Rev. 101: 269-295 (2001). Tholey, A., Reed, J ., Lehmann, W.D. Electrospray tandem mass spectrometric studies of phosphopeptides and phosphopeptides analogues. J. Mass Spectrom. 34: 117-123 (1999). Shukla, A.K., Futrell, J.H. Tandem mass spectrometry: dissociation of ions by collisional activation. J. Mass Spectrom. 35: 1069-1090 (2000). de Hoffmann, E. Tandem mass spectrometry: a primer. J. Mass Spectrom. 31: 129- 137 (1996). DeGnore, J .P., Qin, J. Fragmentation of phosphopeptides in an ion trap mass spectrometer. J. Am. Soc. Mass Spectrom. 9(11): 1175-1188 (1998). Dixon, H.B.F., Karpeisky, M.Y. Vitamin Bé Pyridoxal Phosphate Chemical, Biochemical, and Medical Aspects, Part B. Edited by David Dolphin, Rozanne Poulson, and Olga Avramovic. John Wiley & Sons, New York. p. 71—117 (1986). Morino, Y., Nagashima, F. Vitamin B9 Pvridoxal Phosphate Chemical, Biochemical, and Medical Aspects, Part A. Edited by David Dolphin, Rozanne Poulson, and Olga Avramovic. John Wiley & Sons, New York. p. 477-497 (1986). Sugiyama, Y., Mukohata, Y. Modification of one lysine by pyridoxal phosphate completely inactivates chloroplast coupling factor 1 ATPase. FEBS Lett. 98: 276 (1979) Asara, J .M., Allison, J. Enhanced detection of phosphopeptides in matrix assisted laser desorption/ionization mass spectrometry using ammonium salts. J. Am. Soc. Mass Spectrom. 10: 35-44 (1999). 143 l3. Karas, M., Gluckmann, M., Schafer, J. Ionizationin matrix-assisted laser desorption/ionization: singly charged molecular ions are the lucky survivors. J. Mass Spectrom. 35: 1-12 (2000). 144 CHAPTER 6 THE DEVELOPMENT OF A METHOD FOR LOCATING DEOXY- PYRIDOXAL-L-PHOSPHATE MODIFIED PEPTIDES USING TANDEM MASS SPECTROMETRY AND NEUTRAL LOSS ANALYSIS 6.1 Introduction Tandem mass spectrometry of deoxy- pyridoxal-l-phosphate (dPLP) modified peptides results in two characteristic neutral losses. One has been identified as a neutral loss of phosphoric acid and the other as a neutral loss of dPLP. These neutral loss products dominate MS/MS spectra of these peptides and no sequence information can be obtained. Now that this has been established, the next step was to develop a strategy for locating sites of PLP modifications in proteins. The development of two approaches will be described here. Both approaches begin by stabilizing the PLP modification in the protein by sodium borohydride reduction. The next step is to digest the protein with a site-specific protease, breaking down the protein into smaller pieces (peptides) that can be separated by HPLC and analyzed. Two different mass spectrometric strategies will be explored to analyze these peptides. One will be mass mapping in which the peptides are analyzed with MALDI TOF and correlating the masses detected to a sequence within the protein (1). The protein sequence is available, and the digestion products can be predicted (including the dPLP peptide), so theoretical masses can be used to match MALDI TOF data to a peptide sequence. The other approach was to analyze the protein digest with LC-MS/MS and look for a spectrum that exhibits the characteristic neutral loss pattern of a dPLP peptide. 145 Whether using mass mapping or the tandem mass spectrometry approach, it was imperative to be able to predict the masses of peptides from proteolytic digestions. For mass mapping, a mass can only be matched to a peptide if the theoretical mass is known. For tandem mass spectrometry, observing a MS/MS spectrum that displays the characteristic spectral features of a dPLP peptide is useless if it doesn’t match any of the theoretical possibilities for dPLP modified peptides. Trypsin is the most commonly used protease for protein analysis using mass spectrometry and it was applied in this investigation. Trypsin cleaves amide bonds of proteins on the C-terminal side of lysine (K) and arginine (R) residues. This specificity allows us to determine the sequences of resulting peptides and calculate the predicted masses of all proteolytic peptides. As noted earlier, once a lysine residue has been modified by dPLP, it is no longer recognized as a cleavage site by trypsin (2). This must be taken into account when determining the peptides formed and calculating the corresponding masses. Figure 6.1 illustrates a general example of what would be expected. In the unmodified protein (column 1) trypsin cleaves at all of the predicted sites (K and R residues). The digest results in peptides with m/z 901.1, 1013.1, and 2047.3. Matching the peaks in a mass spectrum (bottom) to those in a theoretical list, such as proteins in a database, is the general basis to mass mapping. If one of the cleavage sites is blocked by a modification, such as dPLP, then we would not see the same three mass spectrometric peaks as column 1. Instead, we would observe two peaks: one for peptide with m/z 901.1 and a peak at higher m/z representing the sum of the peptide m/z values 1013.1 and 2047.3, the mass of the dPLP (231), minus a water molecule and a proton. If a precursor peptide ion is subjected to CID and the resulting MS/MS spectrum exhibits neutral loss 146 l 2 dPLP I YFVS EGFELFCAQSFSKN- YFVSEGFELFC AQSFSKN- FGLYNERV GNLTVVAK F GLYNERV GN LTVVAK l .‘ .................. Trypsm .................. ’ l dPLP YFVS EGFELFC AQSF SK, W (013.) = 2047.3 . Y'FVS EGFELFC AQSF SKN- NFGLYNER, 4431(ng = 1013.1 FGLYNER. mm.) = 32 72-5 VGNLTVVAK, VGNLTVVAK, Wflwg.) = 901.1 1MB+ (avg) = 901.1 901.1 5‘ | 1013.1 3. 901 1 o o . a» -... . > .— 3. :5, 20473 ‘ Contsrasung Mass '3 3 3272.5 a """"" t IIIIIIII o If”: .5 pet ra Di 5 | m/z m]: Figure 6.1. This figure demonstrates the application of mass mapping to identify modifications in proteins. 147 features characteristic to dPLP peptides, then the precursor mass can be compared to theoretical peptide masses to determine which one carries the modification. 6.2 TIP39 Peptide: The Model for Method Development In order to assess our strategy for locating dPLP peptides by peptide mass mapping and tandem mass spectrometry with neutral loss analysis, we synthesized a small dPLP-modified protein to test it on. TIP39 is the model we chose. Figure 6.2A displays the sequence of this protein. This protein was chosen because it is small and simple. It contains only one lysine residue that we can modify with PLP, yet it also contains several arginine residues that can be cleaved by trypsin. The potential modification site is lysine-32 (K-32). With TIP39 as the developmental model, we set out to modify the protein, digest it with trypsin, and determine the site of the modification. 6.3 Procedure for the Synthesis of dPLP-modified TIP39 dPLP-modified TIP39 (TIP39PLP) was prepared in much the same way as modified Fibrinogen (FPLP) in Chapter 5 (3). lOOug of TIP39 was dissolved in SOOuL of saturated PLP containing SOmM HEPES buffered at approximately pH 8. The solution was left at room temperature for 15 minutes in the dark. lOuL of 1M NaBH4 was added and the mixture was incubated for 45 minutes at room temperature in the dark. The NaBH4 reaction was halted with an addition of two drops of concentrated acetic acid. The pH of the solution was about 4.5 after the acid was added. 148 A TIP39 Amino Acid Sequence SLALADDAAF'” RERARLLAALZ" ERRHWLNSYM” HKLLVLDAP” Predicted Digestion Products of TIP39 Seguence Mass (avg) 1 6—22 786.0 1 6-23 942.2 1-1 1 1 1 50.3 24-32 121 6.4 23-32 1372.6 1-13 1 435.6 24—39 1 938.3 23-39 2094.5 *24-39 21 69.5 ‘23-39 2325.7 * dPLP modified, K-32 Figure 6.2. This figure displays the amino acid sequence of TIP39 (A) along with the theoretical tryptic digestion products of TIP39 (B). 149 The sample was dried down to approximately IOOuL and injected onto an HPLC column for purification. It should be noted that we were hoping to separate TIP39PLP from remaining TIP39, but our main goal with HPLC was to remove residual reagents, NaBH4 and PLP, as well as salts that may have been endogenous to the TIP39 sample. A Reliasil C18 reversed phase column with the dimensions 2.0mm x 150mm was used. The column contained 300A particles with a pore size of 511m. Two mobile phases were used for gradient elution. Mobile phase A consisted of 95% water, 5% acetonitrile, and 0.1% trifluoroacetic acid (TFA). Mobile phase B consisted of 5% water, 95% acetonitrile, and 0.1% TFA. The mobile phase gradient began at 5% B and increased to 65% B at 30 minutes. A steeper gradient increased mobile phase B to 95% at 32 minutes where it remained for 3 minutes. The composition of B then decreased to 5% (at 38 minutes into the run) where it remained though the remainder of the run. Total time of the run was 45 minutes. The flow rate throughout was 65 uL/minute. Peaks were detected with a UV-visible detector set at 214nm. Fractions, corresponding to chromatographic peaks, were collected, analyzed with mass spectrometry (MALDI TOF), and dried down to a minimal volume (< 5 11L). As it was, the separation of the two forms was not successful according to the chromatogram (not shown) so both TIP39 and TIP39PLP were collected together and analyzed as a mixture. 6.4 Procedure for Tryptic Digestion of dPLP-Modified TIP39 The sample was then reconstituted in 0.1mL of 25mM ammonium bicarbonate (pH 7.8). 0.5 ug of trypsin was added and the mixture was incubated at 37°C for 18 150 hours. After incubation, one drop of concentrated acetic acid was added to stop digestion (to minimize autoproteolysis of trypsin). 6.5 Mass Spectrometric Analysis of the dPLP-Modified TIP39 Tryptic Digest MALDI TOF was used to analyze the digest before and after HPLC separation of the peptides. Measurements were made with a PerSeptive Biosystems Voyager-DE STR MALDI TOF instrument in linear mode. Typical settings were applied including 20000V accelerating voltage, grid voltage at 95% of the accelerating voltage, guide wire was set to 0.05% of the accelerating voltage, and the delay time was 100 nsec. A nitrogen laser (wavelength 337 nm) was used for desorption/ionization of the sample. luL of sample was co-deposited onto a designated spot of a stainless steel sample plate with 1 11L of a saturated solution of 2,5-dihydroxybenzoic acid (DHB) as the matrix. DHB was prepared by saturation in a 50:50 mixture of water and acetonitrile. The digested TIP39PLP sample was also analyzed by LC-MSn which was performed with a ThermoFinnigan LCQDECA ion trap mass spectrometer. About 20uL of sample was injected into the HPLC. The same column (described above) was used for the LC-MS" experiments as the one used for fraction collection and MALDI TOF analysis. The mobile phase and gradient were changed. Mobile phase A was composed of 5% acetonitrile, 95% water, and 0.1% formic acid while mobile phase B was composed of 95% acetonitrile, 5% water, and 0.1% formic acid. The mobile phase composition began as 5% B at the start. It was maintained that way for the first two minutes. It was then increased to 80% B by the 40 minute mark, then it was maintained tat 80% B for five minutes and then reduced to 5% B again over another five minutes. It 151 was maintained at 5% B for the duration of the run. The total time of the run was 60 minutes. The flow rate was maintained at 7511L/minute throughout. Peptides eluting from the HPLC were immediately analyzed by the LCQDECA. Peptides were converted to gas phase ions by electrospray and transferred to an ion trap mass analyzer. Ion m/z values were determined over time as peptides eluted from the HPLC. MS/MS experiments were carried out according to data-dependent settings. A low-limit abundance threshold was set. Any peak more intense than the threshold could be isolated and induced to fragmentation by CID. The most intense peak was selected first and induced to fragmentation. It was set to perform MS/MS on the same m/z value three times in a row. After that, the instrument isolated the second most intense ion for MS/MS experimentation and continued to the third most abundant ion. 6.6 Results and Discussion Figure 6.3 displays a MALDI TOF spectrum of the HPLC purified mixture of TIP39 and dPLP-modified TIP39 (TIP39PLP). The molecular weight of TIP39 is 4504 Da. TIP39PLP has a molecular weight of 4735 Da with the addition of dPLP (231 Da). Therefore, the peak at m/z 4502.5 in Figure 6.3 has been assigned to TIP39 and m/z 4733.2 has been assigned to TIP39PLP. The peak at m/z 4637.4 is about 96 Da less than the peak assigned to TIP39PLP. This most likely represents the modification of TIP39 by pyridoxal (a degradation product of PLP lacking phosphoric acid). The peaks at m/z values 4600.0 and 4817.8 were not assigned. The spectrum confirmed that some of the protein had been modified by dPLP and further experimentation could be carried out. Figure 6.2 displays the sequence of TIP39 along with a list of theoretical peptide 152 70000 — 60000 — 4sozs,ntnr 50000 — 40000 4 30000 — 4733.2, [1m + aprr 20000 — 10000— WM 0 11.1.2. 1 I 1 l l 4200 4400 4600 4800 5000 5200 m/z Relative Intensity Figure 6.3. The MALDI TOF spectrum of the mixture of TIP39 (MH+) and TIP39PLP ([MH + dPLPY). 153 products from tryptic digestion. The list includes digestion products with and without annotated according to the sequence they represent within the overall sequence of the protein. The peak assignments were designated by matching m/z values in the spectrum to theoretical masses listed in Figure 6.2B (mass mapping). Peaks that correspond to dPLP peptides are marked with an asterisk. Multiple dPLP peptide digestion products were observed as the result of skipped cleavage sites by trypsin. According to the sequence, trypsin should cleave on the C—terminal side of R-23 resulting in peptide 24-39. If modified, this peptide has a theoretical, protonated, monoisotopic mass of 2169.5 Da. A peak at m/z 2169.6 was observed in Figure 6.4. However, if trypsin cleaves on the C- terminal side of R-22 and skips R-23 then peptide 23-39 would result. This has a theoretical, protonated, monoisotopic mass of 2325.7 Da. A peak at m/z 2326.0 has also been observed in Figure 6.5. Fractionation of the peptides with HPLC resulted in the chromatogram in Figure 6.5. All of the fractions corresponding to chromatographic peaks were collected and analyzed with MALDI TOF. Each peak in the chromatogram has been annotated according to the sequence within TIP39 that it represents. These assignments were based on matching peaks in the spectra to theoretical masses of peptides in Figure 6.28 (mass mapping). The mass spectra corresponding to the last two chromatographic peaks in Figure 6.5, marked 24-39' and 23-39', (22.3 and 22.8 minutes respectively) indicate that these peaks represent both dPLP peptides previously discussed. The MALDI TOF spectrum of peak 22.3 minutes is displayed in Figure 6.6. The appearance of a peak at m/z 2170.8 confirms that this is the dPLP-modified peptide 24-39. The mass spectrum also shows that the unmodified peptide 24-39 has co-eluted with the modified counterpart 154 70000 — 1 11 24-32 60000 4 ' 23-32 §‘ 50000 - = *2 40000 - .3 24-39 23-39 .2 30000 - E m/z 2169.6 & 20000 ~ . 1-13 “'39. ' 111/: 2326.0 23.-39* 1500 2000 2500 Figure 6.4. MALDI TOF spectrum of the tryptic digestion products of the mixture TIP39 and TIP39PLP. The peaks are annotated according to the sequence they represent. * represents products modified by dPLP. 155 S E 500 t “'32 E 16-23 23.32 g 400 b 11 2459* g 300 - 3339 \ 2,39. 1-13 - o 200 - 1 0 5 100 ' 16-22 1 . .1 . 2 12 15 18 21 24 27 Retentlon Time (mln) Figure 6.5. The HPLC chromatogram displays the separation of peptides from the tryptic digestion of TIP39 and dPLP-modified TIP39. The peaks are annotated according to the sequence they represent within TIP39. The peaks marked with * correspond to peptides modified by dPLP. 156 30000 - 1939.8, MH“ 25000 ~ 3* § 20000 - ‘5 3 15000 - g 1962.2 7": 10000 _ 21703,[1m+ dprr :4 5000 _ J 2075.4 WM 0 l l l l 1700 1900 2100 2300 2500 m/z Figure 6.6. MALDI TOF spectrum corresponding to the chromatographic peak marked 24-39‘ from Figure 6.5. 157 as indicated by the presence of a peak at m/z 1939.8. Figure 6.7 displays the MALDI TOF mass spectrum of peak 22.8 minutes. The peak at m/z 2328.2 matches the theoretical mass of dPLP-modified peptide 23-39. As in the fraction at 22.3 minutes, the co-elution of the unmodified peptide 23-39 has occurred at 22.8 minutes too. It also appears that there is still some of the modified and unmodified 24-29 peptide still eluting off the column at this point. Figure 6.8 demonstrates the use of mass mapping for assigning peaks to sequences within TIP39 and TIP39PLP. This is a real example of the generic explanation of mass mapping in Figure 6.1. Only residues 16 to 39 of TIP39 are displayed for simplicity. Column A represents the tryptic digestion of unmodified TIP39. Assuming that missed cleavages are observed, the product peptides will be 16-22, 23—32*, 24-32, 23-39*, 24-39*, and 33-39 (* indicates a missed cleavage). Column B illustrates the peptides that would be present if TIP39PLP (modified at K-32) was digested with trypsin. Since K-32 is blocked by PLP, this is no longer a recognized tryptic cleavage site. Therefore, peptides 24-32 and 23-32* would no longer be observed in a mass spectrum. Instead, a peak representing the masses for 24-32, 33-39, and dPLP would be observed. The same could be said for 23-32*, 33-39, and dPLP. Figure 6.8 takes into account that missed cleavages were observed at K-32 even for the unmodified TIP39. This figure helps to clarify how mass mapping can be used to determine the sites of post—translational modifications. The TIP39PLP digest was also analyzed by LC-MSZ. Figure 6.9A displays a base-peak ion chromatogram of the digest. As peptides elute off of the column, they are measured with a full scan of the ion trap mass spectrometer. The more abundant ions are 158 50000 1 2097.2, ME 40000 - 30000 - 1939.8 20000 - Relative Intensity 10000 ‘ 2328.2, [MH + amp 0 .W' 1 1800 1900 2000 2100 2200 2300 2400 2500 m/z Figure 6.7. MALDI TOF spectrum corresponding to the chromatographic peak marked 24-39‘ from Figure 6.5. 159 A B dPLP LLAALEWWLNS YMHISLLVLDAP LLAALEWWLNS YMHI'CLLVLDAP l < Trypsin > (16-22) LLAALER (23-32) RHWLNSYMHK (16-22) LL11 ALER dPLP (“'32) HWLNSYMHK (23-39)* RHWLNSYMHK LLVLDAP (33-39) LLVLDAP .1pr (23 39) RHWLNSYMHKLLVLDAP (24‘3” HWLNSYMHI'CLLVLDAP (24-39) HWLNS YMIIKLLVLDAP A = Cleavage site “ = dPLP modified peptide Figure 6.8. A comparison of the predicted peptides that would be formed from the digestion of TIP39 (column A) versus dPLP-modified TIP39 (column B). 160 1°" Base Peak Ion Chromatogram 80 60 40 324 20 '\ 33.0 a o M 2.; g 100 80 3 so Neutral Loss of 32.7 Da g 40 20 I“ a o 1. AA ‘5‘. 3 100 I! 00 00 Neutral Loss of 77 Da 40 20 0 l1 .1. 1 11 OIJITTIJIIIIIIIIIITII llllélllll Igllll 1111 11111111111111] 10 2 2 3 3 50 55 60 Time(min) Figure 6.9. (A) A displays of the base peak ion chromatogram of the LC-MS2 experiment of the tryptic digestion products of the mixture of TIP39 and dPLP-TIP39. (B) and (C) display neutral loss chromatograms of 32.7 and 77 Da respectively. 161 isolated and MS/MS experiments are performed. A list of MS/MS spectra get stored for future interpretation. The software (Excalibur) has the capability of searching through this list of MS/MS spectra and locating the specific spectra corresponding to the neutral loss inquiries. Figure 6.98 and C are neutral loss chromatograms from the same run in which the search was for the neutral loss of 32.7 Da and 77 Da corresponding to the loss of H3PO4 and dPLP respectively from a +3 precursor. Chapter 5 described the correlation between the charge state of the precursor and the neutral loss masses observed for H3PO4 and dPLP (Figure 5.128). This indicates that the two small chromatographic peaks at 32.4 and 35.0 minutes (in Figure 6.9A) represent the two dPLP peptides. The conclusion that the peaks in the neutral loss chromatograms (Figures 698 and C) are dPLP peptides could only be made if a response was observed at the same retention time for both neutral losses (H3PO4 and dPLP). The CID experiments on dPLP peptides (described in Chapter 5) established a characteristic neutral loss pattern in which the dominant peaks in the mass spectra corresponded to the neutral losses of H3PO4 and dPLP. None of the dPLP peptides studied with tandem mass spectrometry exhibited just one of the two neutral loss features in the mass spectra. Both were always observed, with few other peaks present in the spectra. Therefore, if a peak was present in Figure 6.93 (- 32.7) at time t, then a peak should also be present at time t in Figure 6.9C (-77) if CID was performed on a dPLP peptide ion at time t. Figure 6.10A and 6.108 displays full scan ESI spectra at chromatographic retention times of 32.6 and 34.8 minutes respectively. Both spectra contain peaks at m/z values corresponding to the +2 and +3 charge states of dPLP-modified peptides 162 1085.6, [MH2]2* A 723.7, [MH,]3+ a? a a S 15 ...... “7+. . :1" .+,~.+r ‘1. .H. Talmuvfl—r: L. . z: . 1 . .‘w . .fi $200 400 600 800100012001400160018002000 '43 1163.0, [MH,]2+ E 04 775.9, [MH,]3+ B MALTMLQ I? .32 .- .._ is 1.1 200 400 600 800 1000"" 1200 1400 1600 1800 2000 2 Figure 6.10. LC-MS full-scan spectra corresponding to retention times 32.6 and 34.8 minutes (A and B respectively). Spectrum A displays the +2 and +3 charge states for dPLP-modified peptide 24-39 and Spectrum B displays the same for dPLP-modified peptide 23-39. 163 (sequences 24—39 and 23-39 respectively). The neutral loss chromatograms from the corresponding +2 precursors are not shown but also can be used to determine the MS/MS spectra that reflect the neutral loss patterns that are characteristic of dPLP peptides. The neutral loss chromatograms (Figures 698 and 6.9C) suggest that the two broad, low intensity peaks at 32.4 and 35.0 minutes correspond to peptides that are modified by dPLP. The MS/MS spectra of the +3 precursors of both dPLP-modified peptides are shown in Figure 6.11A and 6.1 13 respectively. The two spectra look almost identical so only the spectrum in Figure 6.11A will be discussed, but will apply to both. There are many more peaks in both MS/MS spectra than what was observed in the MS/MS spectra of FPLP (Chapter 5). However, the characteristic neutral losses were present and are annotated in the spectra. A search was performed for neutral losses of 32.7 Da and 77 Da corresponding to neutral losses of H3PO4 and dPLP from a +3 precursor ion respectively. The peak at m/z 691.2 represents a triply charged ion and loss of 32.6 Da. The triply charged product ion representing neutral loss of dPLP was also present, but it was also quite small (not annotated). Figure 6.12A and 6.12B are the corresponding MS/MS spectra of the +2 precursors of the same dPLP peptides respectively. In both spectra, the dominant peaks are 49 Da lower in mass than the precursors. This indicates that the loss of H3PO4 is the most common CID product for both peptides. Both spectra exhibit peaks that represent neutral losses of dPLP (-115.5 Da) from the doubly charged precursors too, although they are small. They are annotated in the two spectra. The MS/MS spectra from the +2 precursors indicate the CID fragmentation of the doubly charged ions behave much more like the dPLP-modified Fibrinogen (F PLP) than the corresponding +3 precursors. 164 § 691.2. [11111, - 11,0043+ 88888388 :> A1 Ll Jr 111 llLl 11 l 1-1.114L171 1 1 L § 7423,0014, - H,POJ3+ Relative Intensity 3 888388 on 88 .n O l 200 400 600 800 1000 1200 1400 Figure 6.11. MS/MS spectra of +3 precursors (A) 24-39 and (B) 23-39 of dPLP-peptides from the tryptic digestion of dPLP-modified TIP39. 165 100 1036.3, [MH2 — H,Po.]2+ ‘970.1, [MI-12 - dPLPP‘ A 5’ E g 400 600 800 1000 1200 1400 1600 1600 2000 '5 a 2 10° 1114.6, [MHz — H3POJ2* ‘1048.1, [MHZ — dPLPF“ B t vvvvvvvvvvv l_l.ll'lll_lll_lvl"j'VVVt"""V“ 400 600 800 1000 2200 1400 1600 1600 2000 Figure 6.12 (A) MS/MS spectrum of +2 precursor of dPLP peptide 24-39 (m/z 1085.5). (B) MS/MS spectrum of +2 precursor of dPLP peptide 23-39 (m/z 1163.0). 166 6.7 Conclusions TIP39 was developed as a model to assess the possibilities of using mass spectrometry to determine sites of dPLP modification in proteins. This represented a simple case in which a small protein (~4.5 kDa) was modified by dPLP and the site of the modification was determined. MALDI TOF was used with mass mapping to determine where the dPLP modification was located. LC-MS/MS and neutral loss analysis was used to identify dPLP peptides from the digestion of TIP39PLP. The results indicate that both approaches may be successful in determining the site of dPLP modification in a real protein. 167 6.8 Chapter 6 References l. Aebersold, R., Goodlett, D.R. Mass spectrometry in proteomics. Chem. Rev. 101: 269-295 (2001). Buffoni, F ., Cambi, S. A method for the isolation and identification of pyridoxal phosphate in proteins. Anal. Biochem. 187: 44-50 (1990). Sugiyama, Y., Mukohata, Y. Modification of one lysine by pyridoxal phosphate completely inactivates chloroplast coupling factor 1 ATPase. FEBS Lett. 98: 276 (1979) 168 CHAPTER 7 IDENTIFICATION OF DEOXY—PYRIDOXAL-L-PHOSPHATE MODIFICATION SITES IN PROTEINS WITH TANDEM MASS SPECTROMETRY AND NEUTRAL LOSS ANALYSIS 7.1 Introduction: Initial Strategy for Identifying Deoxy-Pyridoxal-l-Phosphate Binding Sites in Proteins Chapter 5 of this report described the characteristic fragmentation pattern for deoxy-pyridoxal-l-phosphate (dPLP) modified peptides in CID experiments. It was shown that MS/MS spectra of these peptides were dominated by two main peaks. One represented the neutral loss of phosphoric acid, H3PO4, and the other represented the neutral loss of dPLP. Chapter 6 then described a simple experiment in which a small, dPLP-modified protein (TIP39PLP) was digested with trypsin in order to assess the potential of identifying dPLP binding sites by LC-MS/MS and searching for the characteristic neutral loss pattern exhibited by dPLP-peptides. It was also determined that detection of dPLP-peptides may be feasible with MALDI TOF mass spectrometry and mass mapping. The next step in the development of this protocol was to apply both LC-MS/MS and mass mapping with MALDI to a larger, naturally occurring protein that binds PLP. Based on the results described in chapters 5 and 6, an approach was outlined as summarized in Figure 7.1. First, the PLP—modification would have to be stabilized against hydrolysis. Therefore, the modification was reduced with sodium borohydride. Then the protein would have to be digested with trypsin or Glu-C. The digestion 169 Protein. 40 - 50 kDa l N 8311‘ Reduction 1 D139“ (WPSln, Glu-C) 1 Separate (LC) MALDI — search ESI (MS ) __ search for expected for eJ‘lleched mass (mass ma lug) fragmentation pp pattern Figure 7.1. The initial strategy employed to locate dPLP modifications in PLP-binding proteins. 170 products (peptides) would be fractionated with liquid chromatography and either analyzed with MALDI TOF and/or ESI and tandem mass spectrometry. Two proteins were used in the development of this protocol: Alanine Racemase (AlRace) and Aspartate Aminotransferase (AAT). Both are known to bind PLP and the sites of the modification have already been determined in both cases. Since the answer is already known, we would be able to evaluate our own method by determining whether our answer agrees with earlier experiments. Since we were only interested in determining the dPLP binding site in both proteins, no background information on them was necessary outside of the amino acid sequences. Figure 7.2 displays the amino acid sequence of AlRace. Lysine 39 (K-39) is the known binding site. It is the underlined residue in Figure 7.2. Figure 7.3 provides a similar look at the amino acid sequence of AAT. Lysine 258 (K—258) is the known PLP binding site in AAT (underlined in Figure 7.3). 7.2 Procedure for Sodium Borohydride Reduction of Proteins Sodium borohydride, NaBH4, reduction was performed with a modified protocol from Sugiyama et al. (1). About 0.3mg of protein (AAT or AlRace) was dissolved in about 200uL of lOOmM HEPES (pH 7.8). A UV-visible spectrum (scanned 280 to 500nm) was obtained against a blank containing lOOmM HEPES. Bound PLP was reduced with the addition of 5 1.1L 1M NaBH4. The mixture was left undisturbed for 45 minutes at room temperature in the dark. The protein solution was dialyzed against 2 x 1L of 25mM ammonium bicarbonate (pH 7.8). A second UV-visible spectrum was obtained against a blank containing 25mM ammonium bicarbonate. 171 MN DFHRDTWA EVDLDAIYDN VENLRRLLPD DTHIMAVVIgA NAYGHGDVQV ARTALEAGAS RLAVAFLDEA LALREKGIEA PILVLGASRP ADAALAAQQR IALTVFRSDW LEEASALYSG PFPIHFHlKM DTGMGRLGVK DEEETKRIVA LIERHPHFVL EGLYTHFATA DEVNTDYFSY QYTRFLHMLE WLPSRPPLVH CAN SAASLRF PDRTFNMVRF GIAMYGLAPS PGIKPLLPYP LKEAFSLHSR LVHVKKLQPG EKVSYGATYT AQTEEWIGT I PIGYAD GWLR RLQHFHVLVD GQKAPIVGRI CMDQCMIRLP GPLPVGTKVT LIGRQ GDEVI SIDDVARHLE TINYEVPCT I SYRVPRIFFR HKRIMEVRNA IGRGESSA Figure 7.2. The amino acid sequence in one letter designations for Alanine Racemase. The underlined lysine residue is the known PLP binding site (K-39). 172 APPSVFAEVP QAQPVLVFKL IADFREDPDP RKVNLGVGAY RTDDCQPWVL PVVRKVEQRI AN DSSLNHEY LPILGLAEFR TCASRLALGD DSPALQEKRV GGVQSLGGTG ALRIGAEFLA RWYNG'I'NNKD TPVYVSSPTW ENHNGVF'I'TA GFKDIRSYRY WDTEKRGLDL QGFLSDLENA PEFSIFVLHA CAHNPT GT DP TPEQWKQIAS VMKRRFLFPF FDSAYQGFAS GNLEKDAWAI RYFVSEGFEL F CAQSF SIiN'F GLYNERVGNL TVVAKEPDSI LRVLSQMEKI VRVTWSNPPA QGARIVARTL SDPELFHEWT GNVKTMADRI LSNIRSELRAR LEALKTP GT W NHI'I'DQIGMF SFTGLNPKQV EYLINEKHIY LLPSGRINMC GL'ITKNLDYV ATSIHEAVTK IQ Figure 7.3. The amino acid sequence in one letter designations for Aspartate Aminotransferase. The underlined lysine residue is the known PLP binding site (K-258). 173 7.3 Procedure for the Reduction, Alkylation, and Tryptic Digestion of Proteins The reduced protein was thermally denatured at 65°C for 15 minutes after the addition of SuL of 200mM dithiothreitol (DTT) to reduce disulfide bonds. Cysteine thiols were modified with the addition of 20pL of 200mM iodoacetamide. The reaction was placed in the dark for one hour at room temperature. 201.1L of 200mM DTT was added and the solution was left at room temperature for one hour. Trypsin or Glu-C was added at a 1:50 w/w ratio relative to protein. The mixture was incubated at 37°C for about 18 hours. At the end of incubation, 1 pL of concentrated acetic acid was added to stop the digestion in order to minimize autoprotolysis of trypsin. 7.4 Procedures for HPLC and Mass Spectrometric Analysis of Protein Digests Half of the digested sample was dedicated to HPLC separation and MALDI TOF analysis for mass mapping. The other half was dedicated for LC-MS/MS experiments. For MALDI TOF analysis, the digests were injected onto an HPLC column and fractions were collected at one minute intervals. A Reliasil C18 reversed phase column with the dimensions 2.0mm x 150mm was used. The column contained 300A particles with a pore size of 5 pm. Two mobile phases were used for gradient elution. Mobile phase A consisted of 95% water, 5% acetonitrile, and 0.1% trifluoroacetic acid (TFA). Mobile phase B consisted of 5% water, 95% acetonitrile, and 0.1% TF A. The mobile phase gradient began at 5% B and increased to 65% B at 30 minutes. A steeper gradient increased mobile phase B to 95% at 32 minutes where it remained for 3 minutes. The composition of B then decreased to 5% (at 38 minutes into the run) where it remained though the remainder of the run. Total time of the run was 45 minutes. The flow rate 174 throughout was 65 uL/minute. Each fraction off the HPLC column was analyzed by MALDI TOF in an attempt to identify the dPLP-peptide by mass mapping. DHB was used as the matrix. DECA ion trap LC-MSn experiments were carried out with ThermoFinnigan LCQ mass spectrometer. About ZOuL of sample was injected into the HPLC. The same column (described above) was used for the LC-MSn experiments as the one used for fraction collection and MALDI TOF analysis. The mobile phase and gradient were changed. Mobile phase A was composed of 5% acetonitrile, 95% water, and 0.1% formic acid while mobile phase B was composed of 95% acetonitrile, 5% water, and 0.1% formic acid. The mobile phase composition began as 5% B at the start. It was maintained that way for the first two minutes. It was then increased to 80% B by the 40 minute mark. It was maintained there for five minutes and then reduced to 5% B again over another five minutes. It was maintained at 5% B for the duration of the run. The total time of the run was 60 minutes. The flow rate was maintained at 75 uL/minute throughout. Peptides eluting from the HPLC were immediately analyzed by the LCQDECA. Peptides were converted to gas phase ions by electrospray and transferred to an ion trap mass analyzer. Ion m/z values were determined over time as peptides eluted from the HPLC. MS/MS experiments were carried out according to data-dependent settings. A low-limit abundance threshold was set. Any peak more intense than the threshold could be isolated and induced to fragmentation by CID. The most intense ion was selected first and induced to fragmentation. It was set to perform MS/MS on the same m/z value three 175 times in a row. After that, the instrument isolated the second most intense ion for MS/MS experimentation and so on. 7.5 Results of Proteolysis, Fractionation, and Mass Spectrometric Analysis of Protein Digests The UV-visible spectra of AlRace and AAT are displayed in Figures 7.4 and 7.5 respectively. In Figure 7.4, the spectrum of AlRace (at pH 8) is characterized by an absorption band with a maximum at about 420nm (2). The band was bleached by the addition of NaBH4 resulting in the appearance of a band at about 330nm. This band is characteristic of a reduced aldimine between dPLP and the side chain, a-amine of a lysine residue (3). Figure 7.5 displays the spectra of AAT at pH 8. A large band with a maximum absorbance at about 360nm and a small band of maximum absorbance at about 430nm are present. These bands are consistent to AAT at pH 8 (4). After NaBH4 treatment, both bands disappear and are replaced by a band with a maximum at 330nm. The band at 330nm for both proteins signifies a stable dPLP modification somewhere within the protein. For tryptic and Glu-C digestions of AlRace, the average, theoretical, protonated masses of the predicted dPLP-peptides would be 3023.4 and 1916.] Da respectively. For AAT, the corresponding masses would be 3272.5 and 2157.3 Da respectively. None of the predicted masses of dPLP peptides for either protein were detected using MALDI TOF of collected HPLC fractions. Likewise, none of the masses were observed using LC-MS nor was the distinct fragmentation pattern of dPLP-peptides observed in any of 176 0.4 I I _ Alanine Racemase o 0.3 E Reduced Alanine Racemase .D ‘3‘ o 2 .9 <1: 0.1 - 0 I l 1 7 . 300 350 400 450 500 Wavelength, nm Figure 7.4. UV-visible spectra of Alanine Racemase before and after NaBH4 reduction. 177 0.1 _ Aspartate Aminotrans ferase Reduced Aspartate Aminotransf erase (L08 0.06 — 0.04 — 0.02 . 0 1 1 1 305 355 405 455 Wavelength, nm Absorbance “\ Figure 7.5. UV-visible spectra of Aspartate Aminotransferasese before and after NaBH4 reduction. 178 the MS/MS spectra. However, both approaches were capable of identifying several peptides via mass mapping and/or peptide sequencing. A protein search was conducted with the Mascot search engine. It compares the masses form MALDI spectra to predicted tryptic or Glu—C product masses of thousands of proteins to determine a match. This is the automated approach to mass mapping. If the masses in a MALDI TOF spectrum match the theoretical masses of a protein in the database, then a match has been found. The database was able to successfully identify both AlRace and AAT. Both proteins were also identified by comparing the sequences of several peptides (determined from MS/MS spectra) to sequences within several thousand proteins in databases. This was performed by the Sequest search engine. 7.6 Discussion The UV-visible spectra displayed in Figures 7.4 and 7.5 prove that a dPLP modification is present in both proteins. Since both approaches (mass mapping and tandem mass spectrometry) can determine the mass and/or sequence of several peptides from the digestion of both proteins, it was possible that analysis of dPLP-modified peptides with mass spectrometry may be difficult because of the low response of modified peptides. Phosphopeptides have been shown to be difficult to detect with mass spectrometry (5). Since PLP also has a phosphate group, it wouldn’t be unreasonable to assume that dPLP-peptides behave in a similar manner. This becomes more problematic when the peptide mixture in the sample is more complex, as in large protein digests. Peptides compete for charge in both MALDI and ESI. Unmodified peptides ionize much easier than phosphopeptides, and presumably, dPLP-peptides. Therefore, dPLP-peptides 179 may not be ionized very efficiently in the presence of unmodified peptides and this inhibits its detection. Another approach had to be considered which would enhance the detection of dPLP-peptides or somehow simplify the sample (ie. Produce fewer peptides to compete with). 180 7.7 The New Strategy UV-visible spectroscopy demonstrates the stable modification of both AlRace and AAT by dPLP (Figures 7.4 and 7.5). However, mass spectrometric approaches (mass mapping and tandem mass spectrometry) were unsuccessful in detecting dPLP-peptides even after fractionating the digests. Since both approaches were successful in the TIP39PLP experiments described in Chapter 6, it was believed to be plausible for larger proteins too. However, large proteins digested with trypsin or Glu-C typically yield complex mixtures of dozens of peptides. As discussed above, the mixtures may be too complex even after fractionation to detect dPLP-peptides. The tryptic digestion of TIP39 produces only a few peptides, so the competition for charge in MALDI and ESI is less of a concern. Therefore, if the protein could be broken down into smaller pieces (comparable in size to TIP39), and then digested with trypsin or Glu-C, it may lead to the successfill identification of a dPLP-peptide. 7.7.1 Cyanogen Bromide Treatment Cyanogen bromide (CNBr) cleaves amide bonds of proteins on the C-terminal side of methionine (M) residues (6). The reaction typically converts M to homoserine or homoserine lactone, Figure 7.6. Methionine residues are one of the least common amino acids in proteins. AlRace has 10 M residues out of 388 and AAT has 6 M residues out of 412 total residues. Treatment of proteins with CNBr typically yields fewer peptides with much larger mass than trypsin and Glu-C digestions would yield. The few peptides that are produced are typically too large to get sequenced by mass spectrometry. 181 R-—-—NH C—NH-FC —C—R" + HCENL—Br 9' Cyanogen Bromide H3 H20 \_Y__J Methionine Residue 5 CH SCN + Br— 3 f' H \ 0 H o H o R__NH_C // “2° I II I 11 \ —’ R—NH—C—C —OH + HzN—C—C—R“ Hz)C /0 FIB' K 2 Homoserlne Lactone Homoserine Figure 7.6. An illustrates of the reaction between a methionine residue in a protein and cyanogen bromide generating a product mixture of homoserine and homoserine lactone. 182 Tables 7.1 and 7.2 list the theoretical peptides that would be expected from treatment of AlRace and AAT with CNBr. Since the complete protein sequences are available, and we know the site of dPLP modification, we could predict the mass of the peptide containing the modification. We could then use HPLC to separate all of the CNBr products and isolate the suspected dPLP-modified peptide from the rest of the protein. Ultimately, the search for the modified residue would be reduced to a large peptide rather than one of many small peptides from a large protein. This would result in a peptide mass (modified by dPLP) similar in mass to the dPLP-modified TIP39 peptide discussed in chapter 6 which proved successful. 7.7.2 Procedure for Cyanogen Bromide Treatment Proteins were treated with CNBr via a protocol modified from Morrison et al. (7). About 1mg of each protein was used for this experiment. A UV-visible spectrum was obtained against a 50mM HEPES blank. Before CNBr treatment, the proteins were reduced with NaBH., to ensure the stability of the PLP modification. The proteins were then dialyzed against 2 x 1L of water. A post-NaBH4 reduction/dialysis UV-visible spectrum was obtained. The samples were dried down to approximately SuL and reconstituted in lOOuL of CNBr solution (2mg/100uL) in a capped 1.5mL micro- centrifuge tube. Before capping, the sample was purged with ultra pure nitrogen gas. The CNBr solution was prepared by dissolving 2mg of CNBr in lOOuL of 70% formic acid. The sample was incubated at room temperature in the dark for two hours. After the two hour incubation period, the sample was dried down completely under nitrogen, and reconstituted in lOOuL pure water. 183 Table 7.1. The theoretical products of cyanogen bromide treatment of Alanine Racemase. Predicted Peptides Ajg. MH+ Mass (Da) 2-35 4066.4 36-130 10035.6 36-130* 10266.7 135-188 6398.2 189-217 3279.8 225-312 9764.5 317-375 6741.0 376-388 1346.5 Table 7.2. The theoretical products of cyanogen bromide treatment of Aspartate Aminotransferase. Predicted Peptides Ayg. MHJr Mass (Da) 1-212 23484.5 213-287 8608.8 213-287* 8840.0 288-326 4839.0 334-359 2947.3 360-389 3476.0 390-412 2506.9 * dPLP modified peptide 184 7.7.3 Procedure for the HPLC Separation of Cyanogen Bromide Products and Mass Spectrometric Analysis In order to separate and analyze the CNBr products, 20uL of the CNBr digest was injected into an HPLC instrument. Since large peptides were expected, the separation was performed with a reversed-phase C4 column from Vydac. The column dimensions were 1.0mm inner diameter x 150mm in length. The column was composed of Sum particles with approximately 300A pore size. Two mobile phases were used for gradient elution. Mobile phase A consisted of 95% water, 5% acetonitrile, and 0.1% trifluoroacetic acid (TFA). Mobile phase B consisted of 5% water, 95% acetonitrile, and 0.1% TFA. The mobile phase gradient began at 5% B and increased to 65% B at 30 minutes. A steeper gradient increased mobile phase B to 95% at 32 minutes where it remained for 3 minutes. The composition of B then decreased to 5% (at 38 minutes into the run) where it remained though the remainder of the run. Total time of the run was 45 minutes. The flow rate throughout was 65 uL/minute. Peptides were detected with a UV- visible detector set at 214nm. Fractions corresponding to chromatographic peaks were collected for analysis by MALDI TOF. Sinapinic acid was used as the matrix. Matrix preparation involved dissolving a saturating amount of sinapinic acid in 50:50 acetonitrile and water. 7.7.4 Results Figure 7.7 displays the HPLC chromatogram of the separation of the CNBr products of AlRace. The corresponding chromatogram for AAT is displayed in Figure 185 290 - 240 - 190~ 140 - 90- Absorbance (mAU) at 214nm 40- 27.5 min. ._ K —r—'" l I M ‘1 15 20 25 30 35 Time (minutes) Figure 7.7. HPLC chromatogram for the separation of CNBr products of Alanine Racemase. 186 7.8. Fractions corresponding to all peaks in the chromatograms were collected for MADLI TOF analysis. Table 7.1 displays the theoretical peptide products, with corresponding masses, expected from CNBr treatment of AlRace. Table 7.2 displays the corresponding information for AAT. As shown in Table 7.1, the CNBr product of AlRace that should contain the dPLP-modified lysine is the peptide 36-130. This modified peptide has an average, protonated, theoretical mass of 10266.7 Da. Average masses will be considered here since MALDI TOF is incapable of monoisotopic resolution in this mass range. Since both AlRace and AAT were purchased from Sigma and the manufacturer claims that PLP was not present in quantities high enough to saturate either enzyme, we were prepared to observe the unmodified peptide 36-130 as well. This unmodified peptide has an average, protonated, theoretical mass of 10035.6 Da (Table 1). According to Table 7.2, the CNBr product of AAT that should contain the dPLP-modified lysine (K-258) is the peptide 213-287. The modified peptide 213-287 has an average, protonated, theoretical mass of 8840.0 Da and the unmodified peptide has a corresponding mass of 8608.8 Da. Figure 7.9 displays a MALDI TOF mass spectrum of the collected fi'action corresponding to the chromatographic peak at 27.5 minutes in Figure 7.7. Figure 7.7 is the HPLC chromatogram of the separation of the CNBr products of AlRace. The two large peaks register m/z values of 10037.8 and 10266.8 Da which are very close to the predicted masses of the unmodified and dPLP-modified peptide (AlRace 36-130) respectively. These two masses differ by 229 Da which correlates closely to the 231 Da difference that would be expected between an unmodified and dPLP-modified peptide of the same amino acid sequence. 187 § E v 5m p 8 278ml : . n. a: 9 250 - O to .0 < 0 J n n 1 4 12 15 18 21 24 27 30 33 13 Retention Time (m in) Figure 7.8. HPLC chromatogram for the separation of CNBr products of Aspartate Aminotransferase. 188 45000 - 40000 ~ 10032.1 35°°° ‘ 10264.5 30000 - 25000 - 20000 - 15000 - 10000 - 5000 - 0 1 I 1 8000 9000 10000 11000 12000 13000 m/z Absolute Intensity Figure 7.9. MALDI TOF spectrum of the fraction corresponding to the chromatographic peak at 27.5 minutes in the HPLC chromatogram in Figure 7.7. 189 Figure 7.10 displays a MALDI TOF mass spectrum of the collected fraction corresponding to the broad chromatographic peak at 27.8 minutes in Figure 7.8. Figure 7.8 is the HPLC chromatogram of the separation of the CNBr products of AAT. The spectrum shows two annotated peaks at m/z values of 8665 and 8899 Da respectively. The peaks have been assigned (and annotated) as the unmodified peptide (AAT 213-287) of AAT and the corresponding dPLP modified peptide for 8865 and 8899 Da respectively. However, it should be pointed out that each of these m/z values in Figure 7.10 are about 58 Da higher than the theoretical mass of each peptide. Furthermore, the two assigned peaks in Figure 7.10 are broad compared to the peaks observed in Figure 7.9 for the AlRace peptides (36-130). Since the m/z ratios in Figure 7.9 were so close to the theoretical masses of the unmodified and dPLP-modified peptides 36-130 for AlRace, instrumental error (such as a poor calibration) was ruled out. The mass error and peak broadness in Figure 7.10 may be due to adducts, some unanticipated chemistry, or a combination of the two. This observation will be reintroduced and discussed in more detail later in this chapter. The mass spectrometry just described implies that the isolated HPLC fractions from CNBr treatment of both AlRace and AAT contain the dPLP-modified peptides, as well as the unmodified analogs, that were predicted according to the theoretical lists in Table 7.1 and 7.2. To ensure that this was the case, each fraction was dried down to a minimal volume (<5 uL), reconstituted in 25mM ammonium bicarbonate (pH 7.8), and analyzed with UV-visible spectroscopy. The instrument was scanned from 280 to 500nm against a 25mM ammonium bicarbonate background. The 25mM ammonium bicarbonate was use for the subsequent tryptic and/or Glu-C digestions. The 190 35000 - 30000 1 8665 25000 ~ 20000 . 15000 - 8899 10000 - Absolute Intensity 5000 . o T 1 f I l 1 7500 8000 8500 9000 9500 10000 10500 m/z Figure 7.10. MALDI TOF spectrum of the fraction corresponding to the chromatographic peak at 27.8 minutes in the HPLC chromatogram in Figure 7.8. 191 corresponding UV-visible spectra are displayed in Figures 7.11 and 7.12. Figure 7.11 is the spectrum for AlRace peptide 36-130. There is a band at approximately 325nm which would indicate the presence of a reduced aldimine bond between PLP and lysine (K-39). This provides strong assurance that the peptide carrying the dPLP modification (AlRace 36-130) has been successfully isolated from the rest of the AlRace peptides. Figure 7.12 offers similar assurance that the dPLP-modified peptide in AAT (213-287) has also been isolated from the rest of the protein. 7.8 Proteolysis of the Isolated dPLP-Modified Products from Cyanogen Bromide Treatment Alanine Racemase (AlRace) and Aspartate Aminotransferase (AAT) were treated with cyanogen bromide (CNBr) and the products were separated by HPLC. All HPLC fractions that corresponded to chromatographic peaks were collected for analysis by MALDI TOF MS. Mass mapping was used to assign m/z values to corresponding peptides within each protein. Masses representing dPLP-modified peptides were observed. In conjunction with mass spectrometry, UV-visible spectroscopy was used to ensure that these same peptides were dPLP modified. The UV-visible spectra supported this conclusion. The initial strategy described earlier (Figure 7.1) was to reduce the proteins with NaBH.,, digest with trypsin or Glu-C, fractionate with HPLC, then analyze the digestion products (peptides) with mass spectrometry. This work was conducted with proteins that had a mass around 45 kDa. Tryptic and Glu-C digestion resulted in a large number of 192 0.06 0.05 Absorbance P o w l 285 335 385 435 485 Wavelength, nm Figure 7.11. UV-visible spectrum of the HPLC fraction corresponding to the chromatographic peak at 27.5 minutes from Figure 7.7. 193 0.09 - 0. 06 — Absorbance 0.03 1 0 l l 1 I 285 335 385 435 485 Wavelength, nm Figure 7.12. UV-visible spectrum of the HPLC fraction corresponding to the chromatographic peak at 27.8 minutes from Figure 7.8. 194 peptides that obscured the presence of weakly responsive dPLP peptide products. By treating these proteins with CNBr and isolating the dPLP-modified, CNBr product, we were left with only about 20 to 30% of the initial protein mass to consider. Tryptic or Glu-C digestions of these peptides would yield only a few peptides making it much more manageable in our attempt to locate the dPLP binding site. The mass is now much more similar to TIP39PLP, described in Chapter 6. Once it had been determined which of the CNBr products were dPLP modified, the next step was to narrow it down to which residue was modified. The remaining amino acid sequence of AlRace (36-130) is listed in Figure 7.13. There are three lysine (K) residues remaining and all are potential PLP binding sites. Figure 7.13 also lists the theoretical peptides that would result from tryptic digestion of AlRace 36-130. Included is the theoretical sequence and corresponding mass of the peptide with the modified residue (K-39). Figure 7.14 displays the theoretical peptides resulting from the Glu-C digestion of AlRace 36-130. Likewise, the list contains the predicted sequence and mass of the peptide that is expected to be modified by dPLP. The corresponding theoretical digestion products of AAT 213-287 by both trypsin and Glu-C are listed in Figures 7.15 and 7.16 respectively. 7.8.1 Trypsin and Glu-C Digestion Procedures The CNBr products suspected of dPLP modification were dried down and reconstituted in 25mM ammonium bicarbonate. Ammonium bicarbonate provides a suitable pH environment buffered at about pH 8. After analysis by UV-visible spectroscopy, the peptides were ready for digestion by trypsin and Glu-C. An estimation 195 H— Alanine Racemase Sequence 36-130: AVVEANAYGH GDVQVARTAL EAGASKLAVA FLDEALALRE KGIEAPILVL GASRPADAAL AAQQRIALTV F RSDWLEEAS ALYSGPFPIH FHLKh Theoretical tryptic digestion products: Predicted Peptides Avg. Mass (2a) 116-52* 1987.1 53-61 876.0 6274 1402.7 77- 100 2390.0 101-107 820.0 108-129 2545.9 * = suspected dPLP product h = homoserine or homoserine lactone residue _K_ = suspected dPLP modified residue italicized residues represent cleavage sites Figure 7.13. The sequence (36-130) of the isolated CNBr product of Alanine Racemase. The predicted products of trypsin digestion are also listed. 196 Alanine Racemase Sequence 36-130: AvngNAYGH GDVQVARTAL EAGASRLAVA FLDEALALRE KGIEAPILVL GASRPADAAL AAQQRIALTV FRSDWLEEAS ALYSGPFPIH FHLKh Theoretical Glu-C digestion products: Predicted Peptides Avg, Mass (2a) 3647* 1433.5 48-56 987.2 57-68 1191.4 6974 672.8 68-74 801.9 75-91 1708.0 79-91 1280.5 92-108 1832.1 * = suspected dPLP product h = homoserine or homoserine lactone residue 5 = suspected dPLP modified residue italicized residues represent cleavage sites Figure 7.14. The sequence (36-130) of the isolated CNBr product of Alanine Racemase. The predicted products of Glu-C digestion are also listed. 197 Aspartate Aminotransferase Sequence 213-287: KRRFLFPF F D SAYQGFASGNL EKDAWAIRYF VSEGFELFCA QSFS_KNFGLY NEKVGNLTVV AKEPDSILRV LSQh Theoretical tryptic digestion products: Predicted Pgtides Av . Mass 3 216-235 2286.6 236-241 731.8 242-266* 3215.5 267-275 901.1 276-282 829.9 * = suspected dPLP product h = homoserine or homoserine lactone residue 5 = suspected dPLP modified residue italicized residues represent cleavage sites Figure 7.15. The sequence (213-287) of the isolated CNBr product of Aspartate Aminotransferase. The predicted products of tryptic digestion are also listed. 198 Aspartate Aminotransferase Sequence 213-287: KRRFLFPFFD SAYQGFASGNL EKDAWAIRYF VSEGFELFCA QSFSIgNFGLY NERVGNLTVV AKEPDSILRV LSQh Theoretical Glu-C digestion products: Predicted Peptides Avg. Mass (2a) 213-222 1373.7 223-234 1244.3 237-246 1242.4 250-265* 2100.3 266-276 1186.4 * = suspected dPLP product h = homoserine or homoserine lactone residue _13 = suspected dPLP modified residue italicized residues represent cleavage sites Figure 7.16. The sequence (213-287) of the isolated CNBr product of Aspartate Aminotransferase. The predicted products of Glu-C digestion are also listed. 199 was used to determine how much protease to add in order to have a 1:50 protease to peptide ratio. Since about 1mg of protein was used at the outset, and approximately 20% of the protein was left after CNBr treatment and fractionation, it was assumed that about 200ug of protein was left. Therefore, about 411g of trypsin and Glu-C was used. After addition of the protease, the mixture was incubated at 37°C for 18 hours. luL of concentrated acetic acid was added at the end of the incubation period. 7.8.2 Procedures for the HPLC and Mass Spectrometric Analysis of Tryptic and Glu-C Digestion Products of Alanine Racemase 36-130 and Aspartate Aminotransferase 213-287 For analysis of tryptic and Glu-C digestion products of AlRace 36-130 and AAT 213-287, MALDI TOF was employed before and after HPLC fiactionation. For HPLC fractionation the following parameters were used. A Reliasil C18 reversed phase column with the dimensions 2.0mm x 150mm was used. The column contained 300A particles with a pore size of 5 pm. Two mobile phases were used for gradient elution. Mobile phase A consisted of 95% water, 5% acetonitrile, and 0.1% trifluoroacetic acid (TFA). Mobile phase B consisted of 5% water, 95% acetonitrile, and 0.1% TF A. The mobile phase gradient began at 5% B and increased to 65% B at 30 minutes. A steeper gradient increased mobile phase B to 95% at 32 minutes where it remained for 3 minutes. The composition of B then decreased to 5% (at 38 minutes into the run) where it remained throughout the duration of the run. Total time of the run was 45 minutes. The flow rate throughout was 65 uL/minute. Each fraction off the HPLC column was collected at one 200 minute intervals and analyzed by MALDI TOF in an attempt to identify the dPLP-peptide by mass mapping. DHB was used as the matrix. DECA ion trap LC-MSn experiments were carried out with ThermoFinnigan LCQ mass spectrometer. About 20uL of sample were injected into the HPLC. The same column (described above) was used for the LC-MSn experiments as the one used for fraction collection and MALDI TOF analysis. The mobile phase and gradient were changed. Mobile phase A was composed of 5% acetonitrile, 95% water, and 0.1% formic acid while mobile phase B was composed of 95% acetonitrile, 5% water, and 0.1% formic acid. The mobile phase composition began as 5% B at the start. It was maintained that way for the first two minutes, then it was then increased to 80% B by the 40 minute mark. It was maintained there for five minutes and then reduced to 5% B again over another five minutes. The gradient was maintained at 5% B for the duration of the run. The total time of the run was 60 minutes. The flow rate was maintained at 75 uL/minute throughout. Peptides eluting from the HPLC were immediately analyzed by the LCQDECA. Peptides were converted to gas phase ions by electrospray and transferred to an ion trap mass analyzer. Ion m/z values were determined over time as peptides eluted from the HPLC. MS/MS experiments were carried out according to data-dependent settings. A low-limit abundance threshold was set. Any peak more intense than the threshold could be isolated and induced to fragmentation by CID. The most intense peak was selected first and induced to fragmentation. It was set to perform MS/MS on the same m/z value three times in a row. After that, the instrument isolated the second most intense peak for MS/MS experimentation and so on. 201 7.9 Results: Mass Spectrometric Analysis of Tryptic and Glu-C Digestion Products of Alanine Racemase 36-130 and Aspartate Aminotransferase 213-287 The first analysis was performed with MALDI TOF in an attempt to detect the predicted dPLP-peptides by mass mapping. The digest was analyzed directly prior to fractionation by HPLC. While several of the peptides in Figures 7. 13-16 were observed, none of the predicted dPLP-peptides were detected. HPLC was employed in each case to fractionate the digests. Fractions were collected at one minute intervals, dried down to a minimal volume (<10uL) and analyzed by MALDI TOF. Again, while many of the peptides predicted in Figures 7. 13-16 were observed, none of the predicted dPLP- peptides were observed. Although this approach was successful with TIP39PLP (Chapter 6), dPLP peptides may be difficult to detect with so many peptides in the same mixture, which is analogous to the behavior of phosphopeptides with MALDI analysis. This marked the end of our efforts to identify dPLP-peptides with MALDI TOF and mass mapping. The rest of the discussion will be directed toward the LC-MS/MS approach. 7.9.1 LC-MS/MS Analysis of Tryptic Digestion of Alanine Racemase 36-130 Figure 7.17A displays the base-peak ion chromatogram of the tryptic digestion of AlRace 36-130. To determine if any of the peaks in the chromatogram correspond to dPLP peptides, a search for the characteristic neutral loss pattern among the MS/MS spectra was carried out. Figure 7.178 and 7.17C are neutral loss chromatograms for 32.7 Da (loss of H3PO4) and 77 Da (loss of dPLP) respectively, from a triply charged precursor. The neutral loss chromatograms imply that a dPLP-peptide elutes over a broad time frame from about 20 to 27 minutes. It doesn’t appear obvious from the base peak 202 Relative Abundance 30 81 Base Peak Ion Chromatogram saga Neutral Loss Chromatogram (32.7 Da) [MH3 — H3P0413+ .111... . Neutral Loss Chromatogram (77 Da) [MH3 — dPLP]3+ e.— =— :a—_ O_ 5 1 0 1 5 20 25 30 35 40 45 50 55 60 Time (min) Figure 7.17. (A) The base peak ion chromatogram illustrating the separation/detection of the tryptic products of AlRace 36-130. (B) The neutral loss chromatogram indicating the retention times in which the neutral loss of 32.7 Da was detected in a MS/MS spectrum. (C) The neutral loss chromatogram indicating the retention times in which the neutral loss of 77 Da was detected in an MS/MS spectrum. 203 ion chromatogram (Figure 7.17A) that anything of substantial abundance elutes at that that retention time relative to the distinct peaks observed at 17.8, 19.7, 28.2, 30.8, 35.8, and 39.3 minutes. Yet the neutral loss chromatograms indicate that the dPLP peptide elutes at that time. Figure 7.18 displays a full scan spectrum within that time frame, 23.4 minutes. The average, protonated, theoretical mass of the dPLP peptide (36-52) from AlRace 36-130 is 1987.1 Da (Figure 7.13). The three large peaks in the spectrum (Figure 7.18) represent three charge states that correspond to the theoretical mass. These are annotated in the spectrum. While the detection of an ion with an m/z value that matches the theoretical mass of the dPLP peptide of AlRace is a positive sign that the dPLP peptide has been identified, tandem mass spectrometry could provide conclusive evidence. Each of these ions in the +2, +3, and +4 charge states (Figure 7.18) were subjected to CID and the corresponding MS/MS spectra are displayed in Figure 7.19. Earlier experiments (described in chapter 5) provided information on the fragmentation patterns of dPLP peptides. It was learned that these peptides fragment in such a mechanism that leads to two dominant products characterized as the neutral losses of H3PO4 and dPLP. Very few other fragmentation products are typically observed. The spectra in Figure 7.19 are consistent with this pattern. In Figure 7.19A, the MS/MS spectrum of m/z 994.5 displays both common neutral losses. The peak at 945.4 is 49.1 Da smaller than the precursor mass which corresponds to the neutral loss of H3PO4 from a +2 precursor. The peak at m/z 878.9 is 115.6 Da smaller than the precursor and corresponds to the neutral loss of dPLP from the precursor. The MS/MS spectrum in Figure 7.19B displays two dominant peaks representing the neutral loss of H3PO4 (m/z 630.4, loss of 32.6 Da) and dPLP (m/z 204 100 497.5, [MHJ‘t ,5 663.0, [MH3]3* 80 / 75 70 88888 40 994.5, [Ml-1212* 35 30 Relative Abundance 20 15 10 200 400 600 000 1000 1200 1400 1600 1800 2000 Figure 7.18. A full-scan ESI spectrum at about 23.4 minutes within the LC-MS/MS run represented by the base peak ion chromatogram in Figure 7.17A. The three annotated peaks in the spectrum represent three charge states for the tryptic digestion product 36-52 of AlRace 36-130. 205 100 60 40 20 a 80 60 40 Relative Abundance I 11111 200 94 .4. [Ml-i, - 14,903+ MS/MS of m/z 994.5 * [Mm—mm 8783‘ 869.9 995.9 5712 .6 746.5 .\ 1 1/ 630.4, [mg- H,Po,]3+ 1202.7 . MS/MS of m/z 663.0 * IMH. — We 588.1' , , . 749.3 606.9 “a...“ n a . MS/MS of m/z 497.5 573.4 1 _A .LiLlJ. IIII'IIII‘ITIIIIIlll‘llllll‘llllllIIII‘I'IIIIIIIIIIIII 300 400 500 600 700 000 900 1000 1100 1200 1300 mlz Figure 7.19. (A) MS/MS spectrum of precursor ion 994.5 Da. (B) MS/MS spectrum of precursor ion 663.0 Da. (C) MS/MS spectrum of precursor ion 497.5 Da. 206 586.1, loss of 76.9 Da) respectively. Figure 7.19C displays an MS/MS spectrum of m/z 497.5 Da. The presence of two dominant peaks with few other peaks present implies that the precursor is a dPLP peptide. However, the peaks in Figure 7.19C have larger m/z values than the precursor and we were unable to correlate a structure for the products represented by those peaks. Nonetheless, the pattern in this spectrum suggests that the products are somehow structurally related to the neutral loss products that have been characterized for dPLP peptides in this study. 7.9.2 LC-MS/MS Analysis of Glu-C Digestion of Alanine Racemase 36-130 AlRace 36-130 was also subjected to Glu-C digestion. Figure 7.20A displays the base peak ion chromatogram of the LC-MS2 experiment of the Glu-C digestion of AlRace 36-130. The chromatogram displays several features representing peptides eluting over time and analyzed by the mass spectrometer. To determine whether any of the peaks correspond to dPLP modified peptides, all MS/MS spectra were examined for the characteristic neutral loss pattern of dPLP peptides. The two neutral loss chromatograms in Figures 7.20B and C correspond to neutral losses of 32.7 Da (loss of H3PO4) and 77 Da (loss of dPLP) respectively, from a tn'ply charged precursor. The neutral loss chromatograms both display a large response to these queries between 24.0 and 26.5 minutes. This appears to correspond to the chromatographic peak marked 24.4 minutes in Figure 7.20A. Interestingly, when we chose to search for both characteristic neutral losses from a +2 precursor, two main responses were observed. Figures 7213 and C display neutral loss chromatograms for loss of 49 Da (loss of H3PO4) and 115.5 Da (loss of dPLP) from a +2 precursor. Figure 7.21A is a repeat of Figure 7.20A. Like 207 1 29.91 80 Base Peak Ion Chromatogram 60 21-73 a .79 A 40 10.4 a... 20 45.19 , , - - .. IL 211; 0 1 8 60 Neutral Loss Chromatograrn (32. 7 Da) 5 60 [MH3— H3PO4]3+ g B 4o 8 -.: 20 _u_1 0 0 I! 1 8,, Neutral Loss Chromatogram (77 Da) so [MH3— dPLP]3+ C 40 20 ooltrtlélti'1lolI‘I1BIrIIZIOIA-I'”I'Lrjrrlllll'l'é'llr&r111&11'lélll'sb “I"Ime (min) Figure 7.20. (A) Base peak ion chromatogram illustrating the separation/detection of the Glu-C products of AlRace 36-130. (B) The neutral loss chromatogram indicating the retention times in which the neutral loss of 32.7 Da was detected in an MS/MS spectrum. (C) The neutral loss chromatogram indicating the retention times in which the neutral loss of 77 Da was detected in an MS/MS spectrum. 208 Relative Abundance 10 2991 Base Peak Ion Chromatogram -| Neutral Loss Chromatogram (49 Da) 6° 3 [MH2 — dPLP]2+ 2:) 1 ”111.111 1.. .111 Neutral Loss Chromatogram (115.5 Da) [MH2 - dPLP]2+ 0 5 10 15 20 25 30 as 40 45 50 55 60 Time (mln) Figure 7.21. (A) Base peak ion chromatogram illustrating the separation/detection of the Glu-C products of AlRace 36-130. (B) The neutral loss chromatogram indicating the retention times in which the neutral loss of 49 Da was detected in an MS/MS spectrum. (C) The neutral loss chromatogram indicating the retention times in which the neutral loss of 115.5 Da was detected in an MS/MS spectrum. 209 Figures 7.20B and C, Figures 7.21 B and C display a neutral loss response at about 24 minutes. However, there is also a distinct response between 18 and 20 minutes. There doesn’t even appear to be a noticeable peak during this time frame in the base peak ion chromatogram (Figure 7.2 l A). Figures 7.22A and B display full scan mass spectra from both elution times where neutral loss chromatograms imply the presence of dPLP peptide spectra (18-20 and 24-26 minutes). According to Figure 7.14, the product of Glu—C digestion of AlRace 36-130 that is predicted to be modified by dPLP, should be peptide 36-47 which has an average, protonated, theoretical mass of 1433.5 Da. The spectrum in Figure 7.22A displays three peaks that correspond to three different charge states for the dPLP peptide 36-47. The +1 charge state is annotated in Figure 7.22A at m/z 1432.2 Da, the +2 state at m/z 717.0 Da, and the +3 charge state is annotated at m/z 478.6 Da. Figures 7.23A and B display corresponding MS/MS spectra of the +2 charge state (m/z 717.0 Da) and the +3 charge state (m/z 478.6) respectively. Figure 7.23A displays two large peaks that correspond to the neutral loss of H3PO4 (m/z 668.0 Da, loss of 49 Da) and the neutral loss of dPLP (m/z 601.6 Da, loss of 115.4 Da). The two large peaks in the MS/MS spectrum of the +3 precursor, m/z 478.8 Da, (Figure 7.23B) appear to be related to the neutral loss pattern that is distinct to dPLP peptides. Both of the m/z values of the two largest peaks, 632.1 and 583.1 Da, are larger than the m/z value of the triply charged precursor, m/z 478.6 Da. Obviously, the two most abundant products do not correlate to neutral losses of 32.7 and 77 Da that has been consistent with triply charged dPLP peptide MS/MS spectra. This explains why there was a not a neutral loss response in the neutral loss chromatograms for 32.7 and 77 Da at about 18 to 20 minutes (Figures 7.20 B and C). It was assumed that 210 1 , , 717.0, [MH2]2* 80 631.1 70 3+ so 478.6, [MHa] A 50 40 20 3905 10 “-4 _ 12.9 1432.2, MH+ 11.1 \ 1200.9, [MHZF Relative Abundance 0 1151.9 801.0, [MH-J” 10855 038 200 400 600 900 1000 1 200 1 400 1 800 1 800 2000 Figure 7.22. (A) Full scan ESI spectrum at 18.9 minutes of the base peak ion chromatogram in Figures 7.20A and 21A (they are the same chromatogram). (B) Full scan ESI spectrum at 24.5 minutes of the base peak ion chromatogram in Figures 7.20A and 21A. 211 5° 601.6, [MHZ - «PLP? A 11.111111 l-AJ . 1569.0.[1121112 — H,Po,]2+ 100 632.1 Relative Abundance 8 200 400 800 20 5 1 ‘10 541.2 0 11 11: 1110-1111114] liuih 1.1 I 1 11‘ .1 All i .. j I l | I 1 | I I ' l 800 mlz A. MS/MS of m/z 717.0, [M112]2+ l 11 J._LL. L 11. B. MS/MS of m/z 478.6, [M113]3+ Figure 7.23. (A) MS/MS of precursor ion m/z 717.0. (B) MS/MS of precursor ion m/z 212 both peaks (Figure 7.23B) represent product ions that are somehow related to the loss of H3PO4 and dPLP respectively, but the exact identity of the products could not be determined, nor was it investigated any further. The predicted dPLP peptide 36-47 from Glu-C digestion of AlRace 36-130 was detected. The observation of the proper mass in three different charge states and the observation of an MS/MS fragmentation pattern characteristic of dPLP peptides provides decisive evidence that the correct dPLP peptide has been determined. However, the neutral loss chromatograms in Figures 7.20B, 7.20C, 7.21B, and 7.21C all imply two different dPLP peptides eluting off of the column. The peaks around 24.5 minutes need to be explained. A representative full scan spectrum at 24.5 minutes in Figure 7.22B displays a base peak at m/z 1200.9. Zoom scans (not shown) indicate that this peak represents a doubly charged ion. This corresponds to a peptide mass of about 2401 Da. There is a related peak at m/z 801.0 Da. Zoom scans (not shown) indicate this is a triply charged ion which correlates to a peptide mass of about 2401 Da as well. MS/MS spectra imply that both of these ions represent the same dPLP modified peptide. Figures 7.24A and B display the MS/MS spectra of m/z 1200.9 Da (+2 precursor) and m/z 801.0 Da (+3 precursor) respectively. The main products in both spectra correspond to both neutral loss products characteristic of dPLP peptides as illustrated in Figure 7.24. If there are two different dPLP-peptides detected, then does this mean there are two PLP binding sites? Figure 7.25 helps explain the observation. Missed cleavages are often observed from proteolytic digestions of proteins. If Glu-C skips the cleavage site at aspartic acid 47 (D-47) and cleaves at the next site instead, glutamic acid 56, E-56, then the resulting peptide (36-56) would have an average, protonated, theoretical mass of 213 100 1152.0, {1111112 - H,Po,}2+ 9o 60 70 6° A 50 3 4o ‘m/z1085.6,[M-lz-dPLP]Z* : 30 a 1:1 20 10763,, C a 10 H a 1 1 1 ( o ' l V l i l' "1""I“"I'V'Yl'"'1""T""I""l""I""I""l""l""I""l '"'I""T‘V'VT‘VYVTV'V'I'"VT""I"'VI"‘r 5 100 7685. [ling-11,110,]3+ (I '3 90 I! so 70 60 B 50 40 3° 724.1,[MH,-dpLP]3+ 20 \ 10 o 1111111|11111111|111111111111 111 1 1 '1 1 1111' 11' 11 11 1 111 '1 1111111 11111111|11 1111111111111 400 600 900 1000 1200 1400 ml: Figure 7.24. (A) MS/MS spectrum of precursor ion m/z 1200.9. (B) MS/MS spectrum of precursor ion m/z 801.0. 214 dPLP peptide 36—47 MH+(M) = 1433.5 Da r W AVV_I$_ANAYGHGDXQVARTALEA \_ J V dPLP peptide 36—56 MH+ = 2401.6 Da (Wm) A = Glu-C cleavage sites _lg = suspected dPLP modified residue Figure 7.25. An illustration of the mixture of digestion products possible by cleavage at D-47 or by the missed cleavage at D-47. 215 2401.6 Da. Both peptides (36-47 and 36-56) contain just one lysine residue so the modification can be assigned to K-39 in both cases. 7.9.3 LC-MS/MS of the Tryptic Digest of Aspartate Aminotransferase 213-287 The same experiments, analysis, and interpretation were performed with AAT. Since the material was covered so extensively for AlRace, less time will be spent on the data for AAT since the conclusions are the same. Figures 7.26A, B, and C display the base-peak ion chromatogram (A), the neutral loss chromatogram for 49 Da (B), and the neutral loss chromatogram for 115.5 Da (C) respectively. These neutral losses correspond to the loss of H3PO4 (Figure 7.26B) and dPLP (Figure 7.26C) from a +2 precursor respectively. Both neutral loss chromatograms contain a feature at about 35 minutes. Since two neutral loss products are expected, this most likely corresponds to a dPLP peptide. The large peak at about 25.4 minutes representing a neutral loss of 115.5 Da (Figure 7.26C) has been attributed to a different neutral loss artifact of the same mass. Evidence to support this comes from the fact that there was not a product detected that represented a neutral loss of H3PO4 (loss of 115.5 Da) in Figure 7.26B at the same time. The H3PO4 neutral loss product is normally more abundant than the dPLP neutral loss product so it’s unlikely to observe just the loss of dPLP. MS/MS data at that retention time (25.4 minutes) indicate that the analyte is not a dPLP-peptide (not shown). Figures 7.27 display a full-scan spectrum at 35.1 minutes. From the tryptic digestion of AAT 213-287, the theoretical dPLP peptide 242-266 should have an average, protonated mass of 3215.5 Da. The peak at m/z 1089.0 Da represents a triply charged ion and the peak at m/z 1631.7 represents a doubly charged ion. These charge states 216 lfll78 80 Base Peak Ion Chromatogram 50 1‘. .34 4o 20 \ 8 o g A ‘ W311 3 .97“ A 1551 NeutIal Loss Chromatogram (49 Da) 5 8° [MH2 — dPLP]2+ 2 so B 0 .42., 40 2 12° 1. 10 so Neutral Loss Chromatogram (115.5 Da) 3° C [MH2 — dPLP]2+ 40 20 oI'llll“"l'llll'llllllll llrlll'ljllllllJ'Allll""l'll'lfirll 0 5 10 15 20 25 30 35 40 45 50 55 Time (min) Figure 7.26. (A) The base peak ion chromatogram of the LC-MS/MS analysis of the tryptic digestion of AAT. (B) The neutral loss chromatogram for loss of 49 Da a precursor ion. (C) The neutral loss chromatogram for loss of 115.5 Da a precursor ion. 217 100 16?1 .7, [Ml-12]” 1089.0. [111111313+ 30 1642.4 20 1850.9 1050 1100 1150 1200 1250 1300 1350 1400 1450 1500 1550 1600 1650 1700 1750 mlz Figure 7.27. This figure displays a full scan spectrum at 35.1 minutes within the LC- MS/MS analysis of the tryptic digestion of AAT 213-287. 218 represent a peptide with a mass of about 3263 mass units, about 47.5 Da larger than the predicted mass (3215.5 Da) of the dPLP peptide, indicating a mass discrepancy between the experimental and predicted masses. While the MS/MS spectra in Figure 7.28 support the presence of dPLP peptides, it appears that there is a mixture of dPLP peptides with different masses. Because the instrument was operated with data dependent settings, it performed CID on m/z 1631.7 first then proceeded on with the next most abundant m/z value and so on. Therefore, the instrument performed CID on the m/z values of 1642.4 Da and 1650.9 Da in addition to m/z 1631.7 Da. These peaks all represent the same peptide, but m/z 1642.4 Da is a sodium adduct and m/z 1650.9 Da is a potassium adduct. The MS/MS spectra of these ions are displayed in Figure 7.28. Figure 7.28A displays the MS/MS spectrum of m/z 1631.9 Da. The base peak in the spectrum corresponds to a loss of 48.9 Da and has been assigned as the neutral loss product of H3PO4 from the +2 precursor. The dPLP product was also observed at m/z 1516.2 (loss of 115.7 Da) as annotated in the spectrum. For this peptide, there were many more fragmentation products formed than typical dPLP peptides have displayed thus far, yet the two common neutral loss products remain the most abundant, especially for the loss of H3PO4. The two other MS/MS spectra in Figures 7.28B and C also demonstrate characteristic dPLP CID behavior. The two neutral loss products (annotated in spectra) are more distinct in these spectra and are also consistent with common dPLP peptide MS/MS spectra. 7.9.4 LC-MS/MS of Glu-C Digestion of Aspartate Aminotransferase 213-287 AAT 213-287 was also digested with Glu-C to determine the site of the dPLP modification. According to Figure 7.16, the theoretical peptide containing the 219 1582.8, [MHz — H3P0J2* 100 A 1517.2, [MHZ - dPLP]?+ 3? 600 0001000 1200 1400 1000 I 1000 I 2000 m a 3 100 1593.4, [MHZ — H,P0J2+ '5 B o > 1526.9, [MHZ - dPLP]2+ '5 \ .53. o ‘ v v ‘1 A; vvvvvv 4" ‘r 1“: [Adv ‘fi-Jf“. ‘2‘“3“ Pl ‘ v ‘5‘ 1L ‘1 vvvvv M 000 000 1000 1200 1400 1600 1000 2000 100 1 0601.4, [1111-12 - H,Po.]2+ C 1536.1, [Ml-12 - 1:1PLPP+ \ .1fijyvr.....“...*v..v‘L|w“1 ...... “.fi 600 800 1000 1200 1400 1600 1800 2000 m]: Figure 7.28. (A) MS/MS spectrum of precursor ion m/z 1631.7 Da. (B) MS/MS spectrum of precursor ion m/z 1642.4 Da. (C) MS/MS spectrum of precursor ion m/z 1650.9 Da. 220 modification should be 250-265 with an average, protonated, theoretical mass of 2 100.3 Da. According to neutral 10Ss chromatograms for loss of 49 Da and 115.5 Da from a +2 precursor (not shown) a dPLP peptide appears to have been detected at about 35.3 minutes into the run. A corresponding full scan spectrum at this retention time is displayed in Figure 7.29A. The peaks at 785.5, 1438.2, and 1632.2 Da were analyzed by CID and zoom scans. All three peaks were determined to be unrelated. However, the MS/MS spectrum of m/z 1632.2 Da, Figure 7298, displays some characteristics of dPLP peptides. The base peak at m/z 1583.0 Da corresponds to a neutral loss of 49.2 Da and the peak at m/z 1516.5 Da corresponds to a neutral loss of 115.7 Da. The fact that both of these neutral losses are consistent with loss of H3PO4 and dPLP and that both peaks are the largest in the spectrum suggest strongly that this is indeed a dPLP-peptide. However, if m/z 1632.2 Da is a doubly charged ion, then the peptide would have a mass of about 3262 Da. This is not the theoretical mass that was anticipated according to Figure 7.16. The zoom scan of m/z 1632.2 not helpful in determining to its charge state which is most likely due to the low abundance of the ion. There is a possible explanation. The dPLP peptide that was expected as a product of Glu-C digestion was 250-265. No such product was detected. However, if the cleavage site at E-265 was skipped by Glu-C, and instead cleaved at the next site, E-276, then the product that would result would be AAT 250-276. If modified by dPLP, this peptide has an average, protonated, theoretical mass of 3267.6 Da. Figure 7.30 illustrates this possibility. The MS/MS spectrum in Figure 293 together with an assumption that m/z 1632.2 Da represents a doubly charged ion of the peptide 250-276 suggests that this is the dPLP peptide. However, the ambiguity over the charge state, the absence of other 221 1 00‘ 785 .5 1632.2 1433.2 l..l|...1u .. L111. “LLJILI‘HIII 111-11111] MINI-J... .11 . 111...” . J 1.. .1... 1. . Relative Abundance -l N (J & 01 01 -J 00 0 NO 0 0 fi 0 0 Figure 7. 1583.0 1516.5 600 300 1000 1200 1400 1600 1800 2000 mlz 29. (A) Full scan spectrum (retention time 35.3 minutes) from the LC-MS/MS analysis of the Glu—C digestion of AAT 213-287. All three annotated peaks are unrelated . (B) MS/MS spectrum of precursor ion m/z 1632.2 Da. 222 related charge states, and the slight error in the measured (deconvoluted) mass (3262 Da relative to the predicted mass 3265.6 Da) of the peptide leaves too much uncertainty to make the assignment in this case. 7.10 Summary and Conclusions This chapter summarized an approach that was developed to determine the site of modification by dPLP in a protein. Chapter 5 discussed the observation that dPLP peptides typically fragment in CID experiments in such a way that two main products are formed. This is evident in the appearance of two main mass spectral features consistent in the MS/MS spectra of these peptides. Two characteristic neutral loss peaks are commonly observed representing the neutral losses of H3PO4 and dPLP. Another characteristic of these spectra is the absence of other fragmentation products. The neutral loss products dominate the MS/MS spectra and no other fragmentation products are represented with any significant abundance. This fragmentation behavior served as the basis for identifying peptides modified by dPLP. In chapter 6, a simple experiment involving dPLP-modified TIP39 was used to assess the possibility of digesting a protein and determining the correct peptide with the modification. With tandem mass spectrometry, we were able to measure peptide mass and perform CID on all of the peptides that eluted separately off of a LC column and analyze the resulting MS/MS spectra for the characteristic neutral loss patterns of dPLP- peptides. The use of neutral loss chromatograms simplified the search for these patterns by pinpointing the retention times where the neutral loss patterns were detected. At this developmental stage, the dPLP-peptides were also identified by mass mapping MALDI 223 dPLP peptide 250—265 MHQMJ = 2100.3 Da A r \ LFCAQ SF SEN FGLYNEBVGNLTVVAKE‘ \ J Y’ dPLP peptide 250-276 MHjW = 3267.6 Da A = Glu-C cleavage sites 5 = suspected dPLP modified residue Figure 7.30. An illustration of the mixture of digestion products possible by cleavage at E-265 or by the missed cleavage at E-276. 224 TOF data to theoretical peptide sequences and masses. The same approach was used for large proteins (~45 kDa) such as Alanine Racemase and Aspartate Aminotransferase. However, no dPLP-peptides were detected with mass spectrometry even after fractionation of the peptides. It was decided that perhaps direct proteolysis of large proteins with trypsin and Glu-C results in too many peptides. This could be significant because PLP contains a phosphate group. Phosphopeptides have been shown to be much more difficult to detect with mass spectrometry relative to non-phosphorylated peptides. Therefore, it could be expected that detecting dPLP-peptides would be difficult in the presence of a large number of unmodified peptides which tend to ionize much easier. An additional step was applied to simplify the digest. By treating the two proteins with cyanogen bromide (CNBr), we were able to break them down into a few large peptides. Since we knew both of the protein sequences and the dPLP modified site in advance, we could predict the mass and peptide sequence that should contain the dPLP modified residue, in each case. We were able to isolate the CNBr products containing the dPLP modification from other CNBr products with HPLC. The fractions corresponding to the dPLP products were confirmed with MALDI TOF and UV-visible spectroscopy. This approach effectively eliminated 75-80% of the protein from consideration. The remaining fraction of the protein, still dPLP modified, was now at a more manageable size for trypsin and Glu-C digestion since fewer products would be formed. Furthermore, tryptic and Glu-C products are at masses more suitable for CID experiments than the large peptides from CNBr treatment. We were still unable to detect any dPLP peptides with MALDI TOF even after utilizing the CNBr step. 225 Figure 7.31 summarizes the final approach which proved to be successful. For both proteins, we were able to digest the suspected dPLP modified CNBr product with trypsin and Glu-C and locate the resulting peptide using LC-MS/MS. This was done by analyzing all of the MS/MS spectra and locating the few that displayed the characteristic neutral loss pattern demonstrated for dPLP peptides. This process was made easier by employing neutral loss chromatograms which were generated by automated searches (neutral loss analysis) for the characteristic neutral losses. Neutral loss spectra alone are not enough to assign dPLP modification to a certain lysine residue. The fragmentation behavior prevents sequencing by tandem mass spectrometry. Therefore, the sequence of the protein in question must be known. The mass of the dPLP peptide must be determined so that it can be matched to a possible sequence within the overall protein sequence. Mass determination requires the determination of the charge state of ions represented as peaks in the spectra. These were achieved with zoom scans of the m/z values. As long as the mass of the peptide matches the theoretical mass of a dPLP peptide, the MS/MS spectrum demonstrates a modification by dPLP, and the theoretical peptide contains only one lysine residue, an assignment of dPLP modification can be made with high certainty. This project represents the only systematic study of dPLP patterns and the first application of tandem mass spectrometry to locate the sight of dPLP modification in proteins. Frey et al. (8, 9) have published two reports where they claim peptides with tandem mass spectrometry. This effort reports the first study of dPLP peptide fragmentation to have identified dPLP-modified residues via mass spectrometry and mass mapping. They did not report any MALDI experiments, but they did have ESI data. They did not have to use the CNBr step reported here to detect the dPLP peptide. 226 Protein, 40 - 50 kDa l NaBE Reduction ———n> CNBr Cleavage >1< Digest (trypsin, Glu-C) 4 HPLC: isolate modified Peptide. l - mass spectrometry - UV-visible spectrospcopy Separate (LC) MALDI — search ESI (MS ) _ search for expected for expected mass (mass ma 111g) fragmentation pp pattern Figure 7.31. Summary of the final approach to locating the site of dPLP modification within a protein. Arrows crossed with x symbolize steps from the initial strategy (Figure 7.1) that were ineffective and eliminated. 227 They were able to detect it directly after proteolytic digestion with LC-MS. Mass spectrometric data, along with UV-visible data, provided a convincing argument that they did detect the correct dPLP modified peptide. Their attempts at sequencing the dPLP peptides with tandem mass spectrometry were unsuccessful. They did offer a tabulated list of product peaks which they claim represent a series of dPLP modified fragment ions. This contradicts our findings which clearly indicate that the dominant peaks in the MS/MS spectra of dPLP represent neutral losses of H3PO4 and dPLP, with very few other products observed. The authors did express a lack of faith in the data, however, and did not offer a mass spectrum among the data presented in their two publications. With protein identification via mass spectrometry becoming almost routine, more efforts will be made to decipher protein structure and function. Those investigators that have already branched out into this field have concentrated almost exclusively on protein phosphorylation. However, there are other co- and post-translational modifications of proteins and more attention will directed toward these as methods for determining phosphorylation sites become more reliable. The project described here is a contribution to that next step. The discovery that a protein is modified by dPLP offers both structural and functional information of the protein. This can be said for other modifications as well. With that in mind, this effort should find application in future studies of dPLP- modified proteins and may represent a template for those searching for other modifications to follow. 228 7.11 Chapter 7 References 1. Sugiyama, Y., Mukohata, Y. Modification of one lysine by pyridoxal phosphate completely inactivates chloroplast coupling factor 1 ATPase. FEBS Lett. 98: 276 (1979) 2. Watababe, A., Kurokawa, Y., Yoshimura, T., Kurihara, T., Soda, K., Esaki, N. Role of lysine 39 of alanine racemase from Bacillus stearothermophilius that binds pyridoxal-5’-phoshate. J. Biol. Chem. 274(7): 4189-4194 (1999). 3. Morino, Y., Nagashima, F. Vitamin Bg Pvridoxal Phosphate ChemicalLBiochemicaL and Medical Aspects, Part A. Edited by David Dolphin, Rozanne Poulson, and Olga Avramovic. John Wiley & Sons, New York, p. 477-497 (1986). 4. Torchinsky, Y.M. Vitamin Bé Pyridoxal Phosphate Chemical, Biochemical. and Medical Aspects, Part B. Edited by David Dolphin, Rozanne Poulson, and Olga Avramovic. John Wiley & Sons, New York, p. 169-222 (1986). 5. Asara, J.M., Allison, J. Enhanced detection of phosphopeptides in matrix assisted laser desorption/ionization mass spectrometry using ammonium salts. J. Am. Soc. Mass Spectrom. 10: 35-44 (1999). 6. Lundbald, R.L. Techniques in Protein Modification. CRC Press, Ann Arbor, p. 51- 54 (1995). 7. Morrison, J .R., F idge, N.N., Grego, H. Studies on the formation, separation, and characterization of cyanogen bromide fragments of human A] apolipoprotein. Anal. Chem. 186: 145 (1990). 8. Frey, P.A., Chen, D. Identification of lysine 346 as a functionally important residue for pyridoxal-5’-phosphate binding and catalysis in lysine 2,3-aminomutase from Bacillus subtilis. Biochem. 40: 596-602 (2001). 9. Frey, P.A., Harma, A., Tang, K. Identfication of a novel pyridoxal-5'-phosphate binding site in adenosylcobalamin-dependent lysine 5,6—aminomutase from Porphyomonas gingivalis. Biochem. 41: 8767-8776 (2002). 229 nljlljjjnjjnjjujjjjl