MECHANISM, FUNCTION, AND INHIBITION OF PEPTIDE DEFORMYLASE
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Kiet T. Nguyen *****
The Ohio State University 2005
Dissertation Committee: Approved by
Professor Dehua Pei, Adviser
Professor Ross Dalbey Dehua Pei Professor Sean Taylor The Ohio State Biochemistry Graduate Program
ABSTRACT
Peptide deformylase (PDF) was originally thought as unique and essential only to the prokaryotes and apparently was absent from the eukaryotes. In this work, two peptide deformylase homologs from the eukaryotic Plasmodium falciparum (PfPDF) and Homo
Sapiens (HsPDF) were cloned and characterized. Both proteins were found to be active and contained similar properties to the Escherichia coli peptide deformylase.
Expression of the PfPDF protein was detected inside the Plasmodium cell, and potent
PDF inhibitors were found to reduce cell growth. The HsPDF was fused with green fluorescence protein (GFP) and revealed mitochondrial localization. Human cell growth study was carried out and was unaffected by potent PDF inhibitors. This work provides evidence that PDF indeed exists in the eukaryotes. However, the role for deformylation in humans was apparently unnecessary, and this work may validate the support for a PDF drug target, as well as providing some preliminary evidence that PDF inhibitors are not toxic to human cells.
The next project involved the identification of a novel class of macrocyclic PDF inhibitors, which tailored from the traditional PDF inhibitors in being acyclic and peptide-like in nature. This new class of PDF inhibitors exhibited potent inhibition and antibacterial properties, against both the Gram-negative and Gram-positive bacteria. The cyclization effect improved the PDF inhibition selectivity and biostability. Here, this
ii work may provide a lead toward more effective PDF inhibitors more resistant to proteolysis and demonstrating improved specificity.
The final part of this work involved the purification and identification enzymes degrading chemotactic f-Met-Leu-Phe peptide (fMLF) released from bacteria. This area of study encompasses the PDF drug effect presented by the commensal bacteria within the mammalian intestines. N-acylpeptide hydrolase and N-acylase 1A from rat small intestine mucosal layer were isolated and identified to degrade the fMLF chemopeptide and other N-formylmethionine peptides. The results may provide support for better understanding the host defense mechanism against the commensal bacteria and to identify the potential problems that may encounter as PDF makes an attractive novel therapeutic drug target.
iii
Dedicated to my parents and foster parents
iv ACKNOWLEDGMENTS
I wish to thank my adviser, Professor Dehua Pei, for his intellectual support, professional guidance, and for having patience with me over the years. I also thank him for the environment in which he had created for a suitable professional training. I am
deeply thankful for my former and present lab members for their stimulating discussion
and providing support on multiple levels in the lab, especially Dr. Kirk Beebe, Mike
Sweeney, Grace Zhu, Junguk Park, and Dr. Xubo Hu. As friends and colleagues, they
have motivated me and provided a special place to conduct my research, an environment
that I embrace with a desire to work long hours and to be able to show up the next day
with the same passion. Dr. Xubo Hu was exceptionally helpful in providing the majority of the PDF inhibitors.
I am grateful and dedicated my life to my wife, Valerie, who during my years of study had believed in me and provided emotional support that could only stem from a loved one. Her understanding and sacrifice had tremendously driven me to succeed and provided a place for refuge whenever I need to resolve research frustrations.
This research was supported by a grant from the National Institute of Health.
v VITA
1997……………………………………...B.A. Biology, Wittenberg University
1997-1999……………………………….Consultant, Materials Directorate, Wright- Patterson Air Force Research Lab
2000-present……………………………. Graduate Teaching and Research Associate, The Ohio State University
PUBLICATIONS
1. Nguyen, K.T., and Pei, D. Purification and Characterization of Enzymes Involved in the Degradation of Chemotactic N-Formyl Peptides. (2005) Biochemistry, submitted.
2. Lucas, R.W., Ingasson, B.P., Krumm, B.E., Hu, X., Nguyen, K.T., Verlinde, C.L.M.J., Pei, D., and Hol, W.G.J. Rational Drug Design of Polypeptide Deformylase Inhibitors in Hemophilus Influenzae: Species Variations in Drug Design. (2005) Protein Science, in preparation.
3. Ingasson, B.P., Lucas, RW., Krumm, B.E., Hu, X., Nguyen, K.T., Verlinde, C.L.M.J., Pei, D., and Hol, W.G.J. Structural Basis for the Inhibtion of Plasmodium Falciparum Deformylase by Macrocyclic peptidomimetics. (2005) Protein Science, in preparation.
4. Nguyen, K.T., Hu, X., and Pei, D. (2004) Slow-Binding Inhibition of Peptide Deformylase by Cyclic Peptidomimetics as Revealed by a New Spectrophotometric Assay. Bioorg. Chem. 32,178-191.
5. Robien, M.A, Nguyen, K.T, Kumar, A., Hirsh, I., Turley, S., Pei, D., and Hol, W. G. J. (2004) An Improved Crystal Form of Plasmodium falciparum Peptide Deformylase. Protein Science 13, 1155-1163.
6. Hu, X., Nguyen, K.T., Vernon, C.J., Lofland, D., Moser, H.E., and Pei, D. (2004) Macrocyclic Inhibitors for Peptide Deformylase: A Structure-Activity Relationship Study of the Ring Size. J. Med. Chem. 47, 4941-4949.
vi 7. Nguyen, K.T., Hu, X., Colton, C., Chakrabarti, R., Zhu, M.X., and Pei, D. (2003) Characterization of a Human Peptide Deformylase: Implications for Antibacterial Drug Design. Biochemistry 42, 9952-9958.
8. Hu, X., Nguyen, K. T., Verlinde, C. L. M. J., Hol, W. G. J., and Pei, D. (2003) Structure-Based Design of a Macrocyclic Inhibitor for Peptide Deformylase. J. Med. Chem. 46, 3771-3774.
9. Deng, H., Callender, R., Zhu, J., Nguyen, K. T., and Pei, D. (2003) Determination of the Ionization State and Catalytic Function of Glu-133 in Peptide Deformylase by Difference FTIR Spectroscopy. Biochemistry 41, 10563-10569.
10. Kumar, A., Nguyen, K. T., Srivathsan, S., Ornstein, B., Turley, S., Hirsh, I., Pei, D., and Hol, W. G. J. (2002) Crystals of Peptide Deformylase from Plasmodium falciparum Reveal Critical Characteristics of the Active Site for Drug Design. Structure 10, 357-367.
11. Bracchi-Ricard, V., Nguyen, K. T., Zhou, Y., Rajagopalan, P. T. R., Chakrabarti, D., and Pei, D. (2001) Characterization of an Eukaryotic Peptide Deformylase from Plasmodium falciparum. Arch. Biochem. Biophys. 396, 162–170.
12. Su, W., Cooper, T. M., Nguyen, K. T., Brandt, M. C., Brandelik, D. M., and McLean, D. G. (1998) Structure-optical Property Relationships of Porphyrins. Proc. SPIE, 3472, 136-143.
FIELDS OF STUDY
Major Field: Ohio State Biochemistry Program
vii TABLE OF CONTENTS
P a g e
Abstract...... ii Dedication...... iv Acknowledgments...... v Vita...... vi List of Tables...... xii List of Figures...... xiii List of Abbreviations...... xv
Chapters:
1. General Introduction...... 1 1.1 Biological Function of Peptide deformylase...... 1 1.2 Peptide Deformylase in Eukaryotes...... 3 1.3 Structure and Mechanism...... 5 1.4 Peptide Deformylase as Novel Drug Target...... 8
2. PDF Assays...... 15 2.1 Introduction...... 15 2.2 FDH-coupled Spectrophotometry Assay (Method A)...... 16 2.3 AAP-coupled Spectrophotometry Assay (Method B)...... 17 2.4 Direct-spectrophotometry Assay (Method C)...... 18 2.5 Development of a New Spectrophotometric Assay (Method D)...... 19 2.5.1 Analytical Methods...... 19 2.5.2 Continuous PDF Assay with DPPI...... 20 2.5.3 PDF Inhibition Assay...... 21 2.6 Results and Discussion...... 21 2.6.1 Assay Design...... 21 2.6.2 Continuous DPPI Coupled Assay...... 22 2.6.3 Application to Screening of PDF Inhibitors...... 24 2.7 Conclusion...... 25
3. Characterization of a Eukaryotic Peptide......
viii Deformylase from Plasmodium falciparum...... 32 3.1 Introduction...... 32 3.2 Experimental Techniques...... 33 3.2.1 Cloning of Plasmodium falciparum PDF cDNA...... 33 3.2.2 Nucleic Acid Isolation and Blot Analyses...... 35 3.2.3 Protein Purification...... 35 3.2.4 PDF Assays...... 37 3.2.5 Western Blot Analysis...... 38 3.3 Results...... 38 3.3.1 Cloning, Expression, and Purification of Recombinant PfPDF.....38 3.3.2 Kinetic Properties of PfPDF...... 40 3.3.3 Spectroscopic Properties of PfPDF...... 42 3.3 4 Inhibition of PfPDF Activity and Malaria Cell Growth...... 43 3.3.5 Expression and Localization of PfPDF...... 44 3.4 Discussion...... 45
4. Characterization of a Human Peptide Deformylase...... 60 4.1 Introduction...... 60 4.2 Experimental Techniques...... 61 4.2.1 Cloning, Expression, and Purification of HsPDF...... 61 4.2.2 Site-Directed Mutagenesis...... 64 4.2.3 Peptide Synthesis...... 64 4.2.4 Synthesis of PDF Inhibitors...... 65 4.2.5 PDF Assays...... 65 4.2.6 Fluorescence Microscopy...... 66 4.2.7 Cell Growth Inhibition Assay...... 67 4.2.8 DNA Synthesis Inhibition Assay...... 67 4.3 Results...... 68 4.3.1 Overexpression and Purification of HsPDF...... 68 4.3.2 Catalytic Properties of HsPDF...... 70 4.3.3 Intracellular Localization of HsPDF...... 71 4.3.4 Inhibition of HsPDF...... 71 4.3.5 Effect of PDF Inhibitors on Growth of Human Cells...... 72 4.4 Discussion...... 73
5. Characterization of Macrocycle PDF Inhibitors...... 84 5.1 Introduction...... 84 5.2 Experimental Techniques...... 85 5.2.1 PDF Inhibition Assays...... 85 5.2.2 Antimicrobial Susceptibility Testing...... 87
ix 5.2.3 Inhibition of MMPs...... 87 5.2.4 Antimicrobial Susceptibility Testing...... 88 5.2.5 In Vitro Stability of PDF Inhibitors in Rat Plasma...... 89 5.3 Results and Discussion...... 90 5.3.1 Slow-binding Inhibition of PDF...... 91 5.3.2 In Vitro Antibacterial Activity...... 93 5.3.3 Cyclization Improves Inhibitor Selectivity...... 94 5.3.4 Cyclization Improves the Stability of Inhibitors...... 95 5.4 Conclusion...... 95
6. Purification and Characterization of Enzymes Involved in the...... Degradation of Chemotactic N-Formyl Peptides...... 106 6.1 Introduction...... 106 6.2 Experimental Techniques...... 108 6.2.1 Purification of fMAP...... 108 6.2.2 Purification of fMDF...... 109 6.2.3 Synthesis of N-Formyl-methionyl-p-Nitroanaline...... 111 6.2.4 fMAP Assay...... 112 6.2.5 fMDF Assay...... 112 6.2.6 Gel-Filtration Analysis...... 113 6.2.7 In-Gel Digestion...... 113 6.2.8 Nano-LC MS/MS...... 114 6.3 Results...... 115 6.3.1 Purification of fMAP and fMDF...... 115 6.3.2 Identification of fMAP and fMDF by...... Mass Spectrometer Fingerprinting...... 117 6.3.3 Catalytic Properties and Inhibtion of fMAP (APH)...... 118
6.3.4 Catalytic Properties and Inhibition of N-Acylase IA (fMDF).....119 6.4 Discussion...... 120
7. Materials and General Methods...... 134 7.1 Buffers...... 134 7.2 Materials...... 135 7.3 Synthesis of Substrates for DPPI Assay...... 137 7.4 General Biochemical and Biological Methods...... 140 7.4.1 Materials...... 140 7.4.2 Growth Media...... 141 7.4.3 Growth and Storage of Bacterial Strains...... 142 7.4.4 Preparation of Competent Cells...... 143
x 7.4.5 Extraction with Organic Solvents...... 143 7.4.6 Quantitation of DNA and RNA...... 144 7.4.7 Removal of Small Molecules from High-molecular-weight...... DNA and RNA...... 144 7.5 Electrophoresis...... 145 7.5.1 Agarose Gel...... 145 7.5.2 Polyacrylamide Gels for Protein Separation...... 145 7.6 Recombinant DNA techniques...... 147 7.6.1 Restriction Digestions...... 147 7.6.2 Filling Recessed 3’-Termini and Removing Protruding...... 3’-Termini...... 147 7.6.3 Removal of 5’ Phosphates...... 148 7.6.4 Ligation of DNA...... 148 7.6.5 Transformation...... 148 7.6.6 Small-Scale Preparation of Plasmid DNAs...... 149 7.6.7 Mutagenesis...... 150 7.6.8 Sequencing...... 151 7.7 Purification of Peptide Deformylase (wild type from E. coli)...... 151 7.7.1 Cell growth and induction...... 151 7.7.2 Q-Sepharose purification...... 152 7.7.3 Cell Lysis...... 152 7.7.4 Ammonium sulfate precipitation...... 153 7.7.5 Phenyl Sepharose purification...... 153 7.7.6 Storage...... 154 7.7.7 Protein Quantitation...... 154 7.8 Purification of Substituted Zn(II)-, Fe(II)-, and Co(II)-PDF...... 154 7.8.1 Cell Growth...... 154 7.8.2 Buffers...... 155 7.8.3 Protein Purification...... 155 7.8.4 Purification of His-tagged PDF...... 156
Appendix...... 158
Bibliography...... 160
xi LIST OF TABLES
Table Page
3.1 Comparison of PfPDF Variants...... 49
3.2 Kinetic Constants of Ni-PfPDF(∆N63)-HT toward...... Various Peptide Substrates...... 50
4.1 HsPDF Activity against f-ML-pNA and Human Mitochondrial Peptides...... 76
4.2 Catalytic Activity of Wild-Type vs. Mutant HsPDF and EcPDF...... 77
4.3 Inhibition Constants of EcPDF and HsPDF...... 78
5.1 Inhibition Constants against E. coli PDF...... 97
5.2 In Vitro Antibacterial Activity of PDF Inhibitors...... 98
5.3 Inhibition of MMPs...... 99
6.1 Purification of fMAP from Rat Epithelial Mucosal Layer...... 124
6.2 Purification of fMDF from Rat Epithelial Mucosal Layer...... 125
6.3 MS Analysis of N-acylaminoacyl-peptide Hydrolase under Reducing and...... Denaturing conditions Following.Digestion with Trypsin...... 126
6.4 MS/MS Analysis of N-Acylase I under Reducing and Denaturing...... Conditions Following Digestion with Trypsin...... 127
6.5 Kinetic Activity of N-acyl Aminopeptidase from Rat Intestine Epithelial...... Mucosal Layer...... 128
6.6 Kinetic Activity of N-Acylase I from Rat Intestine Epithelial Mucosal...... Layer...... 129
xii
LIST OF FIGURES
Figures Page
1.1 Protein Biosynthesis in Bacteria, Mitochondria, and Plastids...... 12
1.2 Crystal Structures of PDF complex...... 13
1.3 Proposed catalytic mechanism of PDF...... 14
2.1 PDF Assays...... 27
2.2 Structures of PDF inhibitors...... 28
2.3 Reactions involved in the assay...... 29
2.4 (A) Time courses for the hydrolysis of f-Met-Lys-AMC by PDF...... 30 (B) Plot of initial rates against f-Met-Lys-AMC Concentration...... 30
2.5 Inhibition of PDF by BB-3497...... 31
3.1 Comparison of the deduced PfPDF amino acid sequence with other PDF...... sequences...... 51
3.2 Coomassie blue-stained SDS–PAGE gel (15%) showing purified PfPDF...... Variants...... 52
3.3 Effect of pH on Co-PfPDF(∆N63)-HT Activity...... 53
3.4 Comparison of the electronic absorption spectra of Co-PfPDF(∆N57)-HT...... 54
3.5 Structures of PDF Inhibitors...... 55
3.6 Inhibition of P. falciparum cell growth in erythrocyte culture by PDF...... inhibitor 1...... 56
3.7 Expression of PfPDF during intraerythrocytic malaria life cycle...... 57
xiii
3.8 E. coli and P.falciparum PDF structural Comparison...... 58
3.9 Comparison of the Active Sites in E. coli PDF and P. facifparum PDF...... 59
4.1 Sequence Alignment of Various PDF’s from Eukaryotic and Prokaryotic...... Organisms...... 79
4.2 15% SDS-PAGE gel showing the purity of HsPDF(∆N58) during different...... stages of purification...... 80
4.3 Intracellular Localization of HsPDF-GFP fusion in HEK cells...... 81
4.4 Lineweaver-Burk plot for the inhibition of HsPDF by BB-3497...... 82
4.5 Effect of BB-3497 on human cell growth...... 83
5.1 Structures of PDF inhibitors...... 100
5.2 Slow-binding inhibition of PDF by compound 1c...... 101
5.3 Electronic absorption spectra of Co(II)-substituted PDF in the absence and...... presence of inhibitor 1b...... 102
5.4 Crystal Structure of HiPDF/Macrocycle 1d Complex...... 103
5.5 Inhibition of B. subtilis (A) and E. coli (B) cell growth by inhibitor 1c...... 104
5.6 Comparison of the in vitro stability of PDF inhibitors 1f (cyclic) and 3 (acyclic).... in rat plasma...... 105
6.1 Enzymes from rat small intestine epithelial mucosal layer involved in...... degradation of N-formylmethionine peptides...... 130
6.2 Chromatography of an N-formylaminopeptidase hydrolytic enzyme...... 131
6.3 SDS-PAGE of rat intestinal N-acyl aminopeptidase...... 132
6.4 SDS-PAGE of rat intestinal N-acylase I...... 133
xiv LIST OF ABBREVIATIONS
AAP Aeromonas aminopeptidase
Ac-Met acetyl-methionine
Ac acetyl
APH N-acylpeptide hydrolase br broad (IR and NMR)
β beta n-Bu normal-butyl
t-Bu tert-butyl
Bz benzoyl
°C degrees Celsius
calcd calculated
δ chemical shift in parts per million downfield from tetramethylsilane
d doublet (spectra); day(s)
DBU 1,8-diazabicyclo[5.4.0]undec-7-ene
DEPC diethylpyrocarbonate
DMAP 4-(N,N-dimethylamino)pyridine
DMF N,N-dimethylformamide
DMSO dimethylsulfoxide
DPPI dipeptidyl peptidase I xv eq. equivalent
EcPDF Escherichia coli PDF
Fmoc 9- fluorenylmethoxycarbonyl fMLP N-formyl-methionine-leucine-phenylalanine fM-pNA N-formyl-methionine-para-nitroanalide
FDH Formate Dehydrogenase g gram(s)
GFP green fluorescence protein
GST gluthathione-S-transferase h hour(s)
HBTU 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorphosphate
HEK human embryonic kidney
HEPES N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid
HsPDF Homo sapiens PDF
IPTG isopropyl-β-D-thiogalactopyranoside
J coupling constant in Hz (NMR) k kilo
L liter(s) m milli; multiplet (NMR)
µ micro
xvi M moles per liter
MAP methionine aminopeptidase
Me methyl
MES 2-morpholinoethanesulfonic acid
MHz megahertz min minute(s) mol mole(s)
MS mass spectrometry; molecular sieves m/z mass to charge ratio (MS)
NAD+ nicotinamide adenine dinucleotide
NMR nuclear magnetic resonance p para
PDF peptide deformylase
PfPDF Plasmodium falciparum PDF
Ph phenyl ppm parts per million rt room temperature s singlet (NMR); second(s) t tertiary (tert) t triplet (NMR)
xvii TLC thin layer chromatography
TMS trimethylsilyl
Tris tris (hydroxymethyl) aminomethane
xviii
CHAPTER 1
GENERAL INTRODUCTION
1.1 Biological Function of Peptide Deformylase
The prokaryotic and eukaryotic organelle (mitochondrial and chloroplast)
ribosomal protein synthesis universally initiates with an N-formylmethionine residue
(Fig. 1.1). In its initiation complex, key associated components, along with the messenger
Met RNA, are the initiator fMet-tRNAf , the initiation factors IF1, IF2, GTP, IF3 and the
30S and 50S ribosomal subunits, small and large ribosomal subunits. Once the methionyl
Met Met residue is esterified at the 3’ end on the initiator tRNAf (1, 2), the methionine-tRNAf
Met molecule is N-formylated by a methionyl-tRNAf formyltransformylase. The reaction proceeds through the transfer of a formyl group from N10-formyltetrahydrofolate to the α- amino group of the methionine residue (3) (Fig. 1.1). The formylation of the methionine-
Met tRNAf molecule is crucial for its initial recognition by IF2, and ultimately the complex
is conferred by the P site of the 30S ribosome to proceed with the initiation of protein
translation (4, 5). After the onset of initiation, elongation of protein translation proceeds
through the displacement of IF2 by EF-Tu molecules, which recognize the rest of the
unformylated aminoacylated tRNA molecules. This interaction complex is necessary to
participate in the ongoing translation process to ultimately produce a complete
polypeptide transcript.
1 In all prokaryotes, a formyl-methionine residue is inherently incorporated at the
N-terminus of all nascent polypeptides; however, the vast majority of the N-formyl moiety is removed from the mature proteins. The removal of the N-formyl moiety is catalyzed by peptide deformylase (PDF) (Fig. 1.1) (6, 7). But, significantly only half of all the polypeptides retain the methionine at their N-terminus (8). The removal of the N- terminal methionine residue from the nascent polypeptides is catalyzed by the action of methionine aminopeptidase (MAP) (8). It has been shown by genetic complementation, gene knockout, that MAP is essential in E. coli (9). The removal of the methionine residue by MAP depends on the penultimate amino acid side chain (8). The preference had been revealed showing MAP reaction is more efficient when the penultimate amino acid side chain is either Gly, Ala, Pro, Thr, Ser, Val, or Cys (8, 10). However, MAP action is inhibited when the N-terminal methionine residue of the polypeptide is blocked by an N-formyl group (8, 10). Here, PDF plays, at least, a critical role in mediating the maturation process of the nascent polypeptides, partly due to the necessity of removing the N-formyl group to render the nascent polypeptides for MAP cleavage of the N- terminal methionine residue. In fact, the disruption of the fms gene encoding PDF in E. coli proved critical for cell survival (11, 12), making it an attractive drug target.
The requirement for N-formylation in protein translation is somewhat unclear.
Thus far, N-terminal formylation is a unique feature of eubacterial, mitochondrial, and plastid translation systems. Formylation helps mediate the N-formylmethionine-
Met tRNAf initiator’s recognition by the IF2 and to discriminate from other elongation apparatus, but the formylation process is speculated to be nonessential for bacterial survival (13). However, cell growth is decreased by ~3-12-fold in lower growth rate and
2 is temperature sensitive upon disrupting the methionyl-tRNA formyltransformylase gene
(fmt) (13, 14); thus indeed formylation is deemed not strictly essential for bacterial survival (13, 14). In other words, the genetic complementation did not comprise cell growth completely, suggesting that protein translation in this bacterial strain was capable to support cell growth by bypassing the N-formylmethionine initiation.
1.2 Peptide Deformylase in Eukaryotes
Protein synthesis in the cytoplasm of eukaryotes does not begin with N- formylmethionine, and undoubtedly the need for deformylation in the cytoplasmic process is apparently absent. Since its inception in the 1960’s and recent molecular cloning and genomic sequencing, over 100 putative peptide deformylating genes (def) have been identified throughout the prokaryotes, as well as homologs in the nucleus of a wide range of completed genomes from the lower eukaryotes, the Protista kingdom, to the more complex higher eukaryotes, the Animalia kingdom. However, the majority of the discovered PDFs are from bacterial genomes, some having one copy and in certain cases two different copies are present (15). Thus, deformylation remains to be a conserved and an essential process in all eubacteria; whereas the disruption of the def gene rendered a lethal condition in E. coli (11, 12), Thermus thermophilus (16) and
Staphyloccocus aureus (17). Despite its diversity, there are no PDF homologs found in the complete genomes of the archaea, Saccharomyces cerevisiae(18), or in
Caenorhabtidis elegans (18).
Of all the data accumulated thus far, the need for deformylation has been proven crucial in bacteria. Deformylation is apparently absent from mitochondrial protein
3 translation in yeast (19-24), Neurospora (25, 26), honey bee (27), and bovine (28-32), where their N-formylmethionine moiety is retained. To complicate the matter further, recent data from genome sequencing results have identified putative PDF-like sequences in higher plants such as Arabidopsis thaliana, tomato, corn, and rice. More importantly, deformylation has been shown to occur in chloroplasts of plants (33-35). The cDNA of
Arabidopsis thaliana PDF (AtPDF) has been cloned and overexpressed in E. coli (15).
AtPDF is catalytically active and shares many properties of bacterial PDF, and it is shown to be subcellular localized in the chloroplast and mitochondrion (15, 36).
Additional PDF-like homologs in eukaryotes have increasingly been revealed. In addition to plant, several eukaryotic plastid-containing parasites such Plasmodium falciparum trypanosomal species have a nuclear encoded PDF-gene homolog (18). From sequence alignment, the PDF from Plasmodium falciparum (PfPDF) shows a 33% identity to the bacterial PDF, plus a ~60-amino acid extension at the N-terminus. Thus, it is conceivable that the extra extension is a bipartite N-terminal pre-sequence that consists of a signal peptide for entry into the secretory pathway and a plant-like transit peptide for subsequent import into the apicoplast of the Plasmodium. The PfPDF localization was
presumed to be involved in the apicoplast protein translation processes, similar to the
prokaryotic translation system. More importantly, PDF-like sequences have been
identified in the genomes of Drosophila Melangogaster, as well as in the partial
expressed sequence tags (ESTs) of mouse and human (15). Like PfPDF, all of these eukaryotic PDF sequences contain an extra ~60 amino acid N-terminal extension.
Finally, these PDF-like gene sequences are not known to be expressed and, if expressed
at all, to encode a functional PDF. The role and function of PDF in eukaryotes have not
4 been established. Part of this research is to identify a link between the PDF-like homologs in the eukaryotes and the bacterial PDF.
1.3. Structure and Mechanism
Since first reported by Adams in 1968 (7), PDF has eluded detailed characterization studies due to its extreme instability. The cloning of the deformylase gene, def, in 1993 by Meinnel et al. (12) greatly facilitated our mechanistic understanding of PDF. The PDF protein was first overexpressed in E. coli and purified to apparent
-1 -1 homogeneity. The recovered activity was extremely poor (kcat/KM of 80 M s against
formyl-Met-Ala-Ser substrate) but showed one zinc ion per polypeptide (37). However
in 1997, Rajagopalan et al. (38) reported the purification of a highly active PDF fraction
4 -1 -1 under oxygen-free conditions, with a kcat/KM on the order of 2.9 x 10 M s against the
formyl-Met-Ala-Ser substrate, and contained one ferrous ion per polypeptide.
Subsequently, the Fe2+ was confirmed to be the metal ion cofactor in PDF (39, 40). The
extreme lability of the ferrous-containing PDF was due to the oxidation of the ferrous ion
to ferric, via atmospheric oxygen species (41). This finding explained the initial results
obtained by Meinnel et al. (12), where during their purification, when exposed to air, the
Fe2+ metal ion was oxidized and replaced by Zn2+ ion, which contributed to the observed
lower catalytic activity.
For ease of enzymatic study, the ferrous-containing PDF can be replaced by a
divalent metal such as Ni2+ (40) or Co2+ (42) to produce a highly stable and fully active enzyme, as well as enabling spectroscopic studies. A worthy note here is that the enzyme, in both nickel-containing and cobalt-containing PDF, is oxygen insensitive and
5 catalytically as active as the ferrous-containing PDF. It is important to add, here, that
these surrogate PDFs made possible the biochemical characterization, through structural
and mechanistic studies.
PDF is widely accepted as a novel class of iron metalloenzyme that is related in
structure to the metalloproteinase superfamily. While other regions of its sequence may
vary, all PDFs share three highly conserved motifs GΦGΦAAxQ, EGCΦS, and HEΦDH
(where Φ encodes any hydrophobic amino acid and x is any amino acid). These motifs form the three sides of the active site pocket, a common architecture evidently conserved among all PDFs. Several X-ray crystal and NMR solution structures of Fe-PDF, Zn-
PDF, and Ni-PDF(free) forms have been determined (43-47) . In each case, structural conservation was apparent through its metal ion tetrahedrally coordinated by two histidine residues of the conserved HEΦDH motif, one cysteine residue from EGCΦS motif, and a water molecule (Fig. 1.2A) (45). The co-crystallization of PDF (Fe-PDF,
Zn-PDF, and Ni-PDF) with a reaction product (Met-Ala-Ser) revealed that the P1’ site
having appropriate volume to support a hydrophobic interaction with the methionine side
chain (43). More importantly, key interactions of the P1’ and P2’ are predominantly
formed from hydrogen bonding between the carbonyl and amide protons of the peptide
backbone. Further, the co-crystallization of E. coli Zn-PDF (48) with a transition state
analog, (S)-2-O-(H-phosphonoxy)-L-caproyl-L-leucyl-p-nitroanilide (PCLNA) (Fig
1.2B), additionally revealed crucial interactions with the H-phosphonate moiety. The
crystal complex showed one of the phosphonate oxygen is liganded to the metal,
replacing the water molecule, as well as interacting with the side chain of the E-133. The
other phosphonate oxygen, which mimics the carbonyl oxygen of the formyl moiety, is
6 within hydrogen bonding distance with the polypeptide amide of L-91 and the side chain
NH2 of Q-50. The (L)-caproyl group in the S1’ position, which mimics the methionine
amino acid, sits in a pocket generated by residues Gly-43, Ile-44, Gly-45, Glu-88, Cys-
129, His-132, and Glu-133. In the S2’ position, the (L)-leucine side chain makes van
der Waals contact with Leu-91 and the mainchain carbonyl groups of residues Glu-42 and Gly-43 on one side, while the other side is exposed to solvent. Site-directed
mutagenesis of the key catalytic residue E-133 (in the HEΦDH motif) to Ala and Asp
provides additional insights into PDF’s catalytic mechanism (42). Under the absorption
spectra of various pH conditions, the metal-bound water of the cobalt-substituted mutants
E133A and E133D was shown to have pKa values of 6.5 and 5.6, respectively. Together, it was suggested that the role of the metal ion is to ionize the bound water molecule to
generate a metal-bound hydroxide, which nucleophilically attacks the formyl moiety. In
addition, the pKa, 6.5, of the metal-bound water in E133A suggested that the most
important role of E133 side chain involvement in catalysis is to serve as a H+ shuttle as
well as general acid mechanism, donating a proton to the leaving amide ion of the peptide
(42).
A proposed mechanism of catalysis is depicted in Figure 1.3. In the free enzyme,
the metal-bound water molecule is hydrogen bonded to the side chain of E-133.
Catalysis occurs when the N-formyl moiety of the peptide substrate enters the active site,
. making an E S complex, to position next to the metal-bound water. Since the pKa values of the metal-bound water (6.5) and the carboxyl group of E-133 (~5) are similar, the shared proton may be transferred from the water to the carboxylate group. The reaction presumes to occur instantaneously due to the readily formed hydroxyl species,
7 coordinated to the metal, via by performing nucleophilic attack on the formyl group. The
resulting transition state tetrahedral intermediate, producing an oxyanion derived from
the formyl group, is stabilized by a hydrogen bonding network between the NH backbone
of L-91 and the side chain NH2 of Q-50. The breakdown of the tetrahedral intermediate is facilitated by reforming the original C=O bond. During the second transition state, the leaving NHR is likely hydrogen bonded to the side chain of E-133. A proton transfer process from E-133 to the leaving –HNR group and subsequently is followed by a water- formate exchange to complete the catalytic cycle.
1.4 Peptide Deformylase as Novel Drug Target
The discovery of antibiotics is one of the greatest findings of the 20th century.
Despite all advances in the field, problems arise in the proven ability of bacteria to adapt and to develop resistance to multiple classes of existing antibiotics (49). The emergence of antibiotic resistance bacteria is a question of not “if,” but “when” will the antibiotic resistance occur. To get a sense of the seriousness of this matter, early resistance to multiple drugs was reported among the enteric bacteria, namely, Escherichia coli,
Shigella and Salmonella, in the late1950s to early 1960s (50-52). Recent resistance has been reported in the nosocomial S. aureus strains, both methicillin-resistant (MRSA) (53)
and multidrug-resistance (MDR) (54). More deaths are associated with MRSA than with
methicillin-sensitive strains (55). A steadily increasing, small proportion of MRSA also
now shows low-level resistance to vancomycin (56, 57). These problems pose an urgent challenge to find new avenues to combat these antibiotic resistant bacteria. Therefore, the development of new antibiotics is essential, i.e. by blocking or circumventing the
8 resistance mechanisms or identifying novel targets. Consequently, there has recently been a renewed interest from the pharmaceutical industry to develop novel antibiotics to combat the resistant bacteria. The analysis of increasingly available microbial genomic data has revealed an abundance of potential novel drug targets. One novel target has recently received tremendous recognition and success, peptide deformylase.
Since PDF is required for bacterial survival but apparently unnecessary in animal cells, it provides an attractive target for a novel antibacterial strategy. Moreover, deformylation is a conserved feature throughout the eubacterial kingdom. With the numerous three-dimensional structures of various bacterial PDFs solved (58-62), both
with and without enzyme-bound inhibitor complexes, it appeared that the active site of
PDF is conserved, composing of sequence motifs, motif 1 [GφGφAAXQ], motif 2
[EGCLS], and motif 3 [HEφDH] (where φ is a hydrophobic amino acid and X is any
amino acid) (46, 63). Consequently, PDF appears to be a broad spectrum novel antibiotic
drug target.
One notable advance in PDF inhibitor screening came from the random
screenings by Versicor and Hoffmann-La Roche (64, 65). The work led to the
identification of actinonin, a naturally occurring hydroxamic acid derivative known to
have potent antibacterial activity and it is a PDF target. In vitro activity of actinonin
against PDF is within KI value of 0.3 nM and demonstrated bacteriostatic activity against a wide spectrum of Gram-positive and Gram-negative bacteria. Evidence to support the molecular target for antibacterial activity of actinonin came from the E. coli genetic
construct in which mutant E. coli strain expresses controllable levels of PDF (65). Thus,
the susceptibility to the presence of actinonin is associated with the different expressed 9 levels of PDF within the bacterial cell (at high PDF expression level, the bacterial cell
becomes more tolerable to the presence of actinonin) (65). Despite having strong inhibition for PDF, actinonin was never developed for the treatment of infections due to its poor bioavailiability and consequent lack of in vivo efficacy (66) .
Owing to a structural lead from actinonin, BB-3497 was developed to target PDF
(KI = 9 nM) with moderate antibacterial activity against a range of bacterial pathogens
followed by oral administration (67). BB-3497 was administered in doses to mouse
model carrying a systemic S . aureus infection. Mice were rescued from infection upon
intravenous or oral treatment with BB-3497 at a median effective dose (ED50) of 7 and 8
mg/kg, respectively. When tested orally against a methicillin resistant strain of S. aureus
(MRSA), BB-3497 had an ED50 of 14 mg/kg, which was competively as effective as
ofloxacin, which had an ED50 of 10 mg/kg (67). Similarly in a separate study, VRC
3375, a potent PDF inhibitor, had ED50s of 32 and 21 mg/kg by intravenous and oral
routes, respectively (68). Together, these results strongly demonstrated for the first time
that PDF was the target for novel antibiotic for the treatment of bacterial infections. More
importantly, an inhibitor BB-86398 from British Biotech is currently in phase I clinical trials.
Despite efforts to broaden PDF inhibitor design leading to very potent in vitro activity (subnanomolar range), relatively high frequency of resistance against PDF inhibitors have recently been reported in actinonin-treated Staphylococcus aureus (a resistance rate of ~107) (17). The arisen viability due to resistance was conferred by early
study in the fmt/def double knockout mutants (13, 14), in which the bug was able to
bypass the formylation-deformylation pathway. Due to the fact that these resistant
10 mutants all have a slow growth phenotype similar to that of the fmt-/def – strains (17, 64), it would, by no means, deter the pursuit of PDF as a novel antimicrobial drug target.
11
Methionine (Met)
Aminoacyl-tRNA tRNA Met Synthetase F Met-tRNAMet F Formate (from N10- Transformylase formyltetrahydrofolate)
Met N-formyl-Met-tRNA F (Protein Initiation)
N-formyl-Met-AA1-AA2-AAn-tRNA
Peptide formate Deformylase Met-AA1-AA2-AAn-tRNA Methionine Aminopeptidase Methionine AA1-AA2-AAn-tRNA
AA1-AA2-AAn (Mature Protein)
Figure 1.1. Protein biosynthesis pathway in bacteria, mitochondria, and plastids. Met, methionine; AA, amino acids.
12
A B
Figure 1.2. Crystal structures of PDF (A) E. coli Zn-PDF structure, (B) E. coli Zn-PDF- PCLNA complex
13
Figure 1.3. Proposed catalytic mechanism of PDF.
14
CHAPTER 2
PEPTIDE DEFORMYLASE ASSAY
2.1 Introduction
The need for an effective PDF assay is crucial to facilitate the mechanistic study
of PDF and inhibitor screening. Fostering the development of PDF assay, work done by
Hu et al (69) determined the substrate specificity through the screening of a
combinatorial N-formyl tetratpeptide library. Of which, a consensus sequence of formyl-
Met-X-Z-Tyr (X = any amino acid except for aspartate and glutamate; Z = lysine,
arginine, tyrosine, or phenylalanine) substrates was determined to be efficiently
deformylated by PDF, and as well as a substrate consisting formyl-Phe-Tyr-(Phe/Tyr)
peptide. Thus, the study suggested that PDF has broad substrate specificity, capable of deformylating a wide range of N-formyl peptide substrates. Prior to this work, there were three major PDF assays employed and all of them depend on PDF removal of the N- formyl group: 1) FDH-coupled PDF reaction; 2) AAP-coupled PDF using f-M-L-pNA substrate; and 3) direct assay utilizing N-formyl-β-thiaphenylalanyl peptide substrate.
All of these assays are depicted in Figure 2.1 and will be discussed below.
The major reason why there are so many assays thus far is that there are problems encountered within each assay. These problems will be revealed as each assay is discussed below. However, having several different assay methods makes the
15 characterization of PDF much more meaningful. To make our data more reliable, when possible, we often confirmed the results with a second assay method. One of the major parts of this work was to screen and characterize potent PDF inhibitors. Our need for a new assay stemmed from our recent PDF inhibitor development where we have synthesized a macrocyclic PDF inhibitor 1 (compound 1b in Chapter 5, (Fig. 2.2)), in which the P1’ and P3’ side chains are covalently cross-linked (will be later discussed in
more details in Chapter 5). The macrocycle acts as a potent, slow-binding inhibitor. We experienced major difficulties in its kinetic characterization using the existing assays. In section 2.5, we have developed an additional PDF assay method by employing the reaction dipeptidyl peptidase I (DPPI, also called cathepsin C), a cysteine protease, as the coupling enzyme.
2.2 FDH-Coupled PDF Assay (Method A)
The robustness of PDF to accommodate various N-formylmethionine peptide substrates makes this FDH-coupled assay to be very effective in characterizing PDF activity against the natural peptide substrates, which are different from the synthetic substrates used in the colorimetric assays of Method B, C, and D (Section 2.5). The reaction utilizes the activity of Candida boidinii formate dehydrogenase (FDH) in a
continuous fashion (Fig. 2.1, Method A) (38, 70). Upon the release of the formyl moiety
from the N-formylmethionine peptide, FDH oxidizes the released formate to CO2, while reducing NAD+ to NADH. The rate of deformylation of a substrate by PDF is directly
proportional to the rate of production of NADH. The formation of NADH is monitored
visibly in a continuous fashion on a UV-Vis spectrophotometer at 344 nm (ε 344 nm = 6200
16 M-1cm-1). Typically, the Co2+-PDF activity for f-Met-Ala-Ser was 2.9 x 104 M-1s-1 (38).
However, FDH is sensitive to high salt content (NaCl), which cannot be used to adjust
the ionic strength of the buffered solution. Typically, FDH activity will be inhibited at
>50 mM NaCl concentration. In addition, due to FDH’s low specific activity for formate
(KM of 2.5 mM), the reactions often showed an initial early lag phase (0-20 s), which
could be difficult to characterize the slow-binding inhibitors.
2.3 AAP-Coupled PDF Assay (Method B)
In search of a more sensitive PDF assay to facilitate the characterization and
mechanistic studies, this assay utilizes Aeromonas aminopeptidase (AAP) as the coupled
enzyme (Fig 2.1, Method B) (71). Aeromonas aminopeptidase from A. proteolysis was
purified in our lab by the Prescott method (72). The reaction uses f-Met-Leu-p-
nitroanilide (f-ML-pNA) as substrate. Here, AAP does not have activity against the f-
ML-pNA, which makes it an ideal coupling enzyme. Until the removal of N-formyl
group by PDF from f-ML-pNA to release ML-pNA, then AAP is able to catalyze the
-1 -1 release of the chromogenic para-nitroaniline, pNA, (ε 405 nm = 10,600 M cm ). The
deformylation activity of f-ML-pNA by Co2+-PDF followed Michaelis-Meten kinetics
-1 6 -1 -1 with a KM = 20.3 ± 1.3 mM, kcat = 38 ± 2 s , and kcat/KM = 1.9 x 10 M s (71). Several
advantages gained from this assay: sensitivity, convenience (a continuous fashion), and
relatively inexpensive. It is, thus, the primary assay used predominantly to carry out PDF
reactions. A minor limitation of this continuous assay is the difficulty in examining the
sequence specificity of the deformylase, which is limited to substrate preference by AAP.
In addition, the AAP-coupled assay carries with it some inherent problems due to its
17 requirement for divalent metal ion (Zn2+) for catalysis. A major interest in PDF study is
the novel antimicrobial drug target. The design and screening of PDF inhibitors at the
basic research level needs an effectively sensitive assay. Since PDF is a metalloenzyme,
essentially all PDF mechanism-based inhibitors are designed to chelate the active site
metal. A problem in using this assay for inhibitor screening is that PDF inhibitors also
inhibit AAP at low concentrations (in the sub-micromolar range). Therefore, this assay is
limited to substrate kinetic characterization and cannot be used in a continuous fashion to
carry out inhibition studies.
2.4 Direct Spectrophotometic Assay (Method C)
One of the most intriguing assays used to characterize PDF activity is a method
developed in our lab that does not depend on a coupling enzyme. It is a direct
spectrophotometric assay that utilizes an N-formyl-β-thiaphenylalanyl peptide as
substrate (Fig. 2.1, Method C) (73). Upon the removal of the N-formyl group by PDF,
the compound spontaneously releases a free thiol group and will spontaneously react with
5,5’-dithio-bis(2-nitrobenzoic acid) (DTNB) to produce 5-thionitrobenzoic acid. The
reaction is monitored for the release of 5-thionitrobenzoic acid in a continuous fashion on
a spectrophotometer at 412 nm (ε = 14,000 cm-1M-1). The deformylation activity (Co2+-
-1 PDF) for N-formyl-β-thiaphenylalanyllysyl-p-nitroanilide had a kcat = 86 ± 6 s , KM = 27
6 -1 -1 ± 6 mM, and kcat/KM = 3.2 x 10 M s (73). This method is very sensitive and could be
used for inhibitor screening. There are several problems that exist with employing this
PDF assay. The first is DTNB also reacts with cysteines on the protein surface, sometime
rendering the protein reduced activity. Secondly, the assay cannot be used when β-
18 mercaptoethanol or dithiothreitol are used as reducing reagents, which readily react with
DTNB. Another problem is the difficulty in synthesizing the substrate, which involves
multiple synthetic routes and it is prone for hydrolysis.
2.5 Development of a New Spectrophometric Assay (Method D)
To avoid such problems encountered with the assays described above (Method A,
B, and C), here a new assay method (Method D) was developed that coupled the reaction
of capthepsin C (DPPI) (Fig. 2.3). The new assay greatly improved our ability to carry
out inhibition studies where several inhibitors exerted activity in the subnanomolar
concentration, a condition that requires a highly sensitive assay method. The utility of
the assay was revealed by employing this assay to obtain PDF inhibition constants. In
addition to demonstrating the robustness of this assay method, we used this method to
characterize the slow-binding PDF inhibitors, i.e. inhibitor 1, Fig 2.2, which will be
described later in Chapter 5.
2.5.1 Analytical Methods
Protein concentration was determined by Bradford assay using bovine serum
albumin as the protein standard. For PDF, the total protein concentration as determined by Bradford assay was corrected by a factor of 0.71 (42). Substrate concentrations were determined by base hydrolysis followed by measuring the absorbance of 7-amino-4- methylcoumarin at 360 nm (ε = 1.7 x 104 M-1cm-1).
19 2.5.2 Continuous PDF Assay with DPPI
Prior to use, DPPI was activated by treating with 5 mM dithiothreitol (DTT) for
30 min in 50 mM Hepes (pH 7.0), 10 mM NaCl. Unless stated otherwise, all assays were
performed at room temperature (25 ºC). Assay reactions were performed in a quartz
microcuvette (total reaction volume = 200 µL) containing 5–150 µM substrate, 50 mM
Hepes (pH 7.0), 10 mM NaCl, 5 mM DTT, and 0.1 U of DPPI. Assay reactions were
initiated by the addition of 1–10 µL of Co(II)-PDF enzyme (final concentration of 0.5–
2.5 nM) and monitored continuously at 360 nm with a Perkin–Elmer Lambda 20 UV–Vis
spectrophotometer. The initial rates were obtained from the early part of the reaction
progression curves (<60 s). To insure that the deformylation reaction is the rate-limiting,
reactions at the lowest and highest substrate concentrations were repeated with doubled
amount of DPPI (0.2 U). The amount of background hydrolysis of f-Met-Lys-AMC or f-
Met-Gln-AMC by DPPI was determined under similar conditions but in the absence of
PDF. This background hydrolysis rate was subtracted from the observed reaction rates.
To determine the catalytic activity of DPPI toward Met-Lys-AMC, a stock solution of f-Met-Lys-AMC (10 mM) in 50 mM Hepes (pH 7.0), 150 mM NaCl was incubated overnight with excess Co(II)-PDF to allow complete hydrolysis of the N- formyl moiety. The PDF was inactivated by heating at 95 ºC for 30 min and removed by centrifugation. DPPI assays were performed by adding 0.005U of DPPI to a mixture (200
µL) containing 50 mM Hepes (pH 7.0), 10 mM NaCl, 5 mM DTT, and 0.5–100 µM of
Met-Lys-AMC. The reaction progress was monitored at 360 nm. The catalytic activity of
DPPI against Met-Gln-AMC was determined in a similar manner.
20 2.5.3 PDF Inhibition Assay
PDF inhibition assays were performed in a reaction mixture (200 µL)
containing100 µM f-Met-Lys-AMC, 0–200 nM inhibitor, 50 mM Hepes (pH 7.0), 10 mM
NaCl, 5 mM DTT, and 0.1 U of DPPI. The background hydrolysis rate was measured by
monitoring the reaction for 30 s at 360 nm on a spectrophotometer. The PDF reaction was
then initiated by the addition of Co(II)-PDF (final concentration 4.0 nM) and monitored
continuously for another 60–120 s. Initial rate was calculated from the early region of the
reaction progress curve (0–30 s after addition of PDF) and corrected by subtracting the
background rate. The inhibition constant (KI) was calculated according to the equation:
V = (Vmax x [S])/(KM(1 + [I]/KI) + [S]).
PDF assay in the fluorescence mode was similarly performed except that the reaction was monitored on an Aminco-Bowman Series 2 Luminescence spectrometer. The excitation and emission wavelengths were set at 380 and 460 nm, respectively. Activity and inhibition assays with Fe(II)-PDF were carried out in the same manner as for Co2+-PDF, except that the reaction buffer contained 1mM tris(carboxyethyl) phosphine instead of 5 mM DTT as the reducing agent (40).
2.6 Results and Discussion
2.6.1 Assay Design
An ideal assay should involve a highly efficient substrate for both PDF and the coupling enzyme, and the PDF inhibitors should not significantly inhibit the coupling enzyme. DPPI meets both of these requirements. First, since DPPI is a cysteine protease and has an entirely different catalytic mechanism from PDF (74), we reasoned that the 21 metal-chelating PDF inhibitors should not inhibit DPPI. Second, DPPI efficiently
hydrolyzes a wide variety of dipeptidyl AMC substrates to release the corresponding
dipeptides and the chromophore/fluorophore AMC (75). Substrate specificity studies of
PDF have revealed a strong preference for an L-methionine at the N-terminal position (7,
69, 76). It has broad specificity at the penultimate position, although amino acids with positively charged side chains are slightly preferred (76). On the basis of the above considerations, we designed a dipeptide substrate, f-Met-Lys-AMC (Fig. 2.3), for the new
DPPI coupled assay. The presence of a polar side chain at the penultimate position also improves the aqueous solubility of the substrate. Sequential action by PDF and DPPI should release formate, dipeptide Met-Lys, and the AMC, which can be monitored by either its absorbance at 360 nm or fluorescence emission at 460 nm. Consistent with a previous report (75), the hydrolysis of Met-Lys-AMC by DPPI is extremely fast, with a
-1 6 -1 -1 kcat value of 5.7 s , KM of 2.8 µM, and a kcat/KM of 2.1 x 10 M s at pH 7.0. The low KM value makes DPPI a particularly suitable coupling enzyme.
2.6.2 Continuous DPPI Coupled Assay
Fig. 2.3A shows the time courses for the hydrolysis of f-Met-Lys-AMC (50 µM) by various amounts of Co2+-PDF and DPPI, monitored at 360nm on a UV–Vis
spectrophotometer. In the presence of 0.03U of DPPI and 5.0 nM of PDF, the substrate is
rapidly hydrolyzed and the absorbance increased linearly with time (tracing A). The
initial rate for f-Met-Lys-AMC hydrolysis approximately doubles when the amount of
PDF is doubled (compare tracings A and B). However, when the amount of DPPI is
doubled while keeping the amount of PDF constant, there is no significant increase in
22 hydrolysis rate (compare tracings A and C). Further, there is no visible lag phase at the
early regions of the curves. These results indicate that under the assay conditions, the
PDF reaction is rate limiting. In the absence of PDF, we also observed a small yet clearly
detectable background hydrolysis of the substrate (tracing D). To determine the origin of
this activity, we treated DPPI with p-hydroxy-α-bromoacetophenone (77), which
selectively inhibits cysteine proteases (78) by alkylating their active-site cysteines, prior
to assay with Met-Lys-AMC and f-Met-Lys-AMC. Similar reduction of activity toward
both substrates was observed. This suggests that the background signal was due to direct
hydrolysis of f-Met-Lys-AMC by DPPI (with f-Met-Lys and AMC as reaction products).
-1 Further kinetic analysis (in the absence of PDF) revealed a kcat value of 0.06 s , KM value
-1 -1 of 82 µM, and a kcat/KM value of 760 M s for DPPI hydrolysis of f-Met-Lys-AMC. In
an attempt to minimize the amount of background hydrolysis, we chemically synthesized
another substrate, f-Met-Gln-AMC, but found that the commercial DPPI preparation also
cleaved this substrate with similar catalytic efficiency, providing further evidence that
DPPI is responsible for the background signal. Fortunately, this background reaction can
be easily measured and corrected, and therefore does not complicate the assay in any
substantial manner. To correct for this background rate, we typically monitored the reaction for 30 s prior to the addition of PDF (Fig. 2.4A); the slope of the line gives the background hydrolysis rate. After the addition of PDF, the reaction is monitored for another 60 s to obtain the total hydrolysis rate. Subtraction of the background rate from the latter gives the PDF reaction rate. The total amount of substrate consumed during the entire 90 s assay period is well below <20%. The slightly increased slope of tracing C
23 relative to A reflects the increased amount of background hydrolysis of f-Met-Lys-AMC
as a result of doubling the amount of DPPI.
f-Met-Lys-AMC is an excellent substrate of PDF and exhibits Michaelis–Menten
2+ -1 kinetics (Fig. 2.4B). For Co -PDF, the catalytic constants are kcat of 26 ± 4 s , KM of 40
5 -1 -1 5 - ± 7 µM, and kcat/KM of (6.5 ± 1.1) x 10 M s at pH 7.0. A kcat/KM value of 7.5 x 10 M
1s-1 was obtained for the same substrate by the formate dehydrogenase assay in the end-
point format. The catalytic constants toward Fe(II)-PDF under the same conditions were
similarly determined as 52 ± 1 s-1, 39 ± 2 µM, and (1.3 ± 0.1) x 106 M-1s-1 , respectively.
These activities are only slightly lower than that of f-ML-pNA, the best PDF substrate
6 -1 -1 previously reported (kcat/KM = 1.9 x 10 M s ) (71). Coupled with the large extinction
coefficient of AMC (ε = 17, 000 M-1cm-1), the current method is highly sensitive.
Actually, the sensitivity of this assay is further improved by monitoring the reaction in
the fluorescence mode. This is critical for assaying very potent PDF inhibitors, because a
proper assay condition requires that the enzyme concentration be lower than that of the inhibitor, which should in turn be similar to the KI value.
2.6.3 Application to Screening of PDF Inhibitors
The utility of the new assay in PDF inhibitor screening was first demonstrated
with BB-3497 (Fig. 2.2, compound 2), a competitive inhibitor of PDF (IC50 ~7 nM against Ni2+-EcPDF) (67). Varying concentrations of BB-3497 were directly added into
the assay reaction and the reaction progress was monitored continuously at 360 nm on a
spectrophotometer or at 460nm on a spectrofluorimeter (excitation at 380 nm). Figure 2.5
shows an example of the reaction progress curves obtained with Co2+-PDF, as monitored
24 on a spectrofluorimeter. The fluorescence yield increased linearly with time and the
initial rates were calculated from the slopes of the lines. Data fitting to the Michaelis–
2+ Menten equation gave a KI value of 23 ± 5 nM against Co -PDF. A KI value of 9.3 ± 1.9
nM was similarly determined for Fe2+-PDF. These values are in reasonable agreement
with the reported IC50 value (~7 nM) (67). Using the same assay method, compound 3
2+ (Fig. 2.2) gave a KI value of 17 ± 2 nM against Co -PDF. Assays in the absorption mode
2+ gave very similar KI values for these two inhibitors. Previous AAP assays with Co -PDF
(in the end-point format) gave KI values of 11 ± 1 and 18 ± 1 nM for compounds 2 and 3,
respectively (79). We have also used this new assay method to successfully determine the
KI values for a variety of inhibitors against E. coli and Plasmodium falciparum. Under the
assay conditions, none of the PDF inhibitors showed any inhibition of DPPI (assayed
with Met-Lys-AMC as substrate).
2.7 Conclusion
In summary, we have developed yet another highly sensitive, convenient, and rapid assay for peptide deformylase. Compared to the previously reported assays, a major advantage of the current method is that the assay reaction can be carried out in a continuous fashion in the presence of PDF inhibitors, making it particularly useful for screening PDF inhibitors in a high-throughput setting and the kinetic characterization of
PDF inhibitors. This feature has allowed us to characterize the slow-binding properties of macrocyclic PDF inhibitor 1, compound 1b in Chapter 5, which had been difficult to accomplish by the other methods. It should be noted that, because PDF inhibitors usually do not inhibit the formate dehydrogenase either, the FDH assay (Method A, (38, 70)) has
25 been widely used for PDF inhibitor screening. However, due to the poor specific activity
of FDH, the FDH assays were typically performed in an end-point fashion. As such, the
FDH assay is not suitable (at least not convenient) for the kinetic evaluation of slow-
binding inhibitors. Its lower sensitivity and end-point nature also make the FDH assay
less reliable for determining the KI values of exceptionally potent PDF inhibitors. A
drawback of the current assay is the presence of slight background hydrolysis of N-
formylated substrates by DPPI. This is not a problem for routine kinetic assays or
inhibitor screening involving the wild-type PDF. However, this background reaction may
become significant and thus complicate the assay results involving catalytically impaired
PDF mutants.
Considering the strengths and weaknesses of each of the PDF assays, we make the
following recommendations. For routine kinetic assays of wild-type and mutant PDF in
the absence of PDF inhibitors, the AAP assay (71) is most convenient. Both FDH and
DPPI assays may be employed to screen PDF inhibitors. If kinetic characterization of a
PDF inhibitor is desired or if the PDF inhibitor is exceptionally potent (low KI value), the
DPPI assay is the method of choice. Further, for applications that require high substrate
concentrations, we prefer the DPPI assay in the absorbance mode, as high concentration
can lead to internal fluorescence quenching and other complications. However, for
determining the KI values of potent PDF inhibitors, the fluorescence mode is
advantageous, as it is more sensitive allowing the use of very low enzyme (e.g., 2 nM) and substrate concentrations.
26
Method A:
S S O PDF FDH NH-peptide NH-peptide HCO H CO H O H N H2N 2 2 2 H O O NAD+ NADH λ = 344 nm
Method B:
Method C:
NO O S H O 2 NO2 S H O H2O N PDF N NO2 H N N H N H H O H H 2 N N O O H SH NH3 O N Fast O H NH2 HCO2H NH2 NH Fast 2 O2N S S NO2
HO2C CO2H
HS NO O2N S S 2 CO H HO2C 2 -1 -1 ε 412 nm = 14,000 M cm
Figure 2.1. PDF Assays. Method A: FDH-coupled; Method B: AAP-coupled; and Method C: Direct-spectrophotometic.
27
Figure 2.2. Structures of PDF inhibitors.
28
Figure 2.3. DPPI-coupled PDF Assay (Method D).
29
Figure 2.4. (A) Time courses for the hydrolysis of f-Met-Lys-AMC by PDF. In a final volume of 200 µL (pH 7.0), f-Met-Lys-AMC (80 µM) was incubated with varying amounts of Co2+-PDF and DPPI. Tracing A, 2 nM PDF and 0.03U DPPI; tracing B, 4 nM PDF and 0.03U DPPI; tracing C, 2 nM PDF and 0.06U DPPI; and tracing D, 0.03U DPPI (no PDF). (B) Plot of initial rates against f-Met-Lys-AMC concentration. The curve was fitted to the data according to the equation, V = kcat x [E] x [S]/ (KM+ [S]), to give a kcat of -1 26 ± 4 s and KM of 40 ± 7 µM. (Data presented are the mean ±SD for three independent experiments.)
30
Figure 2.5. Inhibition of PDF by BB-3497. Reactions were carried out at pH 7.0 using f- Met-Lys-AMC as substrate (40 µM) and in the presence of indicated amounts of inhibitor. The reaction was initiated by the addition of 4 nM Co2+-PDF as the final component and the fluorescence yield at 460 nm (excitation at 380 nm) was monitored with time (no correction of background rate). Inset, plot of remaining activity (after correction for background hydrolysis and relative to that in the absence of inhibitor) against inhibitor concentration. Data fitting against the equation, V = kcat x [E] x [S]/ (KM + [S]), gave a KI value of 23 ± 5nM.
31
CHAPTER 3
CHARACTERIZATION OF A EUKARYOTIC PEPTIDE DEFORMYLASE
FROM PLASMODIUM FALCIPARUM
3.1 Introduction
Early evidence suggested that peptide deformylase (PDF) only existed in the
prokaryotes and apparently absent from the eukaryotic organisms. In this work, we have
taken this notion into consideration and examine the possibility of an evolutionary
conserved deformylation in the eukaryotic organelles. Recent BLAST searches have
retrieved several PDF orthologues from genomes of plants (e.g., Arabidopsis thaliana,
and rice), and parasites (e.g. plasmodium falciparum and Trypanosome brucei) (18). All
of these eukaryotic PDFs carry an N-terminal ~60-amino acid presequence, show
significant sequence homology to EcPDF, and more importantly, contain all of the key
residues for catalysis (Fig. 3.1). To determine whether these genes encode functional
PDF or merely an evolutionary remnant, we have cloned the PDF orthologue gene from
Plasmodium falciparum (PfPDF). The N-terminal presequence of PfPDF was truncated
and the protein was overexpressed in E. coli and purified.
In this chapter, the biochemical characterization of the N-terminal truncated
PfPDF protein will be reported. Our interests include whether the PfPDF is expressed
32 inside the P. falciparum, where the protein is localized inside the cell, and if PDF is
active, whether we could develop PfPDF as a potential antimalarial drug target.
3.2 Experimental Techniques
3.2.1 Cloning of Plasmodium falciparum PDF cDNA.
A BLAST search was performed against P. falciparum 3D7 unfinished genomic
sequence using EcPDF (11) as the query sequence at http://www.ncbi.nlm.nih.gov/
Malaria/plasmodiumbl.html. Several unique shotgun fragments derived from chromosome 9 were identified, which, after translation into peptide sequences, showed extensive homology to the EcPDF. These fragments were subsequently used to search for other fragments at the 5’ and 3’ ends of the P. falciparum def gene (Pfdef). A total of seven unique fragments were obtained and aligned to generate the full-length Pfdef containing a putative 199-nt intron sequence. The intron sequence was identified by its high A-, T-rich sequence (>90%), and the typical sequences at the intron-exon boundary in P. falciparum. Based on the predicted cDNA sequence, two polymerase chain reaction
(PCR) primers were designed: 5’-
GGGGATCCATATGTTGATGTATTATTCACTTTYC-3’ (Y = 50%
T + 50% C) and 5’-GGGAATTCACTCGAGGGCTGGTTCTTCTGAGTGA-
3’. Reverse transcription was carried out with P. falciparum-mixed stage mRNA as template and with MMLV reverse transcriptase as specified by the supplier (Stratagene).
Approximately100 ng of the resulting cDNA was used in a PCR reaction containing 800
µM dNTPs, 0.2 mM of each primer, and 2.5 U of PfuTurbo DNA polymerase
(Stratagene). Thirty cycles were performed as follows: 95°C for 30 s/55°C for 30 s/68°C
33 for 2.5 min. The 0.75-kb PCR product was purified on Qiaquick columns (Qiagen) to
remove any free nucleotides, digested with restriction endonucleases NdeI and XhoI,
and cloned into prokaryotic expression vector pET-29b (Novagen, WI). This cloning
procedure resulted in the addition of a six-histidine tag to the C-terminus of PfPDF.
Earlier work from this laboratory has shown that addition of a C-terminal histidine tag
does not affect the catalytic properties of EcPDF, but permits complete separation of
a tagged mutant from the endogenous EcPDF on a metal affinity column (42). The entire coding region of Pfdef cDNA was sequenced and found to be identical to the genomic sequence (minus the intron). N-Terminal truncation of PfPDF was carried out by PCR using the above 3’ primer and the following 5’ primers: 5’-
GGGGATCCATATGTCAAAAAATTATAGTAGTAATATAA-
3’, 5’-GGGGATCCATATGTCAAGAAAAGGCTCTTTATATTTAT-
3’, 5’-GGGGATCCATATGTCAAAAAATGAAAAGGATGAGATAAAA-
3’, and 5’-GGGGATCCATATGTCAGAGATAAAAATCGTCAAGTACCC-
3’. These primers resulted in the truncation of 40, 50, 57, and 63 amino acids from the
N-terminus of PfPDF, respectively (Fig. 3.1). The PCR products were cloned into plasmid pET-29b as described above. To produce PfPDF without the histidine tag, a synthetic DNA fragment, 5’-TCGAGTAAGGCCT-3’/3’-CATTCCGGAAGCT-5’, was inserted into the XhoI site of above pET-29b-PfPDF constructs. This insert generates an in frame stop codon before the histidine tag.
34 3.2.2 Nucleic acid isolation and blot analyses
Genomic DNA and RNA were isolated from P. falciparum cells as described
(80). Southern and RNA blots were hybridized with random primed 32P-labeled probes
according to standard methods (81).
3.2.3 Protein purification
To prepare Co2+-substituted, C-terminally histidine-tagged PfPDF, E. coli
BL21(DE3) cells carrying the proper plasmid were grown in minimal media containing
60 mg/ml kanamycin at 37°C until OD600 reached ~0.9. The culture was supplemented with 100 µM each CoCl2 and isopropyl-β-D-thiogalactopyranoside and incubated at 30°C
for 18 h. The cells (3 liters) were harvested by centrifugation and resuspended in 100 ml
of buffer 1 containing 100 µg/ml phenylmethanesulfonyl fluoride, 20 µg/ml trypsin inhibitor, and 100 µg/ml lysozyme. The mixture was incubated for 30 min at
4 °C and sonicated with 5 x 10 s pulses. The crude lysate was centrifuged to yield a clear supernatant, which was loaded onto an SP-Sepharose column (3 x 12 cm) which had been preequilibrated in buffer 2. The column was washed with 80 ml of buffer 2 and then eluted with 200 ml of buffer 2 plus a gradient of 10–1000 mM NaCl. Fractions containing significant PDF activity were pooled and concentrated to ~5 ml using
Centriprep-10 (Amicon). After dilution into 50 ml of buffer 3, the protein was loaded onto a Talon cobalt affinity column (Clontech, 3 x 7 cm). After washing with 200 ml of buffer 3, the bound protein was eluted with 50 ml of buffer 3 containing 60 mM imidazole. PfPDF fractions were pooled, dialyzed against buffer 4 plus 100 µM Co2+, and
then buffer 4 plus 1.5 µM Co2+, quickly frozen, and stored at -80°C. Fe2+-PfPDF was
prepared in a similar manner, except that 100 µM (NH4)2Fe(SO4)2 was added to the
35 minimal medium at the time of induction and all of the buffers used were degassed and
added 10 µg/ml catalase before use. Ni2+-PfPDF was prepared according to a literature
procedure (40). Cells were grown in minimal medium supplemented with 100 µM
2+ (NH4)2Fe(SO4)2 and the cell pellet was lysed in 100 ml of buffer 5 containing 5 µM Ni .
The crude lysate was allowed to stand at 4°C for 65 h. The resulting Ni2+-PfPDF was fractionated on the SP-Sepharose column as described above, and the combined PDF fractions were mixed with an equal volume of 100 mM Tris–HCl buffer (pH 8.0) and loaded onto a Ni2+ affinity column (Novagen, 3 x 5 cm). The column was washed with 10
volumes of buffer 6 and 6 vol of buffer 6 plus 32 mM imidazole, and eluted with 6 vol of
buffer 6 plus 1 M imidazole. The PDF fractions were pooled and brought to 80%
saturation with ammonium sulfate. The precipitate was dissolved in buffer 2 and stored
frozen at -80°C. PfPDF without the C-terminal histidine tag was purified similarly
except for the following modifications. After the SP-Sepharose column, PDF fractions
were brought to 35% saturation with ammonium sulfate. The supernatant was loaded on a
phenyl-Sepharose column (3 x 20 cm). The column was washed with 200 ml of buffer 2 containing 35% (w/v) ammonium sulfate and then eluted with 200 mL of buffer 2 with a gradient of 35–0% ammonium sulfate.
Protein concentrations were determined by Bradford assay using bovine serum albumin as the standard and corrected by a factor of 0.71 (actual conc. = 0.71 x conc. from Bradford assay). This correction factor was determined by careful comparison of the metal contents in several Co2+-PfPDF samples and the Bradford results. Metal
analysis was performed for each sample as well as the corresponding buffer solution
36 (control) by inductively coupled plasma emission spectrometry (ICP-ES) at the Chemical
Analysis Laboratory of the University of Georgia.
3.2.4 PDF Assays
Two different assay methods were employed in this work. Method A was used for all other N-formyl substrates. A typical reaction (total volume of 500 ml) contained
50 mM Mops, pH 7.0, 10 mM NaCl, 1 mM TCEP, and 0–5 mM N-formyl peptide. The reaction was initiated by the addition of 0–10 µg of PfPDF and allowed to proceed for 10 min at room temperature before being quenched by heating at 90°C for 10 min (the inactivation process is usually complete within the first 30 s). After cooling to room temperature, the amount of released formate was quantitated as previously described (38,
70). Method B used f-ML-pNA as substrate (71). A typical reaction (total volume of 1.0 ml) contained 50 mM Mops, pH 7.0, 10 mM NaCl, 1.0 mM Tris(2- carboxyethyl)phosphine (TCEP), 0–200 mM f-ML-pNA, and 2 units of AAP. The reaction was initiated by the addition of 0–10 µg of PfPDF and the reaction progress was monitored at 405 nm on a UV-VIS spectrophotometer. The initial reaction rate was calculated from the early, linear region of the progress curve (usually 0–30 s). Inhibition assays (Method B) were carried out with 50 µM f-ML-pNA as substrate, 0–200 µM inhibitor, and 2.4 µg Ni-PfPDF. The reaction was quenched after 5 min by the addition of
50 ml of 10% TFA and neutralized by the addition of 20 ml of 1 M NaOH and then 100 ml of 500 mM potassium phosphate buffer (pH 7.0). After the addition of AAP (1.0 unit) and incubation for 20 min, the absorbance increase at 405 nm was measured. For pH profile analysis, assay reactions (total volume of 500 µl) contained 50 mM buffer of
37 various pH values, 10 mM NaCl, 100 mM f-ML-pNA, and 16 µg of Co-PfPDF. The
buffers used were NaOAc for pH 4.0 –5.5, Mes for pH 6.0–6.5, Mops for pH 7.0, Tricine
for pH 7.5–8.0, and Bicine for pH 8.5–9.0. Reaction products were quantitated in a
manner similar to that of inhibition assays. For all end-point assay reactions, the substrate
to product conversion was kept at <20%.
3.2.5 Western blot Analysis
Partially purified protein extracts (0–40% ammonium sulfate precipitation of the
S-100 fraction) of different stages of parasite growth were analyzed on a 10% SDS-
PAGE, and then transferred to Hybond-ECL membrane (Amersham). The membrane was then probed with affinity purified anti-PfPDF rabbit polyclonal antibody (Cocalico, PA)
and signals were detected using ECLSuperSignal West Pico chemiluminescent substrate
(Pierce).
3.3 Results
3.3.1 Cloning, expression, and purification of recombinant PfPDF
Using EcPDF amino acid sequence as a query, we conducted a BLAST search
(82) of the genomic DNA sequence database stored at the NCBI website
(http://www.ncbi.nlm.nih.gov/). In addition to dozens of putative def genes from various bacteria, the search revealed several eukaryotic genomic DNA fragments that encode proteins with significant sequence homology to EcPDF. A 1.0-kb genomic DNA fragment on chromosome 9 of P. falciparum was found to encode a putative PfPDF protein (241 amino acid residues) plus a 199-nt intron sequence. Southern analysis
38 showed that Pfdef is localized on chromosome 9 as a single copy (data not shown).
Sequence alignment shows that PfPDF contains a catalytic domain, which shares 33% identity to EcPDF, plus a 65-amino acid extension at the N-terminus (Fig. 3.1). The extreme N-terminal region of PfPDF has the characteristic of a classic “von Heijne” signal peptide cleavage site between residues 20 and 21 (83). The full-length cDNA was cloned by a reverse transcription-polymerase chain reaction (RT-PCR) from a mixed stage mRNA library and inserted into prokaryotic expression vector pET-29b. In anticipation that it may be difficult to express the full-length PfPDF in E. coli, we constructed various truncation mutants that lack the N-terminal 40 (∆N40), 50 (∆N50),
57 (∆N57), and 63 amino acids (∆N63). Indeed, E. coli cells harboring the full-length and
∆N40 genes failed to produce any soluble PfPDF protein, whereas ∆N50, ∆N57, and
∆N63 mutants were produced as active enzymes at high levels. Giglione et al. also reported that the full-length A. thaliana PDF could not be expressed in the active form in
E. coli, but a variant lacking the N-terminal extension was produced in a soluble, active form (36). To facilitate purification by metal affinity chromatography, a 7-residue histidine tag (EHHHHHH) was added to the C-terminus of PfPDF.
When grown in Luria broth, E. coli cells harboring the plasmid-encoded Pfdef gene exhibited only low levels of PDF activity, which was rapidly lost during purification. This is reminiscent of the difficulty encountered during the earlier attempts to purify PDF from bacterial lysates (7, 84, 85). It is now clear that the instability of native PDF is due to oxidative damage of its catalytic Fe2+ ion by molecular oxygen and/or trace amounts of H2O2 in the solutions (40, 41). This may suggest that the native
PfPDF, like EcPDF, is an iron enzyme. It was previously shown that substitution of Co2+
39 or Ni2+ for the ferrous ion in bacterial PDF gives highly active and stable enzymes (40,
42, 86). We therefore produced Co2+-substituted PfPDF by growing the cells in minimal
2+ medium supplemented with 100 µM CoCl2 (18). Fe -PfPDF was similarly prepared by
supplementing the minimal medium with 100 µM Fe2+. Ni2+-PfPDF was prepared from
Fe2+-PfPDF by in vitro metal substitution (40).
A number of PfPDF variants that contain different N-terminal truncations (∆N50,
∆N57, or ∆N63), C-terminal structures (with or without the histidine tag), and metal ions
(Fe2+, Ni2+, or Co2+) were purified to apparent homogeneity (Table 3.1 and Fig. 3.2).
Thus, Ni-PfPDF(∆N63)-HT is referred to as the PfPDF protein that has its N-terminal 63
amino acids removed, contains Ni2+ as the metal cofactor, and has a C-terminal histidine tag. Metal analysis indicated approximately 1.0 g-atom Co2+ per polypeptide for the Co-
PfPDF samples. The Ni-PfPDF samples contained 0.47–0.60 g-atom Ni and small
amounts of Zn (~0.1 g-atom). The substoichiometric Ni content is likely due to the low
2+ affinity of Ni to the protein (KD ~ 12 µM) (data not shown) and thus partial loss of
metal during purification. The Fe-PDF samples contained only a small amount of Fe and
Zn; the majority of the metal ion was lost during purification or the handling steps prior
to metal analysis. This is not surprising, since bacterial PDF has previously been shown
to lose activity at a half life of ~1 min, due to air oxidation of the Fe2+.
3.3.2 Kinetic Properties of PfPDF
The catalytic activities of the PfPDF variants were assessed with f-ML-pNA as substrate (71). The length of N-terminal truncation has little effect on activity [compare
Co-PfPDF(∆N63)-HT, Co-PfPDF(∆N57)-HT, and Co-PfPDF(∆N50)-HT in Table
40 2.1] indicating that the N-terminal 63 amino acids are not part of the catalytic domain.
Among the enzymes with different metals, Ni-PfPDF showed the highest activity, with a
4 -1 -1 kcat/KM value of 1.4 x 10 M s . This activity was further increased by ~2-fold when 100
µM Ni2+ was added to the assay buffer, consistent with the above observation that a
significant fraction of the Ni-PfPDF sample was in the apoenzyme form. Fe-PfPDF and
-1 -1 Co-PfPDF had kcat/KM values of 7200 and ~4000 M s , respectively. These activities are
more than two orders of magnitude lower than those of bacterial PDF (38, 42). The lower activity is at least partially due to the poorer binding affinity of PfPDF to the substrate.
While both E. coli and Bacillus subtilis PDFs have KM values of ~20 µM toward f-ML-
pNA (42, 87), PfPDF showed no sign of saturation up to 200 µM substrate (its solubility
limit). We believe that the lower activity of Fe-PfPDF (as compared to Ni-
PfPDF) is due to the low Fe content in the sample (Table 3.1). Simple addition of Fe2+
into the assay buffer did not increase the enzymatic activity, consistent with our earlier
observation with EcPDF (41). Note that the enzymes containing the C-terminal histidine
tag have slightly lower activity than their untagged counterparts [e.g., compare Co-
PfPDF(∆N57) and Co-PfPDF(∆N57)-HT]. The simplest explanation is that the untagged
enzymes are contaminated by small amounts of EcPDF, which has much higher activity
toward f-ML-pNA, whereas EcPDF is completely removed from the histidine-tagged
enzymes by the metal affinity column. We have previously shown that addition of a C-
terminal histidine tag has no effect on the catalytic activity of EcPDF (42).
To determine whether the poor activity of PfPDF toward f-ML-pNA is due to the
different substrate specificity of PfPDF, Ni-PfPDF(∆N63)-HT was assayed with several
other peptides (Table 3.2). For hexapeptides f-MLAFBR (B = β-alanine), f-MAFNBR,
41 -1 -1 and f-MYAFBR, the kcat/KM values generally fall in the range of 2000–4000 M s . Its
-1 -1 kcat/KM value toward f-MA is 55 M s . These values are again two orders of magnitude lower than the corresponding ones for EcPDF (38, 69). Like f-ML-pNA, these substrates
do not show saturation over the concentration ranges tested (up to 1 mM). These results
demonstrate that the cloned Pfdef cDNA indeed encodes a functional PDF and the
eukaryotic PDF is substantially less active than bacterial PDF.
The effect of pH on PfPDF activity was determined with f-ML-pNA as substrate.
Over the pH range tested (pH 4–9), the enzyme undergoes a single ionization event with
a pKa of ~5 and has a flat profile above pH 6 (Fig. 3.3). The reduced activity at low pH
was not due to enzyme denaturation, as treatment of the enzyme at pH 4.0 for the
duration of PDF reactions followed by neutralization to pH 7.0 did not change the
enzyme activity. This pH profile is very similar to that of EcPDF, in which the single
ionization event (pKa ~5.2) has been assigned to the deprotonation of the metal-bound
water/Glu-133 network (42).
3.3.3 Spectroscopic Properties of PfPDF
The electronic absorption spectrum of Co-PfPDF is similar to and yet
distinguishable from that of Co-EcPDF (Fig. 3.4). It has an intense ligand-to-metal
charge transfer band at ~330 nm (ε = 860 M-1cm-1), indicating that the conserved cysteine in the EGCLS motif is ligated to the metal ion. In the visible region, PDF exhibits three d–d transition bands at 572, 634, and 665 nm. The extinction coefficient of 430 M-1cm-1
for the 572 nm band suggests that the cobalt ion is four-coordinate, presumably in the
tetrahedral geometry as found in EcPDF (42). A distinctive feature of the PfPDF is that it
42 has stronger absorption at 634 nm than 665 nm, whereas the trend is reversed in EcPDF.
Circular dichroism of PfPDF shows a single minimum at 222 nm (not shown), as
opposed to the double minima at 210 and 225 nm of EcPDF (39). The spectral
differences indicate that there are subtle differences in their active site structures, which
cause the observed large difference in catalytic efficiency.
3.3.4 Inhibition of PfPDF Activity and Malaria Cell Growth.
N-[(α-mercaptomethyl)caproyl]-L-lysyl-p-nitroanilide (compound 1, Fig. 3.5)(88)
is a potent competitive inhibitor of bacterial PDF (e.g., KI = 19 nM for EcPDF) and
abroad-spectrum antibacterial agent (87). This compound is an apparent competitive
inhibitor of PfPDF, with an IC50 value of 57 µM when tested with f-ML-pNA as
substrate. Binding of inhibitor 1 to Co-PfPDF resulted in both red shift and increase in
intensity of the d–d bands, and a dramatic increase in the ligand-to-metal charge transfer
band intensity (data not shown), as previously observed for EcPDF (42, 87). Actinonin
(compound 2, Fig. 3.5), a naturally occurring PDF inhibitor (KI = 0.3 nM for EcPDF)
(65), also inhibited PfPDF with an IC50 value of 2.5 µM.
Compound 1 was next tested for its ability to inhibit the growth of P. falciparum
in an erythrocyte culture system. In the presence of reducing agent tris(carboxyethyl)
phosphine (TCEP), which is necessary to keep compound 1 in the reduced form (active
form), compound 1 inhibited the growth of P. falciparum with an IC50 value of 65 µM
(Fig. 3.6). In the absence of TCEP, compound 1 was much less potent, presumably
because it was partially oxidized into the inactive disulfide dimer form by air. TCEP by itself had no effect on cell growth (data not shown). The inhibitor was added to the
43 intraerythrocytic malaria culture at the ring development stage. In the presence of the
active inhibitor, the malaria parasite did not mature beyond the ring stage, whereas the control cultures progressed into the schizont stage. Wiesner et al. have recently reported that actinonin inhibits the growth of both wild-type P. falciparum (IC50 = 3.0 µM) and a
multidrug-resistant strain (IC50 = 3.6 µM) (89).
3.3.5 Expression and Localization of PfPDF
RNA blot analysis revealed a steady level of PfPDF transcript of 1.0 kb in ring, trophozoites and early schizonts (Fig. 3.7B). The transcript level decreased progressively as schizonts matured into segmenters. Ethidium bromide (EtBr) staining of the large and
small ribosomal RNAs showed equal loading of samples for all parasite stages
(Fig. 3.7C). Western blot analyses were then performed with malaria extracts from
different stages of intraerythrocytic growth, using rabbit anti-PfPDF polyclonal
antibodies that had been affinity purified against the recombinant PfPDF. Three protein
bands were observed, with a major band at ~24 kDa and two minor bands at ~29 and ~32
kDa, respectively (Fig. 3.7D). None of the three bands appeared in a control experiment
conducted with preimmune rabbit serum (not shown). Note that the size of the 24-kDa
band is very similar to that of the recombinant PfPDF lacking the N-terminal 63 residues
(Fig. 3.2). We assign this band to the mature form of PfPDF that has lost its N-terminal
signal/transit sequence. We are less certain about the identity of the two minor bands,
since their signals were not significantly above background. However, their molecular
masses (relative to the mature form) are consistent with the preprocessed and partially
44 processed PfPDF forms (e.g., cleavage between residues 20 and 21, the putative signal
peptide cleavage site). Interestingly, the peak of PfPDF protein expression was detected
predominantly in late trophozoites and schizonts. A low level of expression was also
present in rings and early trophozoites. Preliminary immunofluorescence experiments
with polyclonal antibodies and the parasite in the segmenter stage indicated localization of PfPDF at foci distinct from nuclear or mitochondrial stains (data not shown). This is
most consistent with localization of PfPDF in the apicoplast (see Discussion).
3.4 Discussion
We have cloned, overexpressed, and purified a eukaryotic PDF from the malarial- causing Plasmodium faciparum organism. Indeed, the recombinant protein is active toward N-formylmethionyl peptides, shares many of the properties of bacterial PDF (e.g., instability and pH profile), and is inhibited by EcPDF inhibitors. Since PfPDF is orders of magnitude less active than EcPDF, there is a possibility that the observed PDF activity could be due to a small amount of EcPDF contaminant. However, several lines of evidence argue against this notion. First, PfPDF exhibits some unique kinetic properties such as much higher KM value toward f-ML-pNA (>200 µM for PfPDF vs 20 µM for
EcPDF). Second, the observed PDF activity is much less sensitive to inhibition by
actinonin or inhibitor 1. Finally, we have previously shown that after a cobalt affinity
column, the amount of endogenous EcPDF in a C-terminally histidine-tagged EcPDF
mutant was less than 0.2 ppm (42). Northern and Western blot analyses demonstrate that
the Pfdef gene is expressed during the parasite life cycle (Fig. 3.7). Our data therefore
firmly establish the presence of a functional PDF in P. falciparum.
45 Although sequence alignment shows 33% identity, plus a 65-amino acid extension at the N-terminus (Fig 3.1), we could not explain the large differences between the kinetic and inhibition activity of PfPDF and EcPDF. Why is PfPDF less active?
Here we collaborated with Dr. Win Hol’s group to obtain a crystal structure of Co2+-
PfPDF at a resolution of 2.8 Å with ten subunits per asymmetric unit (90). The overall
PfPDF structure is superimposable with EcPDF, with the residues involved in catalysis, the position of the bound metal ion, and a catalytically important water molecule structurally conserved between the two enzymes (Fig 3.8) (90). Although both structures may seem to be similar on the surface, however, several key differences exist between them in their active sites. We have modeled the PfPDF protein complexed with the product peptide Met-Ala-Ser (MAS) and compared this to the previously solved structure of EcPDF/MAS complex (44). First, the charge density around the PfPDF active site is more positive than EcPDF (Fig. 3.9A and B) (90). Second, the conserved residue of
Ile105 in PfPDF lies closer to the active site cavity than the equivalent Ile44 in EcPDF, making a ridge on the active site floor and decreasing the cavity volume. Third, the side chain of Arg97 in EcPDF acts as a lid over the active site region, helping to bind substrates and inhibitors (90). The Arg97-Glu42 pair in EcPDF is replaced by the
Glu161-Lys103 pair in PfPDF. Although this change preserves the overall electrostatic charge lining the active site cavity, it leads to important consequences with respect to binding EcPDF inhibitors.
To bring things into perspective, a possible reason behind the decrease in the binding affinity of compound 1 for PfPDF, an IC50 value of 57 µM, where the KI
value of 19 nM for EcPDF, lies in the interaction between the protein S2’ and the lysyl
46 side chain of the inhibitor P2’ position. Whereas the lysyl side chain has a favorable
interaction with Glu42 in EcPDF, it is in close proximity to Lys103 in the model of the
complex of compound 1 with PfPDF, generating unfavorable interactions. Furthermore
based on the key structural differences observed between the EcPDF and PfPDF, it is
possible to develop effective inhibitors to singly target either the eukaryotic or
prokaryotic PDF. For instance, replacing the lysyl side chain of inhibitor
1 by a negatively charged side chain of similar length is likely to remove the repulsive
interactions between 1 and the malarial enzyme (Fig 3.9C).
Where is PfPDF located in the cell? Although encoded by chromosomal DNA,
PfPDF is unlikely to function in the cytoplasm, since there are no N-formylated
polypeptides in the eukaryotic cytoplasm. Therefore, unless PfPDF has another unknown
function, one would expect it to be targeted to an organelle(s) where ribosomal protein
synthesis takes place. P. falciparum contains two such organelles: the mitochondrion and
the recently discovered plastid remnant apicoplast (91, 92). Several lines of evidence are
suggestive of apicoplast localization. First, PfPDF has a 65-amino acid N-terminal
extension (Fig. 3.1), which is not essential for catalysis. The hydrophobic nature of the
first 20 or so residues suggests that this is a signal sequence. Nuclear encoded proteins
that are targeted to the apicoplast are routed through the secretory pathways (93). These proteins often possess a bipartite N-terminal extension containing both a signal peptide and a transit peptide for trafficking across the four apicoplast membranes (94). The N- terminal extension in PfPDF is also bipartite in nature and, like other transit peptides found in P. falciparum (94), the putative PfPDF transit peptide is not rich in serine or threonine. Second, analysis of its subcellular localization with indirect
47 immunofluorescence indicated cytoplasmic localization with foci that are distinct from
either mitochondrial or nuclear stains. Third, while this work was being completed,
Meinnel and co-workers reported the identification of two PDF homologues in A.
thaliana, PDF1A and PDF1B (36). By fusing green fluorescent protein (GFP) to their N- terminal extensions, these investigators have shown that PDF1A is mitochondrial localized, whereas PDF1B is found in both mitochondria and plastids. There is a notable sequence similarity between the N-terminal extensions of PfPDF and PDF1B (36). Taken together, these data strongly suggest that PfPDF is localized in the apicoplast.
Finally, eukaryotic organelles are believed to have evolved from an ancestral endosymbiotic bacterium and share many of the features of bacterial protein synthesis
including the translational initiation with N-formylmethionine. From our germinal
perspective, PfPDF may be involved in the conserved N-terminal methionine excision
(NME) pathway in the apicoplast of P. falciparum, which contribute to the N-end
rule, i.e. the mechanism that links protein half-life to the nature of its N-terminal amino
acid (95). As a consequence, PfPDF is required for MAP action. Here, we and others
(89) observed the disruption of the P. falciparum intraerythrocytic cell growth by the
treatment of PDF inhibitor 1 (Fig 3.6). Together, this observation and the biochemical
results suggest that PfPDF is a potential new target for antimalarial drug design.
48
Metal content (g-atom)
-1 -1 Enzyme Fe Ni Co Zn kcat/KM(M s )
Fe-PfPDF(∆N63)-HT 0.16 0.00 0.00 0.09 7,200 ± 420
Ni-PfPDF(∆N63)-HT 0.00 0.60 0.00 0.06 13,700 ± 1,000
Co-PfPDF(∆N63) 0.00 0.00 0.98 0.02 6,100 ± 400
Co-PfPDF(∆N63)-HT ND ND ND ND 3,100 ± 300
Co-PfPDF(∆N57) 0.00 0.00 0.98 0.01 6,000 ± 850
Co-PfPDF(∆N57)-HT ND ND ND ND 4,000 ± 500
Ni-PfPDF(∆N57)-HT 0.00 0.47 0.00 0.10 14,300 ± 140
Co-PfPDF(∆N50)-HT ND ND ND ND 4,100 ± 60
Table 3.1. Comparison of PfPDF Variants. All activities were measured using f-ML- pNA as substrate. ND, not determined.
49
-1 -1 Substrate kcat/KM (M s )
f-ML-pNA 13,700 ± 1,000 f-MA 55 ± 3 f-MAFNBR 3,400 ± 180 f-MYAFBR 1,900 ± 80 f-MLAFBR 2,800 ± 260
Table 3.2. Kinetic Constants of Ni-PfPDF(∆N63)-HT toward Various Peptide Substrates. Data reported represent the mean ±SD for a minimum of three sets of experiments. B, β-alanine.
50
Figure 3.1. Comparison of the deduced PfPDF amino acid sequence with other PDF sequences in the National Center for Biotechnology Information database using the MegAlign program of the lasergene navigator (DNAstar). Pf, P. falciparum; Syn, Synechocystis (sp:P73441); Ec, E. coli (sp:P27251); and At1 and At2, A. thaliana (AF250959 and AF269165).
51
Figure 3.2. Coomassie blue-stained SDS–PAGE gel (15%) showing purified PfPDF variants. Lane 1, Co-PfPDF(∆N63)-HT; lanes 2, Co-PfPDF(∆N57)-HT, lane 3, Co- PfPDF(∆N50)-HT; lane 4, molecular weight markers.
52
Figure 3.3. Effect of pH on Co2+-PfPDF(∆N63)-HT activity. The reported activities are relative to the activity at pH 6.5.
53
Figure 3.4. Comparison of the electronic absorption spectra of Co-PfPDF(∆N57)-HT (140 µM) and Co-EcPDF (110 µM) at pH 7.5.
54
Figure 3.5. Structures of PDF Inhibitors.
55
Figure 3.6. Inhibition of P. falciparum cell growth in erythrocyte culture by PDF inhibitor 1. DNA synthesis (measured by the incorporation of [3H]hypoxanthine) was used to monitor parasite growth.
56
Figure 3.7. Expression of PfPDF during intraerythrocytic malaria life cycle. Total RNA and proteins were extracted from a highly synchronized P. falciparum culture, every 6 h as follows: ring stage (lane 1), young and mature trophozoites (lanes 2 and 3), schizonts (lanes 4 and 5), and mature schizonts/segmenters (lane 6). Ring, 1–18 h after invasion; trophozoite, 18–28 h after invasion characterized by extensive RNA and protein synthesis and accumulation of hemozoin; schizonts, 28–38 h after invasion when parasites are undergoing multiple rounds of DNA replication and nuclear division; segmenter, individual merozoites are formed by budding of nuclei and cytoplasmic organelles. (A) Morphology of P. falciparum. (B) Northern blot analysis of the levels of PfPDF transcripts at various stages of malaria life cycle. Approximately 10 µg of total RNA from each time point was loaded and [32P]-labeled Pfdef cDNA was used as the probe. (C) Ethidium bromide (EtBr) stained gel showing the equal loading of RNA for each parasite stage. The bands shown represent the large and small ribosomal RNA’s. (D) Western blot analysis using rabbit polyclonal anti-PfPDF antibodies.
57
EcPDF PfPDF
Figure 3.8. E. coli and P. falciparum PDF structural comparison.
58
A B
C
Figure 3.9. Comparison of the Active Sites in E. coli PDF (A) and P. falciparum (B). The surface electrostatic potential rendition is generated by the program GRASP (96). The blue regions denote positive surface potential, whereas the red regions denote negative surface potential. The E. coli structure is complexed with the product peptide Met-Ala-Ser (MAS; Protein Data Bank code 1BS8) (44), whereas the same peptide structure has been modeled in the P. falciparum protein by superposition of the enzymes. (C) Structural model complex of PfPDF with inhibitor 1. These images were imported from the reference (90).
59
CHAPTER 4
CHARACTERIZATION OF A HUMAN PEPTIDE DEFORMYLASE
4.1 Introduction
In chapter 2, a eukaryotic peptide deformylase was cloned from Plasmodium
Falciparum (PfPDF) and demonstrated to be catalytically active (59) . The PfPDF
protein was shown to be expressed in P. falciparum, where growth was inhibited with
potent PDF inhibitor. It is conceivable that PfPDF may play a role in the deformylation
of the apicoplast protein synthesis. Deformylation has been demonstrated to occur in the
chloroplasts of plants (33-35). Additionally, Meinnel and co-workers have shown that
the two def-like genes in Arabidopsis thaliana indeed code for functional PDFs (36), one mitochondria-targeted PDF (PDF1A) and one chloroplast- and mitochondria-targeted
PDF (PDF1B) (15, 36). It was later shown through genetic complementation and actinonin-growth inhibition that, in both Arabidopsis thaliana and Chlamydomonas reinhardtii, the PDF activity is essential for photosynthetic function and the chloroplast
PDF (PDF1B) is a specific target of the antibiotic actinonin in vivo (97) .
Despite the preliminary evidence of functional PDFs presented above, the need for deformylation within the mammalian cells is arguably absent. First, the cytoplasmic
60 protein synthesis in eukaryotes does not involve N-formylation, and therefore, there is no need for deformylation (98). Second, available evidence also suggests that there is no deformylation in the mitochondrion of mammals. For example, the intramitochondrial synthesized proteins from bovine and rat typically retain their N-terminal formyl group or have their N-terminal signal sequences removed (reviewed in refs (15, 99, 100)). The presence of PDF-like sequences in the mammalian genome raises several important questions such as the function of PDF in mammals and the suitability of PDF as a novel drug target. To begin to address these questions, we have undertaken the cloning and characterization of human PDF (HsPDF). Biochemical and kinetic analyses show that the recombinant protein is an active PDF, although its activity is considerably lower than that of the bacterial enzyme. In addition, known PDF inhibitors inhibit its catalytic activity in vitro but had little effect on the growth of human cells. The results are consistent with the hypothesis that HsPDF is a mere evolutional remnant of no current function.
4.2 Experimental Techniques
4.2.1 Cloning, Expression, and Purification of HsPDF
On the basis of the reported HsPDF genomic DNA sequence at http://www.ncbi.hlm.nih.gov, two polymerase chain reaction (PCR) primers were designed as follows: 5'-
GGGGATCCATATGGCCCGGCTGTGGGGCGCGCTGAGTCTT-3' and 5'-
GGGAATTCTTAGTCATTCACCTTCATCCAATAG-3'. Approximately 0.1 µg of a human fetus Marathon-Ready cDNA library (Clontech, CA) was used in the PCR reaction. The Clontech Advantage-GC PCR mixture was employed to destabilize any
61 secondary structures in G,C-rich DNA (101). The PCR reaction (50 µL total volume)
contained 800 µM dNTPs, 0.2 M of each primer, 1.0 M GC melt mixture, and 1 µL of
50× Advantage-GC 2 polymerase mix. The PCR was performed for 32 cycles with the
following conditions: 94 ºC for 30 s/52 ºC for 60 s/68 ºC for 70 s. The 0.73 kb PCR
product was purified on a Qiaquick column (Qiagen) to remove any free nucleotides,
digested with restriction endonucleases NdeI and XhoI, and cloned into the prokaryotic
expression vector pET-22b (Novagen, WI) to produce plasmid pET22b-HsPDF. The
entire coding region of HsPDF was sequenced and found to be identical to the published
sequence.
N-terminal truncation of HsPDF was carried out by PCR using the above 3'
primer and the following 5' primers: 5'-
GGCCCATGGAACGGCGCTCCTATTGGCGCCA-3', 5'-
GGCCCATGGAACCTGAGGCGTCTGGTGCTGG-3', and 5'-
GGCCCATGGAACCTCCCGAACCGCCGTTCTCG-3'. These primers resulted in the
truncation of 44, 52, and 58 amino acids, respectively, from the N-terminus of HsPDF.
The PCR products were cloned into the plasmid pET-42b to generate plasmids pET42b-
HsPDF44, pET42b-HsPDF52, and pET42b-HsPDF58, respectively. This cloning
procedure resulted in the in-frame addition of an N-terminal glutathione-S-transferase
(GST) tag along with a Factor Xa cleavage site.
E. coli BL21 (DE3) Rosetta cells (Novagen) carrying the proper plasmid were
grown in minimal media containing 60 µg/mL kanamycin and 35 µg/mL
chloramphenicol at 37 ºC until OD600 reached ~0.9. The culture was supplemented with
100 µM CoCl2 and 100 µM isopropyl-β-D-thiogalactopyranoside and incubated at 30 ºC
62 for an additional 3 h. The cells (12 L) were chilled on ice for 1 h and harvested by
centrifugation. The cell pellet was suspended in 200 mL of buffer 8 plus 50 µg/mL phenylmethanesulfonyl fluoride, 0.5% protamine sulfate, 20 µg/mL trypsin inhibitor, and
100 µg/mL lysozyme. The mixture was stirred at 4 ºC for 30 min and briefly sonicated (5
× 10 s pulses). The crude lysate was centrifuged to yield a clear supernatant, which was
mixed with 10 mL of GST bind resin (Pharmacia). The column was washed with 300 mL
of buffer 13 and eluted with 50 mL of buffer 13 containing 0.8 mM reduced glutathione.
The GST-HsPDF fractions were pooled and concentrated to ~2 mL using an Amicon
YM-3 cellulose membrane filter. The resulting solution was passed through a Pharmacia
FPLC Fast-desalting column (eluted with buffer 10) to remove the gluthathione and salts.
Fractions containing GST-HsPDF were pooled (~3 mL) and treated with 30 units of
Factor Xa (Novagen) at 4 ºC for 8 h to cleave the GST tag. HsPDF was purified by
passing through a monoQ HR 5/5 anion-exchange column equilibrated in buffer 10. The
column was eluted with buffer 10 plus a linear gradient of 10-1000 mM NaCl. Fractions
containing HsPDF were pooled and concentrated in an Amicon YM-3 cellulose filter.
Protein concentration was determined by Bradford assay using bovine serum albumin as
the standard. Typically, ~1 mg of pure HsPDF was obtained from a 12-L culture. Metal
analysis was performed for the GST-HsPDF fusion protein by inductively coupled
plasma emission spectroscopy (ICP-ES) at the Chemical Analysis Laboratory of the
University of Georgia.
63 4.2.2 Site-Directed Mutagenesis
Mutation of Glu-173 to leucine in HsPDF was performed by the Quick-Change mutagenesis method using the following primers: 5'-CCGAGGGCTG-
CGCTAGCGTCGCCGGCT-3' and 5'-AGCCGGCGACGCTAGCGCAGCCCTCGG-3.
The DNA amplification reaction contained 800 µM dNTPs, 0.1 µg of the plasmid pET42b-HsPDF58 DNA, 0.2 M of each primer, 1.0 M GC-melt mixture, and 2.5 units of
PfuTurbo DNA polymerase (Stratagene). Twenty cycles were performed as follows: 95
ºC for 30 s/54 ºC for 60 s/72 ºC for 14 min. The identity of the mutants was confirmed by
DNA sequencing. The mutant was expressed and purified in the same manner as the wild-type enzyme. Mutation of Leu-91 of EcPDF to a glutamate was similarly carried out with the following primer pair: 5'-
GAAGAAGGTTGCGAGTCGATCCCTGAACAACGTG-3' and 5'-
CACGTTGTTCAGGGATCGACTCGCAACCTTCTTC-3', except that the plasmid pET22b-EcPDF (38) was used as the template. Expression and purification of L91E
EcPDF were carried out as previously described (42).
4.2.3 Peptide Synthesis
All peptides were prepared by solid-phase synthesis on Rink resin as previously described (38). HPLC analysis showed generally >85% purity. The identity of the peptides was confirmed by matrix-assisted laser desorption ionization mass spectrometry.
64 4.2.4 Synthesis of PDF Inhibitors
PDF inhibitors 1-4 were synthesized by Dr. Xubo Hu from commercially (88)
available starting materials.
4.2.5 PDF Assays
Two different methods were employed to assay for PDF activity. Method A was
used for all other N-formylated substrates. It couples the PDF reaction with formate
dehydrogenase, which oxidizes formate into carbon dioxide while reducing NAD+ to
NADH (38, 70). A typical reaction (total volume of 500 µL) contained buffer 13, 1 mM
TCEP, and 0-2 mM N-formylated peptide. The reaction was initiated by the addition of
1-10 µg of HsPDF, and allowed to proceed for 30 min at room temperature before being
quenched by heating at 95 ºC for 10 min (the inactivation process is usually complete
within the first 30 s). After cooling to room temperature, the amount of released formate
was quantified as previously described (38, 70). Method B employed f-ML-pNA as
substrate which, upon deformylation by PDF, is further processed by Aeromonas
aminopeptidase (AAP) to release p-nitroaniline (71). The assay reaction (total volume of
1.0 mL) typically contained buffer 12, 1 mM tris(2-carboxyethyl)phosphine (TCEP), 0-
200 µM f-ML-pNA, and 2 units of AAP. The reaction was initiated by the addition of 1-
10 µg of HsPDF, and the reaction progress was monitored at 405 nm on a UV-vis
spectrophotometer. The initial reaction rate was calculated from the early, linear region of the progress curve (0-45 s). Inhibition assays (method B) were carried out with 150 µM f-
ML-pNA as substrate, 0-400 µM inhibitor, and 1.1 µg of HsPDF. The reaction was quenched by heating at 95 ºC for 10 min and cooled to room temperature. After the
65 addition of AAP (1.0 unit) and incubation for 15 min, the absorbance increase at 405 nm was measured. For all end-point assay reactions, the substrate to product conversion was kept at <20%.
4.2.6 Fluorescence Microscopy
A PCR was performed with plasmid pET22b-HsPDF as template and primers 5'-
TTGCTCGAGATGGCCCGGCTGTGGGGCGCGC-3' and 5'-
GCCGGATCCTCATTCACCTTCATCCAATA-3'. The resulting full-length HsPDF cDNA was digested with Xho I and BamH I and cloned into a mammalian expression vector pEGFP(N1) (Clontech). This cloning procedure resulted in the in-frame fusion of the enhanced green fluorescence protein (EGFP) at the C-terminus of HsPDF to produce the plasmid pEGFP-HsPDF. Approximately 800,000 human embryonic kidney
(HEK293) cells in 2 mL of Dulbecco's minimum essential medium (DMEM) with high glucose containing 10% fetal bovine serum (FBS) were plated onto a sterile 35 mm plastic plate and incubated at 37 ºC overnight. Transient transfection of pEGFP-HsPDF into HEK cells was performed according to LipofectAMINE 2000 protocol (Life
Technologies). The transfected cells were incubated at 37 ºC in a CO2 incubator for 48 h and were subsequently trypsinized and plated onto glass cover slips. After 24 h, the cells were stained with MitoTracker Red CMXRos (Molecular Probes), and the glass cover slip was removed with the adhered HEK cells. Cells were fixed with 4% paraformaldehyde. Visualization was performed under a Bio-Rad MRC-1024 confocal laser scanning unit equipped with a krypton/argon laser, a photomultiplier tube, and an upright Nikon microscope. Images were taken with a 60× oil objective at an Iris setting of
66 3. The 488 and 568 nm laser line was used separately to acquire images for the GFP and
MitoTracker, respectively. Merged images were generated using the Confocal Assistant
software (Bio-Rad).
4.2.7 Cell Growth Inhibition Assay
To facilitate the monitoring of growth rate using a fluorescence plate reader, HEK
293 cells were transfected with the mammalian expression vector pEGFP-IRESneo
(Clontech) as described above. Cells were grown on a 60 mm polystyrene plate with 4
mL of DMEM + FBS media supplemented with 400 µg/mL G418. To maintain a stable
cell line, the transfected cell line was grown to >95% confluency (typically 3 days),
trypsinized, and subcultured for 8 weeks. To test the effect of PDF inhibitors, cells were plated onto a polyornithine-treated 96-well polystyrene plate in 150 µL of DMEM+FBS
(~20,000 cells per well), and grown overnight. The media was removed and replaced
with fresh DMEM+FBS containing 0-128 µM PDF inhibitor 1. Fluorescence signal was
measured every 8 h over a 3 day period using the FLEXstation (Molecular Devices) at an
excitation wavelength of 485 nm and an emission wavelength of 525 nm at 23 ºC.
4.2.8 DNA Synthesis Inhibition Assay
The cell line used, L5178YD10/R cells, is an acute lymphoblastic leukemia cell
line resistant to L-asparaginase (obtained from ATCC). Cells were maintained in DMEM
with high-glucose-containing 10% FBS, 1% antibiotic/antimycotic and 1% L-glutamine.
Cells in log phase were seeded in a 96-well plate at a dilution of 5 × 105 cell/200 µL in
three wells, which was served as the negative control. Log phase cells at a concentration
67 of 2.5 × 106 cells /ml were then mixed with 3H-thymidine (1 Ci/mL of cell suspension) and added to the wells (200 µL/well). Cells in individual wells were treated with PDF inhibitor 1 or 2 at the specified concentrations (in triplicate) ranging from 50 nM to 150
µM and incubated at 37 ºC for 18 h in a CO2 incubator. At the end of incubation, cells were harvested in a cell harvester (Tomech Harvester), and labeled DNA was captured on filter membranes. Membrane-bound DNA was washed in distilled water and air-dried for
3 h. Radioactivity retained on the membrane was measured in a liquid scintillation counter (Beckman).
4.3 Results
4.3.1 Overexpression and Purification of HsPDF
A BLAST search of the human genomic sequence stored at the NCBI website
(http://www.ncbi.hlm.nih.gov/) with EcPDF sequence as query resulted in a single PDF- like homologue, located on chromosome 16 (GenBank accession number AF23915). On the basis of the predicted protein sequence (36), we cloned the full-length cDNA, which encodes a protein of 243 amino acids, from a human fetal tissue cDNA library by PCR.
Sequence alignment shows that HsPDF contains a catalytic domain, which shares ~30% sequence identity to EcPDF, and a 61-amino acid extension at the N-terminus (Figure
3.1). The N-terminal extension resembles that of P. falciparum PDF (PfPDF) (59). We anticipated that the full-length HsPDF would be difficult to express as a soluble protein in
E. coli; therefore, we constructed various truncation mutants that lack the N-terminal 44
(∆N44), 52 (∆N52), and 58 (∆N58) amino acids. We first cloned the truncation mutants in prokaryotic expression vector pET22b and attempted to overexpress them in E. coli
68 BL21(DE3) cells. Unfortunately, neither the mutants nor the full-length protein could be
expressed at significant levels. After considering the different codon bias between human
and bacterial systems, we next constructed HsPDF as a GST fusion protein and employed
an E. coli Rosetta strain that carries several rare tRNA's normally found in E. coli. The
GST fusion would also facilitate its purification on a glutathione column.
Previous studies have shown that bacterial PDF contains a catalytic Fe2+ ion and
is highly sensitive to molecular oxygen and/or H2O2 in solution (41). Replacing the ferrous ion with Co2+ or Ni2+ results in a stable enzyme of essentially wild-type activity
(40, 42, 86). Therefore, we produced HsPDF in the Co2+-substituted form by growing E. coli cells in minimal medium supplemented with 100 µM CoCl2. The ∆N44 and ∆N52
variants did not produce any detectable protein on SDS-PAGE gels. However, we were
able to express the N58 variant and partially purify it on a glutathione column. The ∆N58
protein thus obtained still contained multiple bands, most of them smaller than the 45-
kDa GST-N58 fusion protein (Figure 4.2, lane 4). We believe that the smaller species are
due to proteolysis of the HsPDF portion of the fusion. Consistent with this notion, when
cells were induced for >6 h, the intensity of the 45-kDa band actually decreased (data not
shown). Metal analysis of the GST-HsPDF fusion revealed that the protein contained
approximately 0.67 mol of Co2+ and 0.33 mol of Zn2+, but no significant amounts of any
other divalent metals. Cleavage of the GST fusion with protease Factor Xa, followed by
anion-exchange chromatography, produced Co2+-HsPDF of apparent homogeneity
(Figure 4.2, lane 6). Using the above procedure, ~1.0 mg of HsPDF was obtained from 12
L of E. coli cells.
69 4.3.2 Catalytic Properties of HsPDF
We first evaluated the catalytic properties of HsPDF against an artificial substrate,
-1 f-ML-pNA. HsPDF is a catalytically active enzyme, with a kcat of 0.17 s , a KM of 27
3 -1 -1 µM, and a kcat/KM of 5.9 × 10 M s (Table 4.1). This activity is comparable to that of
another eukaryotic PDF, PfPDF, but is ~200-fold lower than that of EcPDF (42, 59).
Since HsPDF is located in the human mitochondrion (vide infra), its physiological
substrates, if any, would be the mitochondrially synthesized proteins. The human
mitochondrial DNA encodes 13 proteins (102). We synthesized six N-formylated hexapeptides corresponding to the N-terminal sequences of six of the human mitochondrial proteins (cytochrome c oxidase II and III, NADH dehydrogenase subunit II and V, ATP synthase F0 subunit 8, and cytochrome b) (Table 4.1). HsPDF exhibits low
but detectable activity to three of the peptides (f-MAHAAQ, f-MTHQSH, and f-
-1 -1 MTMHTT), with kcat/KM values in the range of 200-800 M s . These activities are again
20-50-fold lower than those of EcPDF against the same substrates (Table 4.2). Two of the
peptides had very poor solubility and accurate activity measurement was not possible for
either EcPDF or HsPDF (f-MNPLAQ and f-MPQKNT). HsPDF showed no detectable
activity toward the remaining peptide (f-MTPMRK).
A striking difference between HsPDF (also mouse PDF) and bacterial PDF is that
the leucine in the highly conserved EGCLS motif is mutated into a glutamic acid in
mammalian PDF (Glu-173 in HsPDF). The leucyl side chain is involved in hydrophobic
' interactions with the P2 side chain of a substrate (44, 48, 61). To determine whether this
mutation is responsible for the low activity of HsPDF, we mutated Glu-173 of HsPDF
back to leucine and the corresponding leucine (Leu-91) in EcPDF to a glutamic acid.
70 When assayed against the above set of substrates, the E173L mutant HsPDF showed
consistently higher activity than the wild-type enzyme (2-4-fold). Vice versa, mutation of
Leu-91 to Glu resulted in a 100-fold reduction of catalytic activity in EcPDF (Table 4.2).
Thus, mutation of the conserved leucyl residue is at least partially responsible for the low catalytic activity of mammalian PDF.
4.3.3 Intracellular Localization of HsPDF
The function of the N-terminal extension of HsPDF was assessed by several prediction software, including signalP (103) and Predotar
(http://inra.fr/Internet/Produits/Predotar/). Both programs predicted that the N-terminal
sequence contain organelle targeting signals. Also, since the mitochondrion is the only
place where N-formylated proteins are synthesized, one would expect that HsPDF be
localized in the mitochondrion, if its function were to deformylate proteins. To determine
the subcellular localization of HsPDF, we fused the green fluorescence protein to the C- terminus of the full-length HsPDF. The fusion protein was transiently expressed in HEK cells. Confocal laser scanning microscopy revealed that the fusion protein is localized in the mitochondria (Figure 4.3). As a control, we transfected the HEK cells with the expression vector pEGFP(N1) alone, which produces only GFP in the cells. Green fluorescence was observed throughout the cytoplasm (data not shown).
4.3.4 Inhibition of HsPDF
Since PDF is being pursued as a target for antibacterial drug design, an important
issue is whether the inhibitors designed against bacterial PDF will also inhibit HsPDF.
71 BB-3497 (compound 1 in Table 4.3) is a potent inhibitor against bacterial PDF (KI = 7
nM against EcPDF) and an orally available, broad-spectrum antibacterial agent (67). Its
N-formylhydroxylamine moiety binds to the catalytic metal of PDF in a bidentate fashion
(67). To address the issue, BB-3497 and three analogues (compounds 2-4 in Table 4.3)
(88) were tested against both HsPDF and EcPDF. All of the compounds showed potent
inhibition of both enzymes, with KI values in the low to medium nM range (Table 4.3).
Among them, BB-3497 is the most potent, showing competitive inhibition pattern (Figure
4.4) and KI values of 8 and 11 nM (this work) against HsPDF and EcPDF, respectively.
Further, with the exception of compound 2, each of the inhibitors was essentially
equipotent against both enzymes.
4.3.5 Effect of PDF Inhibitors on the Growth of Human Cells
BB-3497 was employed to test the effect of PDF inhibitors on human cells.
Initially, we used a stably transfected HEK cell line that expresses GPF and monitored cell growth by following the fluorescence intensity of the cells. We treated the cells with various concentrations of the inhibitor (up to 128 µM) and monitored cell growth over a period of 3 days. The overall fluorescence intensity increased with time, but no significant difference was observed between the treated versus control cells (no inhibitor)
(Figure 4.5A). Direct cell counting also revealed no difference between treated versus untreated cells. No obvious morphological difference could be observed either. Finally, we examined the effect of BB-3497 and inhibitor 2 on DNA synthesis by monitoring the incorporation of [3H]thymidine using a leukemia cell line. Again, neither inhibitor (up to
150 µM) had any significant effect on the rate of DNA synthesis (Figure 4.5B).
72 4.4 Discussion
The question about the roles of mammalian PDF is of both biological and medical
significance. Since numerous earlier studies have shown that mammalian mitochondrial
proteins do not undergo N-terminal deformylation, what is the physiological function of
PDF? As a practical matter, will PDF inhibitors also inhibit human PDF and have
intolerable toxicity? In this work, we have shown unambiguously that the human PDF orthologue encodes a functional PDF. The fact that we and others were able to clone the
cDNA indicates that transcription of the HsPDF gene took place. Indeed, Meinnel et al.
have reported the detection of HsPDF mRNA in a wide variety of human tissues (36).
Since the plasmid-encoded HsPDF-GFP fusion protein is synthesized and targeted to the
mitochondrion, one can assume that the native HsPDF, encoded by chromosome 16, is
also expressed and targeted to the mitochondrion. The lack of deformylation of
mitochondrial proteins is likely due to one or both of the following factors. First, HsPDF
2+ (at least the N-terminally truncated Co form) is an inefficient catalyst; its kcat/KM value
is orders of magnitude lower than that of a typical bacterial PDF. Another possibility is
that the N-termini of the mitochondrial proteins may not be accessible to PDF due to
folding or being imbedded in the membranes. In this regard, it is notable that several
bacterial proteins are known to retain their N-formyl group (104-106).
What is, then, the function of HsPDF? Our view is that HsPDF is a mere remnant of an ancient function that has no current role in mammalian cells, although we cannot
rule out the possibility of it having a function totally unrelated to protein deformylation.
PDF is an essential activity in bacteria apparently due to the inability of methionine aminopeptidase, another essential activity in all living organisms, to process N-
73 formylated polypeptides (15, 98-100). Given that the removal of the N-terminal methionine only occurs in a fraction of bacterial proteins, PDF is required probably because a small number of essential proteins cannot function properly unless their N- formylmethionine is removed. The mitochondrion is believed to have evolved from an ancestral endosymbiotic bacterium and shares many of the features of bacterial protein synthesis including translational initiation with N-formylmethionine. Presumably, HsPDF was an essential activity in the mitochondrion early on, when the organelle genome encoded a larger number of proteins, some of which required N-terminal processing for activity. As the mitochondrial genome shrank to encode just 13 proteins, all of which are apparently functional without deformylation, the essentiality of HsPDF was lost. The lack of any observable effect on human cells by BB-3497, a potent inhibitor against the purified HsPDF, is consistent with this notion. The acquisition of the L173E mutation in mammalian PDF during evolution, which significantly reduces its catalytic activity, also supports our view. It should be emphasized here that this notion may not apply to other organelles that have much larger genomes (e.g., chloroplasts and apicoplasts). In those organelles, the likelihood of having a protein that requires N-terminal processing is much higher, and PDF is more likely to be an essential activity (36).
There are also several possible explanations for the lack of effect on human cells by PDF inhibitors. One possibility is that HsPDF has no catalytic function, and therefore, its inhibition has no effect on the cell. Other possibilities include the inability of the inhibitor to travel into the mitochondrion and rapid degradation of the inhibitor in human cells. Regardless of the cause, the lack of toxicity to human cells by PDF inhibitors makes PDF an attractive target for designing novel antibiotics.
74 In conclusion, we have demonstrated that the human mitochondrion contains a
PDF. However, it apparently has no functional role due to its poor catalytic activity.
Since PDF inhibitors show little toxicity to human cells, PDF is an exciting new target for designing novel antibiotics.
75
-1 -1 -1 Substrate kcat (s ) KM (µM) kcat/KM (M s ) f-ML-pNA 0.17±0.06 27±6 5900 f-MAHAAQa 0.57±0.08 2500±310 230 f-MTHQSHb 1.5±0.5 1850±230 810 f-MNPLAQc ND ND ND f-MTMHTTd 2.2±0.9 4000±800 545 f-MPQLNTe ND ND ND f-MTPMRKf NA NA NA
*These peptides correspond to the N-terminal sequences of human mitochondrial proteins: a, cytochrome c oxidase II; b, cytochrome c oxidase III; c, NADH dehydrogenase subunit II; d, NADH dehydrogenase subunit V; e, ATP synthase F0 subunit 8; and f, cytochrome b. ND, activity not determined due to insolubility. NA, no detectable activity. All assays were performed in triplicates and data reported represent the mean ± SD.
Table 4.1: HsPDF Activity against f-ML-pNA and Human Mitochondrial Peptides*
76
-1 -1 kcat/KM (M s )
Substrate Wt HsPDF E173L HsPDF Wt EcPDF L91E EcPDF f-ML-pNA 5900±1100 21800±3000 (1.2±0.1)×106 12000±2000 f-MAHAAQ 230±55 810±60 12000±1600 ND f-MTHQSH 810±65 1200±400 15000±1300 ND f-MTMHTT 540±50 1320±450 29000±5800 ND f-MTPMRK NA NA 2390±230 ND
ND, not determined; NA, no detectable activity.
Table 4.2: Catalytic Activity of Wild-Type vs Mutant HsPDF and EcPDF
77
Table 4.3: Inhibition Constants (KI, nM) of EcPDF and HsPDF
78
Figure 4.1. Sequence alignment of various PDF's from eukaryotic and prokaryotic organisms: Plasmodium Falciparum (Pf), Escherichia coli (Ec), Homo sapiens (Hs), Bacillus subtilis (Bs), Staphylococcus aureus (Sa), and Arabidopsis thaliana (At). Alignment was performed by inputting PDF sequences into ClustalW software located at http://clustalw.genome.ad.jp/ website.
79
Figure 4.2. A 15% SDS-PAGE gel showing the purity of HsPDF(∆N58) during different stages of purification. Lane 1, molecular-weight markers; lane 2, crude cell lysate; lane 3, flow-through fraction of the GST-bind column; lane 4, proteins eluted from the GST-bind column; lane 5, after factor Xa cleavage; and lane 6, HsPDF after purification on Mono-Q column. Molecular weight standards (kDa) are indicated on the left side of the gel.
80
Figure 4.3. Intracellular localization of HsPDF-GFP fusion in HEK cells. Fluorescence signal was visualized under a confocal microscope. (A) Fluorescence signal of mitochondria stained by MitoTracker (red). (B) Fluorescence signal of GFP (green). (C) An overlay of panels A and B.
81
Figure 4.4. Lineweaver-Burk plot for the inhibition of HsPDF by BB-3497. Assay reactions (total volume of 500 µL) contained buffer E, 1.1 µg of HsPDF, 0-160 nM inhibitor, and 0-80 µM f-ML-pNA.
82
Figure 4.5. Effect of BB-3497 on human cell growth. (A) Fluorescence cell count. HEK cells that stably express GFP were treated with the indicated concentrations of BB-3497, and the total fluorescence of the culture was measured at various times. (B) [3H]thymidine incorporation. Acute lymphoblastic leukemia cells (L5178YD10/R) were grown in the presence of specified concentrations of BB-3497 (0-150 µM). The amount of radioactivity in the isolated DNA was measured by liquid scintillation counting. RFU, arbitrary fluorescence unit.
83
CHAPTER 5
CHARACTERIZATION OF MACROCYCLE PDF INHIBITORS
5.1 Introduction
In Chapter 4, the characterization of HsPDF as well as work carried out in the
human cell culture study validated PDF as a novel antimicrobial drug target (79). In this
chapter, we will continue the development of a more effective, potent class of PDF
inhibitors. Numerous PDF inhibitors have been reported in recent years; essentially, all of
them are metal chelators such as thiols (87, 107, 108), hydroxamates (64, 65, 109-112),
and N-formylhydroxylamines or reverse hydroxamates (67, 113). Many of these
inhibitors exhibit excellent antibacterial activities in vitro and in animal studies. One of
the reverse hydroxamates from British Biotech, BB86398, has been advanced into
clinical trials for the treatment of hospitalized community-acquired pneumonia. However,
because most of these inhibitors are metal chelators appended to peptidomimetics, there
remains some concern about their selectivity (e.g., inhibition of other metalloproteases)
and in vivo stability (e.g., proteolysis of the peptide bonds). A popular approach to
improving both stability against proteolysis and selectivity of peptidomimetic inhibitors
is to form cyclic peptides or depsipeptides (114, 115).
Structural studies of several PDF-inhibitor complexes (48, 61, 67) have revealed
that the inhibitors are bound in an extended conformation and the P1' and P3' side chains
84 are similarly oriented. While the P2' side chain is extended toward solvent, the P1' and P3' side chains are engaged in intimate interactions with the enzyme. The P1' side chain
(usually an n-butyl group) fits into a deep hydrophobic pocket in the PDF active site. The
P3' side chain makes hydrophobic contacts with a shallow pocket near the active site, as
well as one face of the P1' side chain. Moreover, the rigidity introduced by cyclization
may lock the inhibitor into the PDF-binding conformation and thus improve binding affinity as well as selectivity by preventing binding to other enzymes. On the basis of the structure of reverse hydroxamate 2 (BB-3497) (Fig. 5.1) (67), a potent PDF inhibitor and antibacterial agent, we designed a potent class of PDF inhibitor and antibacterial agent, in which the P1' and P3' side chains are cross-linked together by various long linker of
saturated carbon atoms to form a series of “tripeptide-liked” macrocyclic inhibitors (1a-h,
Fig. 5.1) (88). These cyclic inhibitors showed good antibacterial activity and markedly improved metabolic stability and selectivity.
5.2 Experimental Techniques
5.2.1 PDF Inhibition Assays
Method B (Chapter 2, (71)) was employed to determine the KI* values. PDF
inhibition assays were performed by incubating PDF (8 nM) in 50 mM Hepes, pH 7.0,
150 mM NaCl, 100 µM bovine serum albumin, and 0-300 nM inhibitor for 2 h on ice.
The solution was brought to room temperature in a water bath, and the reaction was
initiated by the addition of f-ML-pNA substrate (80 µM). After 10 min at room
temperature, the reaction was quenched by heating at 95 ºC for 15 min and cooled to
room temperature in a water bath. AAP (1.0 unit) was added, and the resulting mixture
85 was incubated at room temperature for 15 min. The absorbance at 405 nm was measured
in a quartz cuvette on a UV/vis spectrophotometer. The background absorbance was
determined by repeating the reaction but in the absence of PDF and subtracted from the
observed absorbance values. The inhibition constant, KI*, was calculated from the initial
rates using the Michaelis-Menten equation: V = (Vmax × [S])/(KM(1 + [I]/KI*) + [S]). The
percentage of substrate to product conversion was kept at <20%.
Method D was employed to determine the KI and k6 values. Assay reactions (total volume 200 µL) were performed at room temperature in a quartz cuvette containing 5-
150 µM f-Met-Lys-aminomethylcoumarin (f-Met-Lys-AMC) as the substrate, buffer 15, and 0.1 U of DPPI. Prior to use, DPPI was activated by the treatment with 5 mM dithiothreitol in 50 mM Hepes (pH 7.0) and 10 mM NaCl for 30 min. The reactions were
initiated by the addition of 1-10 µL (2-10 ng) of PDF and monitored continuously at 360
nm in a UV/vis spectrophotometer. The initial rates were obtained from the early part of
the reaction progression curves (<30 s). The background signal was measured under the
same conditions but in the absence of PDF and was subtracted from the observed PDF
reaction rates. The PDF inhibition assays were performed in a similar manner except that
the reactions contained 0-300 nM inhibitor and fixed concentrations of PDF (4.0 nM),
DPPI (0.1 U), and f-Met-Lys-AMC (130 µM). The inhibition constant, KI, was calculated from the initial rates using the above Michaelis-Menten equation.
To determine the rate constant, k6, PDF (40-160 nM) and inhibitor (80-200 nM) were preincubated in 100 µL of buffer 15 and 100 µM BSA for 2 h on ice. The mixture was rapidly diluted into 900 µL of buffer 15 containing 270 µM f-Met-Lys-AMC and 0.1
U DPPI, and the reaction was monitored at 360 nm on a UV/vis spectrophotometer. After
86 correction for background hydrolysis by DPPI, the progress curve was fitted to the
-k6t equation: Abs360 = Vs[t - (1 - e )/k6], where Vs is the final steady state velocity. The rate
constant k5 was calculated from KI, KI*, and k6 values by using the equation: KI* =
KIk6/(k5 + k6).
5.2.2 Antimicrobial Susceptibility Testing.
For Escherichia coli and Bacillus subtilis, cell growth was monitored on the
Lamda-20 UV/Vis spectrophotometer at 600 nm. A single bacterial colony from a plate
culture was grown overnight in LB media and was diluted 1000-fold into 2 mL of fresh
growth medium containing 0–64 µg/mL of test agent in a 5-mL glass test tube shaken at a
speed of 250 rpm. Cell growth (at 37 ºC) was monitored for 0–600 min.
5.2.3 Inhibition of MMPs
The effect of PDF inhibitors on human MMP-1, MMP-2, MMP-3, and MMP-9
was assessed in reactions (400 µL total volume) containing buffer 16, 15 µM fluorogenic
MMP peptide substrate, and 0, 1, or 10 µM PDF inhibitor. For MMP-1, MMP-2, and
MMP-3, peptide Mca-P-K-G-Dpa-A-R-NH2 was employed as the substrate. For the
detection of MMP-3 activity, peptide Mca-R-P-K-P-V-E-Nval-W-R-K(Dnp)-NH2 was added to the assay. The assay reactions were initiated by the addition of 0.10 µg of the proper MMP enzyme, and the remaining MMP activity was monitored continuously on an Aminco-Bowman Series 2 luminescence spectrometer (excitation wavelength at 320 nm and emission at 405 nm).
87 5.2.4 Antimicrobial Susceptibility Testing
The antimicrobial susceptibility testing was performed using a microdilution
broth assay based on the guidelines of the National Committee for Clinical Laboratory
Standards (NCCLS) document M7-A5 (NCCLS, 2000). Cation-adjusted Mueller-Hinton broth (CAMHB) was used in the microtiter plates for dilution of the agent stock solution
and for diluting the bacterial inoculum. For Streptococci, 5% lysed horse blood was
added to the CAMHB. Haemophilus Test Medium purchased from REMEL Inc. was
used for Haemophilus species. Inoculum was prepared by taking a sample of bacteria
from a 16-20 h plate culture and adjusting it in sterile saline to an OD600 that corresponds
to ~1 × 108 colony-forming units per milliliter (CFU/mL). The saline suspension was
then diluted to ~1 × 106 CFU/mL in appropriate media to produce the standardized
inoculum. A sample of the inoculating culture is diluted in sterile saline through a series
of six 1:10 dilutions, and all dilutions are plated by spotting 10 µL of each dilution,
incubated overnight at 35-37 ºC, and counted the next day to verify the inoculum size.
Microtiter plates (Evergreen 96 well microplate number 222-8032-01R) were prepared
with the use of the Beckman BiomekR 1000 automated laboratory workstation in
combination with manual techniques. The microtiter plate was filled with 50 µL of
diluent broth using the BiomekR 1000 instrument. The agent or control antibiotic stock
solution was manually added to the drug well of the microtiter plate using a Calibra 852
Pipettor. The agent/antibiotic was serially diluted in 2-fold dilutions using the BiomekR
1000 instrument. Fifty microliters of the standardized bacterial inoculum was added to
each well using the BiomekR 1000 instrument, producing a final inoculum of ~5 × 105
CFU/mL. In addition, a nontreated growth control and commercially available control
88 antibiotics were included to validate the assay. The final concentrations ranged from 0.03
to 32 µg/mL for test agents and 0.06 to 64 µg/mL for control antibiotics. For controls, all
stock and working concentrations were 1 mg/mL. The microtiter plates were incubated
overnight (16-20 h) in a 35-37 ºC incubator with the exception of the Haemophilus and
Streptococcus species, which were incubated at 35-37 ºC and in the presence of 5% CO2 for 24 h. The MIC90 is defined as the lowest concentration of agent that prevented visible
growth of the organism. The end point was determined visually using a plate reading
mirror.
5.2.5 In Vitro Stability of PDF Inhibitors in Rat Plasma
Compounds were diluted 1000-fold into rat plasma (Lot 27624, Pel-Freez,
Rogers, AK) to result in an incubation concentration of 5 µg/mL. All samples were
incubated at 37 ºC for 0-5 h. Aliquots were withdrawn at various time points, the proteins
were precipitated with acetonitrile, and internal standard was added (10 µg/mL). After the
proteins were removed by centrifugation, the supernatant was analyzed for the amount of
remaining parent compound by LC-MS/MS. LC was conducted on an Agilent 1100
Binary HPLC equipped with a C18 column. The chromatography parameters were
optimized so that the analyte (1f and 3) coeluted with the internal standard. MS was
performed on a ThermoFinnigan LCQ-DUO instrument. MS parameters were optimized
for maximal ion signal with the AutoTune routine of the Xcalibur software
(ThermoFinnigan). The samples were prepared in duplicate.
89 5.3 Results and Discussion
To demonstrate the effectiveness of the novel macrocylcic PDF inhibitors,
compound 1a-1g were was assayed against Co(II)-substituted E. coli PDF using Method
* D (Chapter 2, (116). These macrocycles acted as potent inhibitors, with KI values
ranging from 0.22-36 nM (Table 5.1). Thus, cyclization of the P1' and P3' side chains
renders compounds 1b-1d, most potent, more than ~10-fold more potent than the acyclic
parent compound 2 (KI = 11 nM) (Table 5.1). To gain insight into the mechanism of
inhibition, the Co-PDF-inhibitor 1c complex was examined by UV-visible spectroscopy.
Binding of the inhibitor resulted in marked blue shift (by ~40 nm) and reduction in the maximum intensity of the D-D transition bands in the absorption spectrum of the cobalt ion (Figure 5.3). This result suggests that compound 1c is directly ligated to the metal ion. The maximum absorptivity of ~200 M-1 cm-1 for the PDF-inhibitor complex is
consistent with a bidentate interaction between the N-formylhydroxylamine group and the
metal and the formation of a pentacoordinated cobalt (117).
To further provide evidence that the macrocycle binds in the PDF active site, we
have recently collaborated with Dr.Wim Hol group at the University of Washington,
Seattle, to obtain a crystal structural complex of Haemophilus influenzae PDF (HiPDF)
with compound 1d at 1.5Å (Fig. 5.4). This compound potently inhibited Co2+-HiPDF at
* 2+ KI = 0.071 ± 0.007 nM. The Co metal ion (pink) is shown coordinated by side chains
of Gln-51, Cys-91, His-133, His-137, and interacting with the two oxygens of the
hydroxamate moiety of 1d.Residues 43-51, which start as a loop then become a strand
followed by a small helix, form a base for the P2' sidechain with the formyl group on the
2+ far side with the Co . The P2' trimethyl-methyl makes interactions with the CD1 and
90 CD2 carbon atoms of Leu92 (Fig. 5.4). The binding groove surface is formed by the
convergence of three peptides. The side chain of Ile45 is then in a position at the hollow
center of the inhibitor. Together, these favorable interactions provided strong affinity of
1d in the HiPDF active site. We believe that the rest of the other macrocycles bind in a similar manner.
The in vitro antibacterial activity of the cyclic inhibitor was tested against E. coli and Bacillus subtilis, the representative Gram-negative and Gram-positive bacteria, respectively. Figure 5.5 shows the bacterial cell growth curves in the presence of varying
concentrations of inhibitor 1b. Compound 1b exhibited potent antibacterial activity
against B. subtilis, with a minimal inhibitory concentration (MIC) of 2-4 M (or 0.7-1.4
g/mL). It is only moderately active against E. coli, with an MIC of ~32 M (~12 g/mL).
The lower activity against E. coli is likely due to its inefficient permeation of the bacterial outer membrane and/or being removed from the cells by the efflux pump.
5.3.1 Slow-binding Inhibition of PDF
The macrocycles 1a-h were initially tested against PDF by Method A (38, 70) or
Method B (71). Surprisingly, these inhibitors exhibited a slow-binding behavior (118)
against PDF. Coupled with their exceptional potency, a detailed kinetic characterization
of the inhibitors using the above assay methods proved problematic. Here, we employed
our newly developed PDF assay method, Method D (Chapter 2, Fig 2.3 (116)). Because
none of the PDF inhibitors that we have tested showed any inhibition against DPPI (DPPI
is a cysteine protease), the method can be carried out in a continuous fashion and is
ideally suited for kinetic characterization of the slow-binding inhibitors 1a-h. Figure
91 5.2A shows the reaction progress curves in the absence and in the presence of inhibitor
1c. In the absence of inhibitors, the reaction progress curve was essentially a linear line.
Straight lines were also observed in the presence of competitive, acyclic inhibitors 2-4
(not shown). However, the cyclic inhibitors 1a-h each resulted in biphasic curves.
The inhibition kinetics can be described by the equation,
k KI 5 E + I E I E I* k 6
where KI denotes the equilibrium inhibition constant of the initial enzyme-inhibitor
. complex (E I) and k5 and k6 are the forward and reverse rate constants for the slow
conversion of the initial EI complex into a tight-binding complex EI*, respectively. The
overall potency of the inhibitor is described by the equilibrium constant, KI* = KIk6/(k5 +
k6). The initial inhibition constant KI was determined by initiating the assay reactions
(which contained varying concentrations of the inhibitor) by the addition of PDF and
fitting the initial rates against the Michaelis-Menten equation. The KI* values were
determined by preincubating the enzyme and various concentrations of the inhibitor for 2
h prior to the addition of substrate. To distinguish slow-binding inhibition from time-
dependent irreversible inactivation and to determine the rate constant k6, the inhibitors
were preincubated with PDF to form the EI* complex, which was then rapidly diluted
into an assay solution containing the PDF substrate, f-Met-Lys-AMC. The slow, time-
dependent recovery of PDF activity demonstrates that the inhibition is reversible and thus
slow binding in nature (Figure 5.2B). Curve fitting gave the reactivation rate constant
-1 (k6), in this case 0.0037 min , for inhibitor 1c. The rest of the macrocycles behaved in a similar manner (Table 5.1). 92 5.3.2 In Vitro Antibacterial Activity
The cyclic inhibitors were first tested against E. coli and Bacillus subtilis, the
representative Gram-negative and Gram-positive bacteria, respectively. For B. subtilis,
there is a general correlation between the antibacterial activity and the KI* values against
PDF, consistent with PDF as the molecular target responsible for the observed
antibacterial activity. Compounds 1b-d, which are the most active inhibitors against PDF
(Table 5.1), also exhibited the most potent antibacterial effect, with minimal inhibitory
concentrations (MIC90 values) of 0.7-1.4, 0.2-0.4, and 0.08-0.16 µg/mL, respectively
(Table 5.2). On the other hand, inhibitors 1a,e-g had both poorer KI* (3.0-36 nM) and
MIC90 values (1.4 to >32 µg/mL). Most of the compounds were less effective against E.
coli, and there was no correlation between the KI* and the MIC90 values (Tables 5.1 and
5.2). This suggests that other factors such as efflux pumps, membrane permeability,
intracellular distribution, and/or intracellular stability are at play. A notable exception is
inhibitor 1f, which is equally active against E. coli and B. subtilis (MIC90 = 4-8 µg/mL).
Compound 1f contains a lysine as the P2' residue (Figure 5.1). It was previously reported
that substitution of a P2' lysine for a leucine substantially improved the antibacterial
activity of another PDF inhibitor (87). It is not yet clear how the inclusion of an amine
group improves the cellular activity of the inhibitors.
Next, compounds 1b,e,g were selected for testing against a panel of clinically
significant pathogens including Enterococcus faecalis, Haemophilus influenzae,
Moraxella catarrhalis, Staphylococcus aureus, and three Streptococcus pneumoniae
strains. Compound 1b had a good antibacterial activity against most of the pathogens,
with MIC values ranging from 0.5-16 µg/mL (Table 5.2). The high activity of compound
93 1b against H. influenzae strain ATCC 31517 is worth noting. This pathogen is generally
difficult to treat with other antibiotics especially with the other reported PDF inhibitors
(113, 119, 120). To gain some insight into the molecular basis of the observed high efficacy, we tested compound 1b against the purified H. influenzae PDF and found 1b to
be an exceptionally potent inhibitor against the enzyme (KI* = 0.074 ± 0.016 nM). A KI* value of 0.071 ± 0.007 nM was also determined for compound 1d (against H. influenzae
PDF). Compound 1e had a good activity against M. catarrhalis and the S. pneumoniae strains (MIC = 0.031-8 µg/mL) but was poorly active against E. faecalis, H. influenzae, and S. aureus (Table 5.2). Compound 1g was active only against M. catarrhalis, a result not unexpected from its relatively poor KI* value.
5.3.3 Cyclization Improves Inhibitor Selectivity
A major concern of any metal-chelating PDF inhibitor is its potential inhibition of
the numerous metalloproteinases in the host, such as the distantly related matrix
metalloproteases (MMPs). We tested several of the cyclic inhibitors (1a,b,e,g,h) and the
acyclic controls 3 and 4 against a panel of human MMPs (MMP-1, MMP-2, MMP-3, and
MMP-9). At 10 µM, the acyclic compound 3 resulted in essentially complete inhibition
of all four MMPs; even at 1 µM, it caused 50-80% inhibition (Table 5.3). Acyclic
compound 4 had a similar inhibition profile. In contrast, none of the cyclic inhibitors showed greater than 20% inhibition of MMP activity at 10 µM inhibitor concentration
(Table 5.3). It has previously been reported that acyclic compound 2 is highly selective
for PDF, showing minimal inhibition of MMPs, enkephalinase, or ACE (67). The
difference between inhibitors 2 and 3 or 4 is likely due to the presence of a bulky tert-
94 butyl group at the P2' position of 2, which prevents binding to the active sites of these
other enzymes. Remarkably, even the cyclic inhibitors that do not contain the tert-butyl group at the P2' position (e.g., compounds 1g,h) showed little inhibition of the MMPs.
Thus, cyclization per se was sufficient to largely prevent the inhibition of MMPs.
Presumably, cyclization locked the inhibitors into conformations that do not fit the active
sites of these MMPs, while more selective for the PDF active site (Fig. 5.4).
5.3.4 Cyclization Improves the Stability of Inhibitors
Inhibitors 1f and 3 were chosen to test for the effect of cyclization on inhibitor
stability. They both have an L-lysine as the P2' residue and are expected to be sensitive to
proteolytic degradation by trypsin-like enzymes. Both inhibitors were incubated in rat plasma at 37 ºC, and aliquots were withdrawn at various times and analyzed by liquid chromatography-mass spectrometry (LC-MS). The cyclic inhibitor (1f) was very stable under the experimental conditions, showing no detectable degradation after 5 h (Figure
5.6). In contrast, the acyclic compound (3) showed time-dependent degradation, with approximately 25% loss after 5 h. Therefore, cyclization significantly improves the stability of the inhibitors against metabolic degradation. Our attempts to identify the degradation products by MS failed.
5.4 Conclusion
In conclusion, based on our earlier observations that the P1' and P3' side chains of
PDF inhibitors are closely packed in the PDF-inhibitor complex, we have developed a
new class of macrocyclic PDF inhibitor by covalently linking the two side chains. The
95 cyclic inhibitor is highly potent against PDF and has excellent to moderate antibacterial activity against both Gram-positive and Gram-negative bacteria. This result demonstrates that cyclization of the P1' and P3' side chains is a viable approach to developing potent
PDF inhibitors. Due to their more rigid structures, cyclic inhibitors of this type may also have improved stability and selectivity.
The ring size greatly affects the inhibitor properties, with 15-17-membered rings as the optimal ring size. As compared to their acyclic counterparts, the cyclic inhibitors of the optimal ring sizes show a much higher potency against PDF (>20-fold), improved stability against proteolytic degradation, and higher selectivity for PDF over other metalloproteases. To the best of our knowledge, the cyclic inhibitors rank among the most potent inhibitors reported to date against the PDF enzyme. Some of the inhibitors have good antibacterial activity against a wide spectrum of pathogens. Further development of these cyclic scaffolds may provide novel PDF inhibitors with improved metabolic stability and higher selectivity against human MMPs.
96
a b -1 c -1 Inhibitor KI (nM) KI* k5 (min ) k6 (min )
1a 63 ± 12 13.7 ± 3.0 0.022 0.0061 ± 0.0015 1b 109 ± 5 0.33 ± 0.15 1.2 0.0038 ± 0.0010 1c 96 ± 17 0.23 ± 0.06 1.6 0.0037 ± 0.0010 1d 77 ± 20 0.22 ± 0.04 0.50 0.0015 ± 0.0008 1e 2100 ± 500 25 ± 6 1.0 0.012 ± 0.0058 1f 92 ± 20 3.0 ± 1.3 0.12 0.0042 ± 0.0011 1g 258 ± 28 36 ± 9 0.11 0.018 ± 0.0045 1h 710 ± 170 12 ± 4 0.28 0.0048 ± 0.0008 2 11 ± 1) N/Ad N/A N/A 3 74 ± 17 N/A N/A N/A 4 23 ± 6 N/A N/A N/A
a b KI values were determined by the DPPI assay in a continuous fashion. KI* values were c determined by the AAP assay in the end-point format. k6 values were determined by diluting a preformed E.I* complex into a large volume of assay buffer containing f-Met- Lys-AMC and DPPI. dN/A, not applicable (no slow-binding behavior). All values reported represent the mean ± SD from three independent sets of experiments.
Table 5.1. Inhibition Constants against E. coli PDF
97
MIC90
Bacteria 1a 1b 1c 1d 1e 1f 1g
E. coli >24 12 >12 >24 >24 4-8 >32 B. subtilis 1.4-2.8 0.7-1.4 0.2-0.4 0.08-0.16 16 8 >32 E. faecalis (ATCC 29212) NDa >32 ND ND >32 ND 32 H. influenzae (ATCC 31517) ND 0.5 ND ND >32 ND 32 M. catarrhalis (ATCC 37054) ND 0.62 ND ND 0.031 ND 1 S. aureus (ATCC 29213) ND 16 ND ND >32 ND 32 S. pneumoniae (ATCC 49619 ) ND 4 ND ND 8 ND >32 S. pneumoniae (ATCC 6301) ND 4 ND ND 8 ND >32 S. pneumoniae (ATCC 6303) ND 2 ND ND 4 ND >32 a ND, not determined.
Table 5.2. In Vitro Antibacterial Activity of PDF Inhibitors
98
MMP-1 % MMP-2 % MMP-3 % MMP-9 %
1.0 10.0 1.0 10.0 1.0 10.0 1.0 10.0 Inhibitor µM µM µM µM µM µM µM µM
1a 0 1 10 15 3 20 8 11 1b 7 14 4 8 3 9 5 9 1e 7 8 0 0 3 4 4 6 1g 1 7 6 15 0 6 4 5 1h 3 13 17 20 0 8 8 8 3 50 100 73 98 81 99 61 94 4 50 97 34 55 68 73 54 89
aThe values reported are the percentages of MMP activity inhibited in the presence of indicated concentrations of inhibitors. Each inhibitor was tested at two different concentrations (1.0 and 10 µM).
Table 5.3. Inhibition of MMPsa
99
O O R H NH2 N H N N H OH O O O H n N H N N H 1a n = 1 R = -C(CH3)3 OH O 1b 3 -C(CH3)3 1c 4 -C(CH3)3 1d 5 -C(CH3)3 1e 8 -C(CH3)3 3 1f 3 -CH2CH2CH2CH2NH2 1g 8 -CH2CH2CH2CH2NH2 1h 3 -CH2CH2CH2CH2NHAc NH2
O O O O H N N H N N H N N H H OH O OH O
2 4
Figure 5.1. Structures of PDF inhibitors.
100
Figure 5.2. Slow-binding inhibition of PDF by compound 1c. (A) Reaction progress curves when PDF was added as the last component. Each assay reaction contained buffer A, DPPI (0.1 U), f-Met-Lys-AMC (130 µM), the indicated concentration of 1c (0, 50, 100, or 150 nM), and PDF (4.0 nM). (B) Reaction progress curves when substrate was added as the last component. PDF (100 nM) and varying concentrations of 1c were preincubated for 2 h and rapidly diluted (10-fold) into buffer A containing f-Met-Lys- AMC (270 µM) and DPPI (0.1 U). a, control reaction with 1.0 nM PDF (no inhibitor); b, 120 Nm inhibitor; and c, 140 nM inhibitor.
101
------PDF PDF + 1b
Figure 5.3. Electronic absorption spectra of Co(II)-substituted PDF (240 M) in the absence and presence of inhibitor 1b (500 µM).
102
L-92
Q-51 C-91
H-137
H-133
Figure 5.4. Crystal Structure of HiPDF/Macrocycle 1d Complex . The electron density and hydrogen bonding network of 1d and an ordered water molecule (green) involved in a protein/inhibitor contact. The cyclic structure is shown in yellow. Key interactions are explained in text.
103
Figure 5.5. Inhibition of B. subtilis (A) and E. coli (B) cell growth by inhibitor 1b. An overnight culture was diluted 1000-fold into 2 mL of fresh LB medium containing the specified concentrations of inhibitor 1b and incubated at 37 ºC. Cell densities were measured at the specified times on a UV-vis spectrophotometer.
104
120
100
80
60 1f 40 3 % Remaining 20
0 012345 Incubation Time (h)
Figure 5.6. Comparison of the in vitro stability of PDF inhibitors 1f (cyclic) and 3 (acyclic) in rat plasma.
105
CHAPTER 6
PURIFICATION AND CHARACTERIZATION OF ENZYMES INVOLVED IN
THE DEGRADATION OF CHEMOTACTIC N-FORMYL PEPTIDES
6.1 Introduction
Upon microbial infection or tissue damage, a cascade of immunologic response
triggers the accumulation of leukocytes to the site of infection or injury. This process is
initiated by the binding of chemotactic N-formylated peptides to the formyl-peptide
receptor (FPR) located on the cell surface (121, 122). FPR receptors belong to the seven-
transmembrane domain Gi-protein-coupled receptor (GPCR) family. It has been
suggested that the N-formyl group is a crucial determinant for ligand binding to the FPR
(123). Since bacterial (105, 124) and mitochondrial proteins (125) are the only sources of
N-formylated peptides in nature, it has been widely accepted that these FPR receptors mediate the trafficking of phagocytes to sites of bacterial invasion or tissue damage
(126). Among the chemotactic N-formylated peptides thus far discovered, peptide formyl-Met-Leu-Phe (f-MLF) is one of the most potent.
Mammalian large intestine is colonized by commensal bacteria that release bioactive (127) and immunoreactive f-Met peptides, including f-MLF (128); however, it would be detrimental if the N-formyl peptides should cross the intestinal mucosal barrier
106 to provoke inflammatory responses. Indeed, colonic infusion or rectal administration of f-
MLF results in experimental colitis in animals (129, 130). Thus, to prevent unwanted
immune response to commensal bacteria, mammals must possess enzymes that can
effectively degrade the N-formyl peptides. Chadwick and co-workers (131) as well as other investigators (132) have previously reported that rat intestine contains two enzymes that act sequentially to degrade the N-formyl peptides (Figure 6.1). The first enzyme, N- formylmethionine (f-Met) aminopeptidase (fMAP), hydrolytically removes N-
formylmethionine from N-formyl peptides including f-MLF, eliminating their
chemotactic activities. Subsequently, f-Met deformylase (fMDF) further converts f-Met
into formate and methionine (132). It was reported that fMAP is specific for N-
formylmethionyl and N-acylmethionyl peptides and has no activity towards other N-
acylated amino acids (e.g., f-Ala, f-Val, f-Leu, f-Arg, and f-Phe derivatives) (131).
Although the enzyme was purified to near homogeneity, its identity has not been
established. Other investigators have purified enzymes with α-N-acylpeptide hydrolase
(APH or acylamino acid-releasing enzyme, EC 3.4.19.1) activities from sheep (133),
rabbit (134) and human erythrocytes (135), rat liver (136-138), and rabbit skeletal muscle
(139), which are capable of releasing f-Met from N-formylmethionyl peptides. However,
the latter enzymes seem to have different substrate specificity profiles from fMAP (131).
The identity of fMDF also remains to be determined. According to Chadwick and co-
workers (131), fMDF is most active against f-Met and N-formylnorleucine, has reduced
activity against f-Leu, but is inactive against other N-formyl amino acids or N-
formylmethionyl peptides. fMDF activities have also been described in crude extracts of
107 rabbit reticulocyte lysates (134), human leukocytes and platelets (140), and Euglena gracilis (141). Again, in neither case has the identity of the enzyme been established.
We have undertaken the purification and characterization of fMAP and fMDF in order to understand their roles in the degradation of N-formyl peptide chemotactic agents.
Our interest stems from our ongoing work on another enzyme, peptide deformylase
(PDF) from bacteria. PDF is an essential enzyme that removes the N-formyl group from newly synthesized proteins in bacteria and certain eukaryotic organelles (99). PDF is currently being pursued as a novel antibacterial drug target. However, treatment of bacteria cells with PDF inhibitors may result in the accumulation of N-formyl peptides
(142), a condition that could potentially induce unwanted inflammatory responses. In this work, we have purified both fMAP and fMDF to homogeneity and established their identities as APH and N-acylase IA, respectively. Detailed characterization of their substrate specificities suggests that they are responsible for the degradation of N-formyl peptides.
6.2 Experimental Techniques
6.2.1 Purification of fMAP
Fifteen frozen rat small intestines (-80 ˚C) were thawed and washed with ice chilled buffer 18. The loosely bound mucosal layer was removed by gently scraping the inside lining of the small intestines and washed with buffer 18 (3 x 100 mL) and centrifuged at 4˚C for 10 min at 500 g (Sorvall GS-3 rotor). The pellet was resuspended in 30 mL of buffer 18 and sonicated for 5 x 10 s pulses (medium probe, Branson). The suspension was adjusted to 10 mM CaCl2 and stirred at 4 ˚C for 30 min and centrifuged
108 at 4 ˚C for 90 min at 200,000 g (Beckman Ti-70 rotor). The solution (35 mL) was diluted
with ice chilled ddH2O (final volume 150 mL) and loaded onto a Q-Sepharose Fast Flow
(Pharmacia) column (2.5 x 12 cm) pre-equilibrated in buffer 19. The adsorbed proteins
were eluted with 300 mL of buffer 19 plus a linear gradient of 10-500 mM NaCl.
Fractions that showed activity towards f-M-p-NA and f-MLP substrates were pooled
(Figure 6.2A) and concentrated to 37 mL in an Amicon YM-10 nitrocellulose concentrator. The solution was adjusted to 1.7 M ammonium sulfate (final concentration)
and loaded onto a Pharmacia Phenyl-Sepharose column (1.6 x 10 cm) pre-equilibrated in
butter 20. Elution was performed with buffer C and a reverse ammonium sulfate linear
gradient with buffer 21 (1.7-0 M; 300 mL at 2.0 mL/min). Active fractions (Figure 6.2B)
were pooled and loaded onto a Pharmacia FPLC Mono Q column (HR 5/5) pre-
equilibrated in buffer 19. Elution was performed with a linear gradient of 10 mM-1.0 M
NaCl over 20 mL. Active fractions were pooled and precipitated with 85% ammonium
sulfate and centrifuged for 30 min at 4 ˚C (30,000 g). The pellet was dissolved in 500 µL of buffer 22 and passed through a Superdex S-200 column (1.0 x 30 cm) equilibrated in buffer 22 at a flow rate of 0.5 mL/min. The active fractions were pooled and concentrated with Millipore YM-10 filter. The protein concentration was determined by Bradford assay using bovine serum albumin as the protein standard.
6.2.2 Purification of fMDF
Twenty frozen rat small intestines (-80 ˚C) were thawed and washed with buffer
H. The fecal matter was discarded and the mucosal layer was removed from the intestinal lining. The suspension was washed with 200 mL of buffer 26 with centrifugation at 500 g
109 and 4 ˚C (Sorvall GS-3 rotor). The loosely packed pellet was resuspended in 150 mL of
buffer I and sonicated for 5 x 10 s pulses (medium probe, Branson). The reddish
suspension was centrifuged at 27,000 g (Sorvall SS-34 rotor) for 60 min at 4 ˚C. The supernatant was diluted with ice-cold water to a final volume of 200 mL. The pH of the solution was adjusted to 5.5 with ice-chilled (1/5 dilution) glacial acetic acid and centrifugation was continued for 15 min. The solution was loaded onto a CM-Sepharose column (2.5 x 10 cm, Sigma) pre-equilibrated in buffer 27. fMDF activity appeared in the flow-through fractions. The flow-through fractions were loaded onto an SP-Sepharose column (2.5 x 10 cm, Pharmacia) pre-equilibrated in 25 mM Mes, pH 6.0, to remove additional proteins. The pH of the flow-through fractions was readjusted to 8.0 with a 1.5
M Tris (pH 8.0) solution. The mixture was loaded onto a Q-Sepharose column (2.5 cm x
16 cm, Pharmacia) pre-equilibrated in buffer 19. The adsorbed proteins were eluted with
300 mL of buffer B plus a linear gradient of 10-500 mM NaCl. Fractions containing activity against f-Met were pooled and brought to 25% ammonium sulfate and loaded onto an Octyl-Sepharose column (2.5 x 8 cm, Pharmacia) pre-equilibrated in buffer 21.
The strongly adsorbed protein was washed with 200 mL buffer D and eluted with 150 mL
of the same buffer plus a linear gradient of 0-4% Triton X-100. Fractions with fMDF
activity were pooled and loaded onto a Mono-Q column (HR 5/5, Pharmacia) pre-
equilibrated in buffer 19. Elution of the adsorbed protein was effected with 20 mL of
buffer 19 plus a linear gradient of 10 mM-500 mM NaCl. The active fractions were
pooled and passed through a Superdex S-200 gel filtration column (1.0 x 30 cm,
Pharmacia) pre-equilibrated in buffer 22. Protein solution was passed thru a small
Concanavalin A-Sepharose column (1 g). Active fMDF fractions were pooled and
110 concentrated with a Millipore YM-10 nitrocellulose filter and stored at –80 ºC. The protein concentration was determined by Bradford assay using bovine serum albumin as the protein standard.
6.2.3 Synthesis of N-Formyl-methionyl-p-nitroanaline
Fmoc-methionine and p-nitroaniline were thoroughly dried over phosphorus pentoxide. Fmoc-methionine (2.0 mmol) and p-nitroaniline (2.0 mmol) were dissolved in
8 mL of dry pyridine. The mixture was brought to –20 ˚C in a methanol/dry ice bath.
Trichlorophosphorus oxide (3.2 mmol) was slowly added to the chilled solution and the reaction was allowed to proceed for 2 h. The reaction was brought to room temperature and stirred for 3 h. The reaction was neutralized with 50 mL of 5% sodium bicarbonate and extracted with ethyl acetate (3 x 15 mL) and washed with 0.5 M HCl (3 x 30 mL).
The solvent was evaporated under reduced pressure to give 389 mg product (40% yield).
The Fmoc group was removed by dissolving the solid in 5 mL of 20% piperidine in dichloromethane for 1 h at room temperature. The product was recrystallized from methanol. The resulting methionyl-p-nitroanilide was N-formylated by treatment with 3 mL of 96% formic acid and 1 mL of acetic anhydride at 0 ˚C. The reaction product was purified by flash column chromatography (silica gel) to give 100 mg of a white solid. 1H
NMR (250 MHz, CDCl3) δ 9.18 (s, 1H), 8.30 (s, 1H), 8.23-7.68 (m, 4H), 6.53 (d, J = 7.5
Hz, 1H), 4.93 (m, 1H), 2.67 (m, 2H), 2.27 (m, 2H), 2.15 (s, 3H).
111 6.2.4 fMAP Assay
FMAP activity was assayed by two different methods. In method 18, assay reactions were performed in polystyrene cuvettes (1 mL total volume) containing buffer
23 and 0-800 µM f-M-pNA. The reaction was initiated by the addition of 0.56 µg of fMAP and continuously monitored at 405 nm on a Perkin-Elmer Lambda 25 UV/Vis spectrophotometer. Method 19 employs N-formyl peptides (0-20 mM) as substrates.
Reactions (100 µL total volume in buffer 23) were initiated by the addition of 0.56 µg of fMAP and incubated at room temperature for 15 min. The reaction was quenched by heating at 100 ºC for 30 min and centrifuged to remove the precipitated proteins. The solution was diluted to 200 µL in ddH2O and 200 µL of buffer 24 was added. To determine the amount of free amine formed, 200 µL of a fluorescamine solution (0.1 mg/mL in ethanol) was added and the fluorescence yield was measured on an Aminco-
Bowman Series 2 Luminescence spectrometer (excitation wavelength at 390 nm and emission wavelength at 475 nm). The amount of product formed was determined by comparison to a standard line generated with methionine. Inhibition of fMAP was carried out in a total volume of 1 mL containing 150 mM NaCl, 50 mM HEPES, pH 7.0, 0.39 mM f-M-pNA and various concentrations of an effector molecule (e.g., 1,10- phenanthroline). The reaction was initiated by the addition of 0.56 µg of fMAP and the absorbance at 405 nm was monitored for 60 s.
6.2.5 fMDF Assay
Two methods were employed to assay fMDF activity. In method 18, the amount of formate released was quantitated by using formate dehydrogenase (FDH) as the
112 coupling enzyme. The reaction (500 µL total volume) contained buffer 23, 5 mM NAD+,
1.0 unit FDH, and 0-10 mM f-Met as substrate. The reaction was initiated by the addition of fMDF (final concentration 24 nM) and monitored continuously at 344 nm on a
Lambda 25 UV/Vis spectrophotometer. In method 19, assay reactions were performed in
200 µL of buffer 23 containing 0-3.0 mM N-acetyl-L-methionine. The reaction was initiated by the addition of fMDF (final concentration 24 nM) and allowed to proceed for
5 min at room temperature. The reaction was quenched by the addition of 10 µL of trichloroacetic acid and centrifuged at 14,000 rpm in a microcentrifuge for 10 min. The pH of the supernatant was adjusted by the addition of 200 µL of buffer 24. Product formation was analyzed by the addition of 200 µL of 0.1 mg/mL fluorescamine solution and the fluorescence yields were measured immediately as described above.
6.2.6 Gel-Filtration Analysis
Analysis was performed on a Superdex S-200 column (Pharmacia, 10/30 GL) connected to an AKTA/FPLC (Amersham) at flow rate of 0.5 mL/min. The column was pre-equilibrated in buffer 18. Protein samples (200 µL) were injected onto the FPLC column and detection was monitored at 280 nm wavelength. Protein molecular weight standards were ovalbumin (45 kDa), albumin (66 kDa), aldolase (158 kDa), catalase (232 kDa), and ferritin (440 kDa).
6.2.7 In-Gel Digestion
Protein samples were separated on 12% SDS-PAGE gels under reducing conditions. Gels were fixed overnight in a 50:10:40 ethanol/acetic acid/water solution and
113 stained with Coomassie blue G-250. The desired protein band was excised and washed
with the ethanol/acetic acid/water solution for several hours. The gel slice was dried with acetonitrile and treated with a dithiothreitol solution to reduce any disulfides.
Iodoacetamide was added to alkylate the cysteines and the gel was washed with cycles of
acetonitrile and ammonium bicarbonate solution. The resulting gel slice was treated
overnight at room temperature with sequencing grade trypsin (Promega, Madison, WI) by
using the Montage In-Gel Digestion Kit (Millipore, Bedford, MA) and following
manufacturer’s recommended protocols. The peptides were extracted from the
polyacrylamide gel with a 50:5:45 acetonitrile/formic acid/water solution several times,
pooled, and concentrated in vacuo to ~25 µL.
6.2.8 Nano-LC MS/MS
This method was employed to identify tryptic digested fMDF peptides. Capillary-
liquid chromatography-nanospray tandem mass spectrometry (Nano-LC/MS/MS) were
performed on a Micromass hybrid quadrupole time-of-flight Q-Tof(tm) II (Micromass,
Wythenshawe, UK) mass spectrometer equipped with an orthogonal nanospray source
from New Objective, Inc. (Woburn, MA) operated in positive ion mode. The LC system
was a LC-Packings-Dionex Capl LC. The solvent A was water containing 50 mM acetic
acid and the solvent B was acetonitrile. 2.5 µL of each sample was first injected into the
trapping column, and then washed with 50 mM acetic acid. The injector port was
switched to inject and the peptides were eluted off of the trap onto the column. A
ProteoPep C18 column (5-cm x 75-µm) packed directly in the nanospray tip was used for
chromatographic separations. Peptides were eluted directly off the column into the Q-
114 TOF system using a gradient of 2-80%B over 30 min, with a flow rate of 40 µL/min with
a pre-column split to about 500 nL/min. A total run time was 55 min. The nanospray
capillary voltage was set at 3.0 kV and the cone voltage at 55 V. The source temperature
was maintained at 100 0C. Mass spectra were recorded using MassLynx 4.0 with
automatic switching functions. Mass spectra were acquired from mass 400 - 2,000
Daltons every 1 s with a resolution of 8,000 (FWHM). When the desired peak was
detected at a minimum of 15 ion counts, the mass spectrometer automatically switched to acquire CID MS/MS spectrum of the individual peptide. Collision energy was set dependent on charge state recognition properties. Sequence information from the
MS/MS data was processed using Mascot Distiller. Database searches were performed using Mascot and Genomic Solutions.
6.3 Results
6.3.1 Purification of fMAP and fMDF
fMAP was purified in a similar manner as described by Chadwick et al. (Table
6.1) (131). Upon removal of loosely bound mucosal layer from the small intestine, sonication, and centrifugation at 200,000 g to remove organelles and brush-border membrane vesicles, the enzyme activity remained in the supernatant fraction. During purification, fMAP activity was initially monitored by assay method A, which employs f-
M-pNA as substrate and is very sensitive and convenient. Throughout the purification procedure, we were able to detect only a single fraction that is capable of cleaving f-M- pNA to release p-nitroaniline (Figure 6.2). To ascertain that this fraction contained the fMAP activity, all fractions were subsequently assayed against f-MLF (assay method B).
115 The same activity profile was obtained. A combination of anionic exchange (Q-
Sepharose and Mono Q), hydrophobic interaction (Phenyl-Sepharose), and gel-filtration chromatographic steps resulted in a 2,061-fold purification of the enzyme. The purified protein showed one major band at Mr ~55,000 Da on SDS-PAGE gels (Figure 6.3). Gel- filtration analysis of the protein gave a native molecular mass of ~280,000 Da. As reported previously (131), the enzyme did not bind to either wheat germ agglutinin or concanavalin A-Sepharose, indicating that the enzyme is not glycosylated. These results suggest that the native fMAP is a homotetramer.
Purification of fMDF from the small intestinal epithelial mucosal layer also followed a literature procedure (132) but with a number of modifications (Table 6.2).
After cell lysis and centrifugation (200,000 g), fMDF activity remained in the supernatant fraction. The enzyme was stable after acidification of the supernatant to pH 5.5. This greatly facilitated its purification, as many proteins precipitated at this pH and were readily removed by centrifugation. The deformylase was also apparently very hydrophobic, binding exceptionally tight to a Phenyl-Sepharose column. Triton X-100 detergent was required to elute the protein off the column. Overall, fMDF was purified
1833-fold by a combination of acidification, two anion-exchange chromatographic steps, a hydrophobic interaction step, a size exclusion step, and an affinity chromatographic step. The purified enzyme showed a molecular mass of ~45,000 Da in both SDS-PAGE and gel-filtration analyses, suggesting that fMDF exists as a monomer under physiological conditions (Figure 6.4).
116 6.3.2 Identification of fMAP and fMDF by Mass Spectrometer Fingerprinting
Protein bands corresponding to fMAP and fMDF were excised from SDS-PAGE gels, digested to completion with trypsin, and the resulting peptides were analyzed by
LC-ESI mass spectrometry. The obtained peptide masses were then used to search against the NCBI database for potential matches. Such analysis of fMAP resulted in a single match with APH from rat liver with a score of 112. Out of a total of 27 peptide fragments derived from the fMAP sample, 19 peptides matched exactly the calculated masses of the corresponding peptides in APH (Table 6.3). However, rat APH contains
732 amino acid residues and has a calculated molecular mass of 81,347 Da (143), which is significantly larger than the observed fMAP molecular mass of ~55 kDa (Figure 6.3).
We noted that all 19 matched peptides in Table 6.3 are located in the C-terminal fragment of APH, with the first peptide (YCTNR) matching the amino acids 274 to 278 of APH.
This suggests that the purified fMAP is a proteolytic fragment of APH. It has previously been reported that APH contains a 55-kDa C-terminal peptide hydrolase domain and a
22-kDa N-terminal domain of unknown function and that treatment with trypsin was able to separate the two domains with little effect on its peptidase activity or in some cases slightly increased peptidase activities (144, 145). Consistent with our interpretation, the native APH also exists as a homotetramer of 340,000 Da (143).
Analysis of fMDF peptides was performed in a slightly different manner. The peptides derived from a tryptic digest of fMDF were analyzed by electrospray-ionization tandem mass spectrometry (ESI–MS/MS). The individual peptides were fragmented in the second spectrometer by collision-induced dissociation (CID) and the resulting product ions were analyzed with Mascot software. Out of a total of 12 peptides examined, 7
117 matched exactly with peptide fragments from N-acylase IA with a score of 254 (Table
6.4). The next two potential matches were a trypsin precursor and a putative calcium binding protein, with scores of 49 and 36, respectively (data not shown).
6.3.3 Catalytic Properties and Inhibtion of fMAP (APH)
To confirm the identity between fMAP and APH, the purified fMAP was examined for its substrate specificity and sensitivity to inhibitors. fMAP is active toward both N-formyl- and N-acetyl peptides of two to six residues in length, but with the
following trends (Table 6.5). First, fMAP shows a strong preference for a dipeptide
substrate, showing progressively lower activities toward longer peptides (Table 6.5,
compare f-MF vs f-MLF, f-MA vs f-MAS, f-MS vs f-MSS, and Ac-AA vs Ac-AAA).
The increasing KM values with peptide length suggest that fMAP binds dipeptides more
effectively, perhaps by recognizing the α-carboxylate group of the penultimate residue.
Second, the enzyme hydrolyzes N-formyl and N-acetyl peptides with essentially equal
efficiency (compare f-MAS and Ac-MAS). Finally, fMAP has a weak preference for a
smaller amino acid at the N-terminus, showing ~2-fold higher activity toward N-alanyl
peptides than N-methionyl peptides (compare f-MA and Ac-AA, Ac-MAS and Ac-
AAA). Among all of the substrates tested, the artificial substrate, f-M-pNA (which
5 -1 -1 mimics a dipeptide), is most active (kcat/KM = 1.2 x 10 M s ). Importantly, fMAP
-1 efficiently hydrolyzes the chemotactic peptide f-MLF, with a kcat value of 4.6 s , a KM
-1 -1 value of 2.0 mM, and a kcat/KM value of 2300 M s . Thus, the catalytic properties of our
purified fMAP are in qualitative agreement with those reported for APH (136, 146-148).
Quantitative comparison is not possible as the literature values were generally relative
118 activities determined at fixed substrate concentrations. Compared to the catalytic
constants reported for fMAP by Chadwick et al. (131), our values are approximately ~10-
fold lower (Table 6.5). This difference is likely caused by different assay conditions; in
this work, all assays were performed at pH 7.0, whereas Chadwick et al. carried out
assays at pH 8.0 (optimal pH for APH is 8.2). Other factors that may cause the observed
activity difference include partial proteolysis (in our case) and difference in purity.
Among the effector molecules tested, fMAP activity was not affected by the
presence of 2-mercaptoethanol (5 mM), dithiothreitol (10 mM), or 100 µM divalent metal ions (Co2+, Zn2+, and Ni2+). However, the enzyme was potently inhibited by diethyl
pyrocarbonate (DEPC), which modifies histidine and cysteine residues in proteins (IC50 =
35 nM). In addition, fMAP is weakly inhibited by the serine protease inhibitor, phenylmethylsulfonyl fluoride (PMSF), with an IC50 value of 150 µM. These results are
in excellent agreement with those reported for fMAP by Chadwick et al. (131) and with the properties reported for APH (136, 146-148), a member of the serine protease family
(147). Further, both fMAP (131) and APH (136) are markedly inhibited to completely inactivated by Cu2+ and Hg2+ ions. Thus, the comparison of catalytic and inhibition
properties also supports the notion that fMAP and APH are the same enzyme.
6.3.4 Catalytic Properties and Inhibition of N-Acylase IA (fMDF)
fMDF hydrolyzed both f-Met and Ac-Met to release free methionine, with slightly
-1 -1 higher activity toward Ac-Met (kcat/KM of 7600 vs 2600 M s ) (Table 6.6). Chadwick et
al. reported the KM values of 7.1 mM and 0.22 mM for f-Met and Ac-Met, respectively,
at pH 8.0 (132). The corresponding values determined in this work (at pH 7.0) are 3.1
119 mM for f-Met and 0.3 mM for Ac-Met (Table 6.6). Comparison of kcat values was not
possible because they were not reported by Chadwick et al. fMDF showed no activity
against either N-formylated or N-acetylated di- and tripeptides (e.g., f-MA, f-AA, f-
MAS, Ac-MAS, and f-MLF). Among the effector molecules tested, dithiothreitol (20
mM) and iodoacetamide (10 mM) strongly inhibited the enzyme activity (85% and 90%,
respectively). The requirement for divalent metal ions was examined. Treatment with
EDTA (50 mM) and 1,10-phenanthroline (10 mM) also reduced the enzyme activity, to
10% and 6%, respectively. Incubation of the enzyme with 100 µM NiCl2 or CoCl2 increased catalytic activity by 2–3-fold. These results are in excellent agreement with those reported for fMDF by Chadwick et al. (132) and acylase I (149, 150).
6.4 Discussion
Protein synthesis in bacteria, chloroplasts, and mitochondria is initiated with N- formylmethionine residue and consequently, all nascent polypeptides contain an N- terminal f-Met. In bacteria, PDF removes the formyl moiety from the vast majority of nascent polypeptides. However, the deformylation process is not complete and some mature proteins retain their N-terminal f-Met residue (99). In addition, bacterial cells are known to release small N-formylated peptides such as f-MLF into their environment
(105, 132). The fate of the N-formyl moiety of mitochondrially synthesized proteins is currently less well understood. There are strong indications that these mitochondrial proteins retain their N-terminal f-Met residue, due to the poor catalytic activity of mitochondrial PDF (79). Therefore, during tissue damage the mitochondria can also release N-formylated peptides. Since the intestinal mucosal epithelium is constantly
120 exposed to these inflammatory N-formylated peptides, there must exists enzymatic activities that can efficiently degrade the chemotactic peptides. Indeed, Chadwick and others (131, 132, 140) have partially purified and characterized fMAP and fMDF, which act sequentially to inactivate N-formylated peptides.
In this work, peptide mass fingerprinting has established that fMAP, as purified in this laboratory, is actually APH. We believe that Chadwick and coworkers (131) had previously purified the same enzyme as we did, based on the following reasons. First, both Chadwick et al. and we detected only one fraction that has detectable fMAP activity during purification. Both laboratories employed similar (though not identical) procedures to purify the enzyme. Second, the enzymes from two laboratories share similar substrate specificities and sensitivities to effector molecules. Several other observations also support the notion that Chadwick’s fMAP is the same as APH. Chadwick et al. showed that their fMAP has a native Mr of 340,000 Da and a Mr 82,000 Da for each subunit
(131), matching the molecular weights reported for full-length APH (143). In contrast,
fMAP purified in this laboratory has a Mr 55,000, corresponding to the C-terminal
peptide hydrolase domain (144, 145). The precise location or mechanism of proteolytic
cleavage of fMAP is presently unknown. APH and Chadwick’s fMAP both have the
same pH optimum (pH 7.9-9.0 for fMAP and pH 8.2 for APH) (131, 146). Chadwick et
al. reported that fMAP is broadly distributed in the body tissues including intestine,
colons, rectum, kidney, and liver (131). The previously characterized rat APH was
purified from liver tissues (136, 143, 146, 147). It appears that Chadwick and co-workers
had mistaken fMAP as an enzyme distinct from APH due to their poor choice of assay
substrates. These investigators examined the substrate specificity of their enzyme by
121 testing against 8 different formyl-amino acid-β-naphthylamide derivatives. They found
that the enzyme was active only against the f-Met derivative but had no activity against
the other derivatives (f-Ala, f-Val, f-Leu, f-Ser, f-Asp, f-Arg, or f-Phe-β-naphthylamides)
(131). All of the peptide substrates they tested contained an N-terminal methionine
residue. These results led them to conclude that fMAP is “specific” for an N-terminal f-
Met or acyl-Met moiety and thus distinct from APH, which has much broader substrate
specificity (136, 143, 146, 147). fMDF activities have previously been described in the
crude extracts of Euglena gracilis (141), rabbit reticulocyte lysates (134), human leukocytes and platelets (140), and a variety of other animal tissues (151). Chadwick et al. partially purified it from rat small intestines and examined its substrate specificity and sensitivity to a variety of effector molecules (131). However, the identity of fMDF has not been established. It was suggested by Grisolia et al. that fMDF might represent a previously unrecognized activity of acylase I (151). We have now shown that fMDF is indeed an acylase I variant. Acylases cleaves N-acylamino acids into free amino acids and are thought to be important in the degradation of N-acylated peptides and proteins.
According to their substrate specificities, the acylases characterized so far can be classified into four different types, I to IV. Acylase I is responsible for the hydrolysis of neutral and hydrophobic N-acyl-L-amino acids. Two different forms of acylase I (IA and
IB) have been found in rat kidney (152) and human liver (153), with 94% sequence identity between the two rat acylases. fMDF purified from rat intestinal mucosa corresponds to acylase IA. All of the peptides in Table 4 matched with tryptic fragments derived from acylase IA but not IB. Acylases IA and IB both contain 408 amino acids with a calculated molecular mass of ~45 kDa (152, 153). They are metalloproteins 122 containing a single Zn2+ ion per polypeptide, explaining why the activity of fMDF
(acylase IA) is stimulated by divalent metal ions (Co2+ and Ni2+) but inhibited by metal chelators such as EDTA and 1,10-phenanthroline.
Despite of the broad distribution of APH and acylase I in animal tissues, their physiological functions have not been well understood other than the degradation of N- acylated peptides and proteins. Our current work reveals a second function for these enzymes, i.e., degradation of N-formylated peptides. Indeed, both APH and acylase I hydrolyzes N-formylated peptides with essentially the same efficiencies as their N- acetylated counterparts (Tables 6.5 and 6.6). Further, acylase I and APH genes in human and porcine have been mapped to chromosomes 3p21.1 and 3p21.3, respectively (154-
158). The co-localization of these genes on similar chromosome suggests that these enzymes may be functionally linked in the sequential degradation of N-acylated peptides.
By efficient degradation of chemotactic peptides such as f-MLF, APH and acylase I may provide the first line of defense against unwanted inflammatory responses at tissue sites that are in constant contact with commensal bacteria (e.g., intestines). The abundance of
these two enzymes coupled with their robust activities toward N-formylated peptides also
suggests that treatment with PDF inhibitors is unlikely to induce inflammatory responses
in a patient.
123
Purification Stepa Total protein Specific Activityb Purification Recovery (mg) (nmol/min per mg protein) (fold) (%)
Crude Sonicate 1944 8 1 100 Anion Exchange 320 240 29 67 Phenyl-Sepharose 3 3143 374 43 Mono Q 1.2 8547 1018 33 Gel Filtration 0.7 17,313 2061 20
aProcedure for purifying N-Acylaminopeptidase from rate intestinal epithelial mucosal layer are detailed in the Materials and methods. bThe substrate for the purification and activity assay was f-Met-p-NA.
Table 6.1. Purification of fMAP from Rat Epithelial Mucosal Layer.
124
Purification Stepa Total Protein Specific Activityb Purification Recovery (mg) (nmol/min per mg protein) (fold) (%)
Crude Sonicate 2212 1.2 1 100 Acidication 575 1.9 2 90 CM-Sepharose 112 3.9 3 85 Q-Sepharose 45 89.0 74 56 Phenyl-Sepharose 5 320.3 267 40 Size Exlusion 0.5 1899.1 1583 32 Concavalin A-Sepharose 0.3 2199.3 1833 26
aProcedure for purifying N-acylase I from rate intestinal epithelial mucosal layer are detailed in the Materials and methods. bThe substrate for the purification and activity assay was fMet.
Table 6.2. Purification of fMDF from Rat Epithelial Mucosal Layer.
125
Peptidea Observed Mr(calc) Mr(expt) (M+H+)
N691VPRV695 584.49 583.34 583.48 R674VPFK678 646.55 645.40 645.54 Y274CbTNR278 713.47 712.30 712.46 Y274CbTNRR279 869.62 868.40 868.61 L307SPDQCbR313 875.63 874.40 874.62 V435GFLPPPGK443 911.70 910.53 910.69 V376VFDSAQR383 921.72 920.47 920.71 Q682GMEYYR688 946.65 945.40 945.65 Q682GMcEYYR688 962.65 961.40 961.64 V340TSVVVDIVPR350 1183.97 1182.70 1182.96 M528GFAVLLVNYR538 1282.99 1281.69 1281.98 T665PVLLMLGQEDR676 1372.03 1370.72 1371.03 T665PVLLMcLGQEDR676 1388.00 1386.72 1386.99 T665PVLLMLGQEDRR677 1528.15 1526.82 1527.15 C292(c)ELLSDGSLAICcSPR306 1678.16 1677.79 1677.16 D562VQFAVEQVLQEEHFDAR579 2160.41 2159.03 2159.41 G539STGFGQDSILSLPGNVGHQDVK561 2313.53 2312.14 2312.53 E444QSVSQVSLEEAEPIPGIHWGVR466 2605.68 2604.30 2604.67 Q351LGESFSGIYCcSLLPLGCcWSADSQR375 2831.67 2830.31 2830.66
aPeptide fragments shown correlate to indicated regions within the N-acylaminopeptidase polypeptide bCarbamidomethyl cysteine cOxidation of methionine
Table 6.3. MS Analysis of N-acylaminoacyl-peptide Hydrolase under Reducing and Denaturing Conditions Following Digestion with Trypsin.
126
Peptidea Observed Mr(expt) Mr(calc)
R161PEFQALR168 508.94 1015.87 1015.56 V222VNSILAFR230 509.97 1017.93 1017.60 E245GAVTSVNLTK255 559.99 1117.96 1117.60 S116VSIQYLEAVR126 633.05 1264.08 1263.68 L393VAALASVPALPGES408 698.12 1394.23 1393.78 A354VGIPALFGSPMNR367 715.61 1429.21 1428.75 A354VGIPALGFSPMbNR367 723.62 1445.23 1444.75
aPeptide fragments shown correlate to indicated regions within the N-acylase I polypeptide bOxidation of Methionine
Table 6.4. MS/MS Analysis of N-Acylase I under Reducing and Denaturing Conditions Following Digestion with Trypsin
127
-1 -1 -1 Substrate KM (mM) kcat(s ) kcat/KM (M s )
f-M-pNA 0.4±0.2 6.2±2.0 120,000±3000 f-MLF 2.0±0.8 4.6±2.1 2300±645 0.18a 12.3a 68,000a f-MF 0.6±0.2 8.0±3.3 13,000±8000 0.63a 70.2a 110,000a f-MA 0.5±0.1 3.2±0.9 6400±644 f-MAS 2.3±0.8 5.6±2.1 2400±500 f-MS 1.0±2.0 6.1±1.2 6100±322 f-MSS 5.5±2.0 4.2±0.9 764±176 f-MSSS 11.4±3.0 4.0±1.1 351±131 f-MSSSS 11.6±2.4 2.1±1.3 181±88 f-MSSSSS 30.3±9.5 1.3±1.2 43±20 Ac-MAS 1.5±0.3 4.4±0.1 2933±66 Ac-AA 1.0±0.3 14.7±4.5 15,000±3000 Ac-AAA 1.7±0.4 13.3±4.8 8,000±2000
aLiterature Values (reference (131))
Table 6.5. Kinetic Activity of N-acyl Aminopeptidase from Rat Intestine Epithelial Mucosal Layer
128
-1 -1 -1 Substrate KM (mM) kcat (s ) kcat/KM (M s ) f-Met 3.1±0.1 7.9±0.1 2550±77 7.1a Ac-Met 0.3±0.1 2.3±0.9 7600±500 0.22a f-Met-Ala NA NA NA f-Ala-Ala NA NA NA f-Met-Ala-Ser NA NA NA Ac-Met-Ala-Ser NA NA NA f-Met-Leu-Phe NA NA NA
NA = No Activity Detected aLiterature Values (reference (132))
Table 6.6. Kinetic Activity of N-Acylase I from Rat Intestine Epithelial Mucosal Layer
129
fMAP f-Met-Leu-Phe f-Met + Leu-Phe
fMDF
HCO2H + Met
Figure 6.1. Enzymes from rat small intestine epithelial mucosal layer involved in degradation of N-formylmethionine peptides.
130
Figure 6.2. Chromatography of an N-formylaminopeptide hydrolytic enzyme. This purification scheme illustrates the isolation of an enzyme catalyzing the degradation of both f-M-pNA and f-MLF chemopeptide substrates. Methods are described under the procedure section. (A) Purification on an anion-exchange chromatography column (Q- Sepharose). (B) Fractions from Q-sepharose with N-formylaminopeptide hydrolase activity loaded onto a hydrophobic-interaction chromatography column (Phenyl- sepharose). The absorbance at 280 nm (solid line tracing), the specific activity for f-M- pNA (dashed line tracing) and f-MLF chemopeptide (dotted line tracing) of each fraction was measured.
131
97.4 66.2 45.0
31.0
21.5
1 2
Figure 6.3. SDS-PAGE of rat intestinal N-acyl aminopeptidase. The electrophoresis was performed with 10 % polyacrylamide gel under reducing agent dithiolthreitol and sodium dodecylsulfate. The gel was stained with coomassie brilliant blue: Lane 1, molecular weight marker, Lane 2, purified N-acylaminopeptidase. The following molecular weight markers (kDa) were used: rabbit muscle (97.4), bovine serum albumin (66.2), hen egg white ovalbumin (45.0), carbonic anhydrase (31.0), and trypsin inhibitor (21.5).
132
97.4 66.2
45.0
31.0
21.5
12
Figure 6.4. SDS-PAGE of rat intestinal N-acylase I. The electrophoresis was performed with 12 % polyacrylamide gel under reducing agents dithiolthreitol and sodium dodecylsulfate: Lane 1, molecular weight marker, Lane 2, purified N-acylase I. The following molecular weight markers (kDa) were used: rabbit muscle (97.4), bovine serum albumin (66.2), hen egg white ovalbumin (45.0), carbonic anhydrase (31.0), and trypsin inhibitor (21.5).
133
CHAPTER 7
MATERIALS AND GENERAL METHODS
7.1 Buffers. All of these buffers were used through out the chapters (2-6) and referred to as the indicated buffer number in the text. Buffer 1, 20 mM Tris.HCl, pH 8.5,
10 mM NaCl, 1% Triton X-100, and 0.5% protamine sulfate; buffer 2, 20 mM Mes, pH
6.0, and 10 mM NaCl; buffer 3, 20 mM Tris–HCl, pH 8.0, 0.5 M NaCl, and 5 mM imidazole; buffer 4, 20 mM Hepes, pH 7.4, and 10 mM NaCl; buffer 5, 20 mM Mops, pH
6.5, 50 mM KCl, 5 mM NiCl2, 1% Triton X-100, and 0.5% protamine sulfate; buffer 6,
20 mM Tris.HCl, pH 7.9, 0.5 M NaCl, and 5 mM imidazole; buffer 7, 20 mM Hepes, pH
7.4, and 150 mM NaCl; buffer 8, 50 mM Hepes (pH 7.2), 100 mM NaCl, 1 mM β-
mercaptoethanol, and 1% Triton X-100; buffer 9, 20 mM Hepes (pH 7.2), 100 mM NaCl;
buffer 10, 50 mM Tris·HCl, 10 mM NaCl, pH 8.0; buffer 11, 50 mM Tris·HCl (pH 8.0), 1
mM NaCl; buffer 12, 50 mM Hepes (pH 7.0), 150 mM NaCl; buffer 13, 20 mM Hepes
(pH 7.2), 100 mM NaCl, 0.2 mM -mercaptoethanol; buffer 14, 50 mM Mops (pH 7.0), 10
mM NaCl; buffer 15, 50 mM Hepes, 10 mM NaCl, 5 mM dithiothreitol, pH 7.0; buffer
16, 100 mM Tris, 100 mM NaCl, 10 mM CaCl2, 0.04% Brij-35; buffer 17, 50 mM Mops,
pH 7.0; buffer 18, 150 mM NaCl, 30 mM Tris, pH 7.5; buffer 19, 20 mM Tris, 10 mM
NaCl, pH 8.0; buffer 20, 1.7 M (NH4)2SO4, 100 mM KH2PO4, pH 7.0; buffer 21, 100 mM
134 KH2PO4, pH 7.2; buffer 22, 50 mM Tris, 75 mM NaCl, pH 7.5; buffer 23, 150 NaCl, 50
mM Hepes, pH 7.0; buffer 24, 200 mM KCl, 200 mM sodium borate, pH 9.0; buffer 25,
10 mM sodium phosphate, 0.5 M NaCl, pH 7.5; buffer 26, 20 mM sodium phosphate, pH
7.4 ; buffer 27, 50 mM sodium acetate, pH 5.5.
7.2 Materials. Escherichia coli PDF was overexpressed in E. coli (BL21(DE3)) and
purified to apparent homogeneity with either Fe2+ or Co2+ as the divalent metal (38, 42).
DPPI (cathepsin C) from bovine spleen and yeast formate dehydrogenase were purchased
from Sigma Chemical (St. Louis, MO). Other chemicals were obtained from either
Sigma–Aldrich (St. Louis, MO) or Bio-Rad Laboratories (Hercules, CA) and Advanced
ChemTech (Louisville, KY). PDF inhibitors were synthesized by Dr. Xubo Hu in this
lab, as previously described (79, 159).
All Fmoc-protected amino acids, 2-(1H-benzotriazol-1-yl)-1,1,3,3- tetramethyluronium hexafluorophosphate (HBTU), and 1-hydroxybenzotriazole (HOBT) were purchased from SynPep (Dublin, CA). Rink resin was from Advanced Chemtech
(Louisville, KY). Formate dehydrogenase was from Sigma. Aeromonas aminopeptidase was purified as previously described (72). Other chemicals, including isopropyl-β-D- thiogalactopyranoside (IPTG), phenylmethanesulfonyl fluoride, kanamycin, ethanedithiol and its derivatives, and β-mercaptoethanol were purchased from Aldrich.
H. influenzae PDF was overproduced in E. coli with cobalt(II) as the metal cofactor and a C-terminal histidine tag and was purified on a cobalt(II) affinity column in a manner similar to that described for E. coli and B. subtilis PDF (42, 87). Mca-P-K-G-
Dpa-A-R-NH2 fluorogenic peptide substrate I and Mca-R-P-K-P-V-E-Nval-W-R-
135 K(Dnp)-NH2 fluorogenic peptide substrate II were purchased from R&D Systems
(Minneapolis, MN). Human MMPs (MMP-1, MMP-2, MMP-3, and MMP-9) were
purchased from Sigma Chemical Co. (St. Louis, MO). Dulbecco' Modified Eagle
Medium, Lipofectamine 2000, fetal bovine serum, and phosphate buffered saline required
for HEK 293 cell growth were purchased from Life Technologies. Other chemicals were obtained from either Sigma-Aldrich (St. Louis, MO) or Bio-Rad Laboratories (Hercules,
CA). Spectroscopic measurements were performed on a Perkin-Elmer Lambda 20 UV/vis
spectrophotometer.
Small intestines from Sprague-Dawley rats were purchased from Zivic
Laboratories (Pittsburg, PA). NAD+, Triton X-100, and Candida boidinii formate
dehydrogenase (FDH) were purchased from Sigma. f-MLF, fluoresamine, p-nitroanaline,
1,10-phenanthroline, and diethyl pyrocarbonate were purchased from Aldrich. f-Met, Ac-
Met, and other peptide derivatives were purchased from Bachem Biosciences Inc.
Reagents for peptide synthesis were purchased from Advanced ChemTech (Louisville,
KY).
PDF substrates and peptide thiol inhibitors were synthesized as previously
described (38, 87). Actinonin was purchased from Sigma Chemical Co. All other
reagents were obtained from commercial suppliers and used directly in the experiments.
136 7.3 Synthesis of Substrates for DPPI Assay
7-(Nα-Benzyloxycarbonyl-Nε-t-butoxycarbonyl-L-lysyl) amino-4-
methylcoumarin (N-Cbz-Lys(Boc)-AMC). To a solution of Cbz-Lys(Boc)-OH (0.42 g,
1.1 mmol), 7-amino-4-methylcoumarin (0.17 g, 1.0 mmol) and HOBt (0.17 g, 1.1 mmol) in dichloromethane (10 mL) was added EDC (0.23 g, 1.2 mmol) at 0 ºC. The reaction was stirred overnight at room temperature. The mixture was diluted with 80 mL of ethyl acetate and washed with dilute HCl (2 x 20 mL), water (20 mL), 5% NaHCO3 (2 x 20
mL), and brine (20 mL). The organic layer was dried over Na2SO4, filtered and
concentrated to dryness. The residue was subjected to flash chromatography using
dichloromethane/ethyl acetate as eluent to afford 190 mg of a white solid (35% yield). 1H
NMR (CDCl3, 250 MHz) δ 9.22 (s, 1H), 7.54 (s, 1H), 7.25 (m, 6H), 6.00 (s, 1H), 5.84 (d,
J = 7.5 Hz, 1H), 4.99 (s, 2H), 4.65 (br s, 1H), 4.27 (m, 1H), 2.97 (m, 2H), 2.23 (s, 3H),
1.82 (m, 1H), 1.35 (m, 4H), 1.28 (s, 9H).
7-(N-formylmethionyl-Nε-t-butoxycarbonyl-L-lysyl) amino-4-
methylcoumarin (f-Met-Lys(Boc)-AMC). N-Cbz-Lys(Boc)-AMC (90 mg, 0.17 mmol)
was dissolved in a solvent mixture containing 1:1 (v/v) methanol and ethyl acetate (10 mL) and hydrogenated in the presence of Pd/C at room temperature for 2.5 h. After the reaction was completed, the catalyst was removed by filtration and the filtrate was concentrated to dryness. The residue was dissolved in dichloromethane (6 mL), N- formylmethionine (37 mg, 0.21 mmol), HOBt (32 mg, 0.21 mmol), and EDC (40 mg,
0.21 mmol) were sequentially added at 0 ºC. The mixture was stirred for 2 h at room temperature. After that the mixture was diluted with 50 mL of ethyl acetate, washed with dilute HCl (2 x 15 mL), water (15 mL), 5% NaHCO3 (2 x 15 mL), and brine (15 mL).
137 The organic layer was dried over Na2SO4, filtered and concentrated to dryness. The
residue was purified by silica column chromatography using dichloromethane/ethyl
1 acetate as eluent to give 60 mg of product (64% yield for two steps). H NMR (CDCl3,
250 MHz) δ 9.76, (s, 1H), 8.28 (s, 1H), 7.84 (d, J = 7.5 Hz, 1H), 7.68 (d, J = 7.5 Hz, 1H),
7.65 (s, 1H), 7.42 (m, 2H), 6.12 (s, 1H), 4.91 (m, 2H), 4.67 (m, 1H), 3.07 (m, 2H), 2.63
(m, 2H), 2.36 (s, 3H), 2.20–2.03 (m, 2H), 2.05 (s, 3H), 1.95 (m, 1H), 1.83 (m, 1H), 1.45
(m, 4H), 1.39 (s, 9H).
7-(N-formylmethionyl-L-lysyl) amino-4-methylcoumarin (f-Met-Lys-AMC).
f-Met-Lys(Boc)-AMC (50 mg) was dissolved in dichloromethane (2 mL) and
trifluoroacetic acid (1 mL) was added. After the mixture was stirred for 2 h, the volatile
substances were removed under vacuum to afford the product in quantitative yield. The
product was neutralized by dissolving in 1:1 (v/v) dichloromethane/isopropyl alcohol (10
mL) containing 10% triethylamine. 1H NMR (D6-DMSO, 250 MHz): δ 10.45 (s, 1H),
8.37 (m, 2H), 8.04 (br s, 1H), 7.76 (m, 2H), 7.65 (br s, 3H), 7.50 (dd, J = 2.0, 8.7 Hz),
6.27 (s, 1H), 4.44 (m, 2H), 2.79 (m, 2H), 2.46 (m, 2H), 2.40 (s, 3H), 2.04 (s, 3H), 1.85–
+ + 1.49 (m, 8H). ESI-HRMS: calcd for C22H31N4O5S (M + H ): 463.2009. Found 463.2004.
7-(Nα-9-fluorenylmethoxycarbonyl-Nγ-trityl-L-glutamyl) amino-4-
methylcoumarin (N-Fmoc-Gln (Trt)-AMC). To an ice-chilled solution of Fmoc-
Gln(Trt)-OH (244 mg, 0.4 mmol) and triethylamine (40 mg, 0.4 mmol) in tetrahydrofuran
(7 mL), ethyl chloroformate (48 mg, 0.44 mmol) was added drop-wise. The mixture was
stirred at 0 ºC for 10 min. 7-Amino-4-methylcoumarin (68 mg, 0.4 mmol) was added and
the solution was stirred overnight at room temperature. The solvent was evaporated and
the residue was dissolved in 50 mL of dichloromethane and washed with 0.5 M HCl (2 x
138 20 mL), water (20 mL), 5% NaHCO3 (2 x 20 mL), and brine (20 mL). The organic layer was dried over Mg2SO4, filtered, and concentrated. The crude product was purified by
silica gel chromatography using dichloromethane/ethyl acetate as eluent to give 40 mg of
1 a white solid. H NMR (CDCl3, 400 MHz): δ 7.77 (d, J = 5 Hz, 1H), 7.49 (s, 1H), 7.44 (s,
1H), 7.40–7.27 (m, 26H), 6.97 (s, 1H), 6.18 (s, 1H), 4.41 (br s, 1H), 4.15 (m, 2H), 2.80
(m, 2H), 2.55 (m, 2H), 2.39 (s, 3H).
7-(N-Formylmethionyl-L-glutamyl) amino-4-methylcoumarin (f-Met-Gln-
AMC). N-Fmoc-Glutamyl(Trt)-AMC (40 mg, 0.053 mmol) was dissolved in 20%
piperidine dichloromethane (5 mL) and stirred at room temperature for 45 min. The
reaction was purified by silica gel chromatography using ethyl acetate/dichloromethane
as eluent to give 20 mg white solid. The residue was redissolved in dichloromethane (7 mL), and N-formylmethionine (8.8 mg, 0.05 mmol), 1-hydroxy-7-azabenzotriazole
(0.037 mmol), and 1-ethyl-3-( (dimethylamino)propyl)-carbodiimide (EDC) (7.6 mg,
0.038 mmol) were sequentially added at 0 ºC. The reaction was stirred overnight at room temperature and followed by dilution into 40 mL dichloromethane. The mixture was washed with 0.5M HCl (2 x 20 mL), water (20 mL), 5% NaHCO3 (2 x 20mL) and brine
(20 mL). The organic layer was dried over Mg2SO4, filtered, and concentrated. The
residue was purified by silica gel column chromatography using dichloromethane/ethyl
acetate as eluent to give 15 mg of 7-(Nα-formylmethionyl-N-trityl-L-glutamyl) amino-4- methylcoumarin (58% yield). This purified compound (15 mg) was dissolved in 4.5 mL trifluoroacetic acid, 0.25 mL thioanisole, 0.1 mL anisole, and 0.15 mL ethanedithiol. The
mixture was stirred at room temperature for 1 h and followed with removing the volatile
substances with nitrogen gas. The remaining residue was triturated with ether (3 x 15
139 mL) to give 10 mg of white solid (quantitative yield). The product was neutralized by
redissolving in 1:1 dichloromethane/isopropyl alcohol containing 10% triethylamine (10
mL). The solvent was evaporated by nitrogen gas and the residue was triturated with
ether (2 x 15 mL). 1H NMR (D6_MSO, 400 MHz) δ 10.42 (s, 1H), 8.41 (d, J = 4 Hz, 1H),
8.34 (d, J = 4 Hz, 1H), 8.10 (s, 1H), 7.77 (m, 2H), 7.52 (d, J ¼ 4 Hz, 1H), 7.3 (s, 1H),
6.80 (s, 1H), 6.28 (s, 1H), 4.46 (m, 1H), 4.37 (m, 1H), 2.5 (m, 2), 2.4–2.2 (m, 6H), 2.15
+ + (s, 1H), 2.05 (s, 3H), 1.92 (s, 3H). ESI-HRMS: calcd for C21H26N4O6SNa (M + Na ):
485.1465. Found: 485.1465.
7.4 General Biochemical and Biological Methods
7.4.1 Materials. Restriction enzymes, T4 DNA ligase, and Vent polymerase were obtained from New England Biolabs (NEB). Calf intestinal alkaline phosphatase was obtained from Sigma. All enzymes were used in buffers suggested by the suppliers.
Phenol was purchased from Fisher Scientific prepared (160) prior to use and stored in 10 mM Tris.HCl, pH 8.0, 1 mM EDTA, 0.1% β-mercaptoethanol at 4 ◦C for up to 1 month. Chloroform mixture is indicated by 24:1 chloroform and 3-methyl-1-butanol.
Chloroform is ACS certified. Deoxynucleotide mixture was from Stratagene. Ethidium bromide, bromophenol blue, xylene cyanol, tris (hydroxymethyl) aminomethane (Tris),
diethylpyrocarbonate (DEPC), dithiothreitol, 2-(N-morpholino) ethanesulfonic acid)
(HEPES), ethylenediamine tetra-acetic acid (EDTA), isopropyl-β-D-thiogalactosidase
(IPTG), all antibiotics, and β-mercaptoethanol were from Sigma.
140 7.4.2 Growth Media. Dry bacterial growth media (bactotryptone, yeast extract, casamino acids, and bactoagar) were obtained from Difco. All growth media were prepared with ddH2O and sterilized by autoclaving for 20 min. at 20 lb/sq. in. on liquid cycle. Inorganic salts, 20% (w/v) glucose, 1 M MgSO4, and 1% thiamine hydrochloride were dissolved in ddH2O and sterilized by passing through sterile 0.45 µM filters
(Acrodisc, Gelman Sciences). Antibiotics were added directly to the growth media after cooling to less than 50 ◦C.
Luria-Bertani Medium (LB): 10 g/L bactotryptone, 5 g/L yeast extract, 10 g/L NaCl.
LB Plates: LB medium plus 15 g/L bactoagar.
Top Agar: 10 g/L bactotryptone, 5 g/L NaCl, 6.4 g/L bactoagar.
Minimal Medium: 2.5 g/L D-(+)-glucose, 5 g/L casamino acids, 10.8 g/L
K2HPO4, 5.5 g/L KH2PO4, 10 g/L NaCl, 1.0 g/L ammonium sulfate, 2 mg/L
thiamine, 1 mg/L (+)-biotin and supplemented with metals 35 mg/L
. . . MgSO4 7H2O, 10 mg/L CaCl2 2H2O, 4.7 mg/L MnCl2 4H2O, 4 mg/L H3BO3, 1.5 mg/L
. (NH4)6Mo7O24 4H2O, 0.45 mg/L CuSO4.
Escherichia coli Strains. The following strains were used in this work.
- - - BL21(DE3) F ompT hsdSB (rB mB ) gal dcm (DE3). This strain carries a copy of T7
RNA polymerase gene on its chromosome (λ lysogen). It has neither
methylation nor restriction function, and therefore it is useful for preparing
DNA free of methylation. It is also deficient in lon protease and is good
for overproduction of foreign proteins.
141 - + r DH5αF’ F’/endA1 hsdR17 (rk mk ) supE44 thi-1 recA1 gyrA (NaI ) relA1
∆(lacZYA-argF)U169 deoR (Ф80dlac∆9lacZ)M15. This train is deficient
in DNA recombination. Plasmid DNA prepared from this strain usually
has very high quality. It is also used to overproduce proteins.
BL21(DE3) (Rosetta)
This strain is similar to the BL21(DE3) except it contains pRARE (CmR).
The strain supplies tRNAs for the codons AUA, AGG, AGA, CUA, CCC,
and GGA on a compatible chloramphenicol-resistant plasmid. This strain
is used to express gene sequences that are codon bias, i.e. HsPDF.
7.4.3 Growth and Storage of Bacterial Strains. E. coli were stored on agar plates at
4 ◦C for periods up to one month, as stab cultures for periods up to 2 years, or as frozen
glycerol cultures at -80 ◦C indefinitely.
E. coli strains DH5αF’ and BL21(DE3) were streaked on LB plate and strain
BL21(DE3) (Rosetta) was streaked on LB plate plus chloramphenicol (35 mg/L). Cells
from a frozen glycerol culture were streaked on an appropriate plate and then incubated at
37 ◦C until individual colonies were 1-2 mm in diameter.
Stab culture was prepared by picking a single, well-isolated colony with a sterile
inoculating needle and stabbing the needle several times through the sterilized LB agar
(by autoclaving) in a 4 mL glass vial. The vial was tightly sealed with a screw-on cap
fitted with a rubber gasket and stored in the dark at room temperature.
To store cells as frozen glycerol culture, 0.5 mL of liquid bacterial culture was added to an equal volume (0.5 mL) of 50% (v/v) glycerol in water (sterilized by
142 autoclaving) in a sterile microcentrifuge tube (1.5 mL). The culture was mixed with the glycerol by vortexing, frozen in isopropyl alcohol-dry ice bath, and transferred to -80 ◦C
for long-term storage.
Overnight culture was prepared by picking a single, well-isolated colony from an
agar plate and using it to inoculate 1-10 mL of the appropriate sterile medium. Cells
were allowed to grow at 37 ◦C for 8-16 hr.
7.4.4 Preparation of Competent Cells. Unless otherwise noted, all sterile pipettes,
tubes, and solutions were pre-chilled to 4 ◦C, and all processes were performed on ice or in a cold room. 250 mL of LB media was inoculated by the addition of 2.5 mL of the
overnight culture of an appropriate strain. The cells were allowed to grow with aeration
◦ at 37 C until OD600 reached 0.4-0.6. The cells were transferred to a sterile GS-3
centrifuge tube, cooled on ice 5-10 min., and centrifuged at 5,000 rpm for 5 min. The
pellet was gently resuspended in 10 mL of ice cold competent solution A (30 mM KOAc,
pH 5.8, 100 mM RbCl, 10 mM CaCl2, 50 mM MnCl2, and 15% (v/v) glycerol) and kept
on ice for 5 min. The cell suspension was centrifuged again (5,000 rpm, 5 min.) and the
resulting pellet was resuspended in 10 mL of ice cold competent solution B (10 mM
MOPS, pH 6.5, 75 mM CaCl2, 10 mM RbCl, and 15% (v/v) glycerol). Competent cells
in solution B were stored at -80 ◦C for up to 6 months.
7.4.5 Extraction with Organic Solvents. Proteins and agarose (after purification in
agarose gels) were removed from preparations of nucleic acids by extraction with phenol
and chloroform. Generally, a solution of nucleic acid was extracted once with phenol,
143 1:1 (v/v) mixture of phenol/chloroform, and chloroform. An equal volume of the organic
solvent was added to the aqueous solution of nucleic acids, and the contents were
thoroughly mixed by gently shaking or inverting the tubes. The mixture was then
separated into two layers by a microcentrifuge (14,000 rpm, 3 min). The upper layer
contains the nucleic acids was carefully pipetted off and transferred into a clean tube.
Extracted nucleic acid samples were precipitated with ethanol.
7.4.6 Quantitation of DNA and RNA. Pure samples of DNA and RNA (i.e. free of
proteins, phenol, agarose, nucleotides, or other nucleic acids) were quantitated by
measuring the absorbance at 260 nm using a UV/vis spectrophotometer as described
(160). An OD260 of 1 corresponded to approximately 50 µg/mL for double-stranded
DNA, 40 µg/mL for large single-stranded DNA, and 27 µg/mL for single-stranded
oligonucleotides. The purity of a preparation of DNA or RNA was estimated by reading
its ratio of OD260/OD280. Pure preparations of DNA and RNA should have OD260/OD280 values of 1.8 and 2.0, respectively. When the amount of a sample was small (<1 µg) or the sample was heavily contaminated with other substances that absorbed at 260 nm, the quantity of DNA (or RNA) in the sample was estimated by comparing the fluorescence yield of the sample with that of a series of standards (160).
7.4.7 Removal of Small Molecules from High-molecular-weight DNA and RNA.
To remove unincorporated dNTPs from DNA polymerase reaction, a mini spin column from Qiagen (Qiaquick) was used according to the manufacture protocol. RNA can be
144 removed simply by adding 1 µg of RNAse A to the DNA solution and passed through a
Qiagen column.
7.5 Electrophoresis
7.5.1 Agarose Gel. Agarose gel electrophoresis was used for separation of large DNAs
(>150 bp). Agarose (Sigma, electrophoresis grade) was added to 100 mL of 0.5x TBE
buffer (44.5 mM Tris, 41.5 mM boric acid, 1 mM EDTA, pH 8.3) to make final
concentration of 0.5-1.5% (w/v). The agarose was melted by heating in a microwave and
2 µL of an aqueous solution of ethidium bromide (10 mg/mL) was added. After mixing by shaking, the agarose was poured into a horizontal slab gel tray (110 x 140 mm) and allowed to solidify at room temperature.
DNA samples were mixed with 1/6 volume of a 6x loading buffer (30% (v/v) glycerol, 0.25% (w/v) bromophenol blue in ddH2O) and loaded into 5 x 2 (1) x 5-10 mm
(length x width x depth) wells. Electrophoresis was carried at constant voltage (100 V)
submerged in 0.5x TBE buffer until the desired separation was achieved. The DNA-
containing bands were visualized under either long-wavelength (preparative) or short-
wavelength (analytical) UV light.
7.5.2 Polyacrylamide Gels for Protein Separation. Proteins were separated by
sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) according to the
method of Laemmli (161). The gel consisted of a stacking layer (175 x 30 x 1.5or 0.75
mm) and a separating layer (175 x 120 x 1.5 or 0.75 mm). The percentage (10-20%) of
the acrylamide of the separating layer depended on the sizes of the proteins in the sample.
145 An appropriate volume of a 30% acrylamide stock solution (bis(acrylamide): acrylamide ,
. 1:29) was diluted with ddH2O, separating gel buffer (325 mM Tris HCl, pH 8.8, 0.1%
SDS (w/v)) and 1.6 mg/mL ammonium persulfate. The resulting solution was degassed,
activated by the addition of 8.0 µL of TEMED, and poured between two assembled glass plates. The surface of the separating gel was smoothed by adding a few drops of n- butanol saturated with water on the top of the gel. The gel was allowed to polymerize at ambient temperature for 15-20 min and the n-butanol was removed by rinsing the gel with water. The stacking gel solution containing 4% acrylamide mixture, 1.6 mg/mL ammonium persulfate, and 8.0 µL of TEMED in 1x stacking gel buffer (125 mM
Tris.HCl, pH 6.8, 0.1% SDS) was then added onto the top of the polymerized separating
gel. The sample loading wells were formed by inserting combs of the appropriate width
into the top of the stacking gel.
Protein samples were mixed with an equal volume of a 2x loading buffer (125 mM Tris.HCl, pH 6.8, 4% (w/v) SDS, 10% (v/v) β-mercaptoethanol, 20% (v/v) glycerol,
0.2% (w/v) bromophenol blue) and heated at 95-100 ◦C for 5 min prior to loading. The
gels were electrophoressed in Tris-glycine buffer (25 mM Tris, 192 mM glycine, 0.1%
SDS, pH 8.0) at 200 V until the bromophenol blue had migrated near the bottom of the
gel. The gels were then removed from the glass plates, stained in 40% methanol, 10%
acetic acid in water containing 0.25% (2 hr staining) or 0.05% (overnight staining)
Coomassie Brilliant Blue R-250, and destained by soaking in a solution of 40% (v/v)
methanol, 10% acetic acid in water.
146 7.6 Recombinant DNA techniques
7.6.1 Restriction Digestions. All restriction digestions were performed in buffers
suggested by or obtained from commercial suppliers. Complete digestions were usually carried out at 37 ◦C for 1-8 hr using 1.5 units of restriction enzymes for 1 µg of plasmid
DNA. More enzymes (0.4-1 unit/pmol DNA) were used for every microgram of smaller
DNA fragments (synthetic DNAs, PCR products, or restriction fragments). If required,
the restriction enzymes were removed after digestion by extraction twice with
phenol/chloroform, once with chloroform, and DNA recovered by ethanol precipitation.
7.6.2 Filling Recessed 3’-Termini and Removing Protruding 3’-Termini. In order
to ligate DNA fragments with incompatible termini, they were converted into compatible
forms by partially or completely filling-in the recessed 3’-termini or removing the
protruding 3’-termini. To fill-in a recessed 3’-terminus, 1 µL of a solution containing the
desired dNTPs (depended on the sequence of the 5’ overhang and whether partial or
complete fill-in was required) at 1 mM was added directly to 0.2-5 µg of DNA (20 µL in
restriction enzyme buffer plus 5 mM MgCl2) digested with appropriate restriction
enzymes. After the addition of the Klenow fragment of DNA polymerase I (1 unit for
every µg of DNA), the reaction mixture was incubated at room temperature for 15 min.
Removal of protruding 3’ termini was carried out similarly except that T4 DNA
polymerase was used instead of the Klenow fragment, height concentrations of dNTPs
were added (1 µL of 2 mM each), and the reaction was incubated at 14 ◦C for 15 min.
After the reaction, enzymes and dNTPs were removed by either agarose gel
electrophoresis or Qiagen Kit (Qiaquick mini column).
147 7.6.3 Removal of 5’ Phosphates. In order to prevent undesired self-ligation of a DNA
fragment, the 5’ phosphate groups were removed with calf intestinal alkaline phosphatase
(CIP). Dephosphorylation was carried out in a 50 µL reaction containing digested DNA
(up to 10 µg) and CIP (0.01 unit/pmol of protruding 5’ termini, 1 unit/pmol of blunt or
. recessed 5’ termini) in 10 mM Tris HCl, pH 8.3, 1 mM MgCl2, 1 mM ZnCl2 buffer. The
reaction mixture was diluted to 100 µL in TE buffer and the CIP was removed by extraction with 1:1 mixture of phenol/chloroform (3x), chloroform (2x), followed by ethanol precipitation.
7.6.4 Ligation of DNA. All ligation reactions were carried out in 20 mM Tris.HCl, pH
7.6, 5 mM DTT, 0.5 mM ATP, 50 µg/mL BSA buffer with T4 DNA ligase at 12-15 ◦C for 4-20 hr. The concentrations of DNA varied from 2 µg/mL (sticky-end ligation) to 30
µg/mL (blunt-end ligation). The ratio of insert DNA/vector DNA varied from 2:1
(restriction fragment to dephosphorylated vector DNA) to 20:1 (short unphosphorylated synthetic duplex to phosphorylated vector). The concentrations of T4 DNA ligase varied from 5 (sticky-end ligation) to 100 (blunt-end ligation) Weis units/mL. The reaction volume was 20-40 µL. The resulting solution was stored at -20 ◦C until used for
transformation.
7.6.5 Transformation. Competent cells in competence solution B were prepared as described previously and dispensed into 100 µL aliquots on ice. For transformations with purified supercoiled plasmid DNA, three dilutions of plasmid DNA (approx. 100 ng, 10 ng, and 1 ng of plasmid in 1-10 µL of TE buffer) were added each to an aliquot of
148 competent cells in a sterile microcentrifuge tube. For transformations with plasmid DNA
from mutagenesis synthesis reactions or ligation reactions, 10 µL, 1 µL, and 1 µL of 10 fold-dilution of the reaction mixture (usually containing 1-10 µg/mL DNA) were added to aliquots of competent cells. The DNA/cell suspensions were gently mixed and kept on ice for 30-35 min, after which the tubes were placed in a 37-42 ◦C heat-block for 3 min.
The tubes were centrifuged for 15 sec in a microcentrifuge and the supernatants were
carefully withdrawn with a pipette. The cell pellet was gently resuspended in 1 mL of
LB medium and incubated at 37 ◦C for 30-60 min. The cells were pelleted again by
centrifugation for 15 sec and resuspended in 100 µL of LB medium. This culture was evenly spread onto an LB plate impregnated with the appropriate antibiotic(s). The plates were inverted and incubated at 37 ◦C for 10-16 hr.
7.6.6 Small-Scale Preparation of Plasmid DNAs. Typically this was performed by
isolating a single E. coli colony from the transformation LB plate and inoculating 5 mL
of LB media with the appropriate antibiotic(s). This method was adapted from that of
Holmes and Quigley (162). After inoculation with aeration at 37 ◦C for 16-24 hrs, cells
were pelleted by centrifugation (5000 rpm, Sorvall SS-34 rotor, 5 min). The liquid
supernatant was removed and the cell pellet was resuspended by vigorous vortexing in
100 µL of ice cold Solution I (50 mM glucose, 25 mM Tris.HCl, pH 8.0, 10 mM EDTA).
200 µL of freshly prepared Solution II (0.2 N NaOH, 1% SDS) was added to the suspension and mixed by inverting 5-6 times and stored on ice for 2-5 min. 150 µL of ice-cold Solution III (60 mL 5 M KOAc, 11.5 mL glacial acetic acid, 28.5 mL ddH2O)
was then added and mixed by vortexing in an inverted position for 10 sec and returned to
149 ice for 3-5 min. The lysed cell suspension was centrifuged for 5 min in a microcentrifuge
at 13,000 rpm. The supernatant was transferred into a fresh tube and washed with 400
µL of 1:1 phenol:chloroform mixture. The solution was centrifuged at 13,000 rpm for 2
min. The top aqueous layer was transferred to another fresh tube. The DNA was
precipitated by adding 2 volumes of ethanol and centrifuged at 4 ◦C for 5 min. The
pelleted DNA was dried and redissolved in 50 µL TE and stored at -20 ◦C.
7.6.7 Mutagenesis. Site-directed mutagenesis was carried out by QuickChange Site-
Directed Mutagenesis Kit (Stratagene). Reactions were carried strictly as directed by the
manufacture protocol. Mutagenesis primers (35-40 nt) were ordered and synthesized by
IDT DNA Technologies. Pfu DNA polymerase was from Stratagene. Reactions
typically were carried out in 50 µL PCR reaction volume (thin wall) by adding 5-50 ng
vector dsDNA template, 125 ng primers, 0.5 mM dNTP mix, and 2.5 unit Pfu turbo DNA
polymerase, and adjust volume with ddH20 to get 50 µL. Cycling temperatures were
performed as followed: 16-18 cycles of 94 ◦C for 30 s/55 ◦C for 1 min/72 ◦C for 2 min per
1 kbp of plasmid. Plasmids typically contained about 6 kbp and required 12 min at the 72
◦C cycle. At the completion of the cycle, 10 units of Dpn I was added to the mutagenesis mixture and incubated at 37 ◦C for 1 hr. Dpn I treatment was required to remove the
parental or wild-type plasmid. The reaction mixture was then transformed into E. coli as
described previously.
150 7.6.8 Sequencing. DNA sequencing was performed by the dideoxy chain-termination
method derived from Sanger (163). Sequencing was performed at the Plant Genomic
Facility at the Ohio State University.
7.7 Purification of Deformylase (wild type from E. coli).
Q-Sepharose buffer A: 20 mM K/PO4, 10 mM NaCl, pH 8.0.
Q-Sepharose buffer B: 20 mM K/PO4, 1.0 M NaCl, pH 8.0.
Phenyl Sepharose buffer A: 20 mM K/PO4, 10 mM NaCl, pH 7.0.
Phenyl Sepharose buffer B: 20 mM K/PO4, 10 mM NaCl, 1.7 M (NH4)2SO4, pH 7.0.
Lysis buffer: 180 mL Q-Sepharose buffer A, 20 mL 10% Triton X-100, 1.0 g protamine
sulfate, and stir to dissolve completely.
7.7.1 Cell growth and induction. E. coli (DE3) cells (from a single colony from LB
plate) containing the appropriate plasmid (i.e. pET22b-Def) were used to inoculate an
overnight 100 mL culture containing 75 mg/L of ampicillin. The flask was aerated at 37
◦C at a shaking speed of 250 rpm. The overnight culture was used to inoculate the 6 L
LB media (proportioned into 6 flasks) by adding 1 mL of 75 mg/mL ampicillin stock
solution and 10 mL overnight culture to each flask. The flasks were shaken at 250 rpm
and the OD600 was monitored. When the reading reached a value 0.5-0.6 (typically 2-2.5
hours), 1 mL of 100 mM IPTG stock solution was added to each flask and the
temperature was lowered to 30 ◦C with a shaking speed of 200 rpm. Cell induction was carried out for 4 hr and harvested by centrifuging at 5000 rpm (Sorvall GS-3 rotor) 5 min at 4 ◦C. The cell pellets were combined.
151 7.7.2 Q-Sepharose purification. A Q-Sepharose column can be prepared as follows.
A XK type or C type column from Pharmacia can be packed with Q-Sepharose to a bed volume of about 20-25 mL. The distance between the bottom of the adapter/plunger and
the top of the bed should be kept to a minimum. The column after packing is washed
with at least 2 bed volumes of Q-Sepharose buffer B using the FPLC (flow rate 2-4
mL/min). A stable baseline form the FPLC detector should be obtained by this. The column can now be equilibrated with 3 bed volumes of Q-Sepharose buffer A at 2-4 mL/min until a stable baseline was observed. The sample was loaded onto the column with a Superloop at a flow rate of 2 mL/min. Once the loading was done, the column was
washed with Q-Sepharose buffer A until the baseline was stabilized. A gradient was used
0%-37.5% Q-Sepharose buffer B in a volume of 150 mL at 2 mL/min. 5-mL fractions
were collected. The deformylase eluted over a wide range. A 12% SDS-PAGE was run
to determine the deformylase fractions, which were combined for (NH4)2SO4.
7.7.3 Cell Lysis. The cell pellets were transferred to a clean 250 mL beaker by a clean spatula. 100 mL of Lysis buffer was added to the pellet and stir to suspend the cells.
After the cells have been resuspended, 200 µL of 10 mg/mL trypsin inhibitor, 200 µL of
10 mg/mL leupeptin, 2 mL of 1 mg/mL pepstatin A, 5 mg phenyl-methyl-sulfonyl fluoride, and 20 mg of chicken egg white lysozyme. The lysis solution was stirred with a magnetic bar (1 inch) for 30 min in a cold room and centrifuged for 30 min at 15,000 rpm
(Sorvall SS-34 rotor) at 4 ◦C. The supernatant was transfer into a clean beaker and loaded
onto a Q-Sepharose column.
152 7.7.4 Ammonium sulfate precipitation. The fractions from the Q-Sepharose
purification (usually 40-50 mL) were poured into a clean 100 mL beaker. A 0% to 80%
(NH4)2SO4 precipitation was done. A stir bar was used to stir the solution while adding
the ammonium sulfate slowly. The addition was carried out for 30 min. After all of the
solid had been added, the solution was stirred for an additional 10 min to ensure complete
deformylase precipitation. The (NH4)2SO4 salt protein pellet was recovered by
centrifugation at 15,000 rpm (Sorvall SS-34 rotor) at 4 ◦C for 15 min.
7.7.5 Phenyl Sepharose purification. A prepacked Pharmacia 16/10 HiLoad Phenyl
Sepharose column was used. The column was washed with Phenyl Sepharose buffer A until a stable baseline was observed (3-4 column volumes) and equilibrated with 3 column volumes of Phenyl Sepharose buffer B. The precipitated deformylase was dissolved in a minimum volume of 70% buffer B and 30% buffer A mixture (10-15 mL).
After dissolving the solution, an Acrodisc (0.45 micron) filter was used to remove the insoluble contaminants. The clear solution was loaded onto the column by a Superloop and washed with 50-60 mL buffer B before elution. A linear gradient was carried out from 100% to 0% Phenyl Sepharose buffer B in 200 mL at 4 mL/min. 4-mL fractions were collected. Around 50% Phenyl Sepharose buffer B, Zn-Deformylase began to elute.
Peak two immediately followed contained Fe-Deformylase.
The relative size of the two peaks varied from one purification to another. The
location of the peaks (% buffer B) may also vary. A specific activity assay of peak one
and two is done by measuring the initial rate with saturating concentrations of f-ML-
pNA. Usually, peak two is about 20-50 times more active than peak one. Often, the
153 resolution between the two peaks is not complete. In such cases, the earlier fractions of
peak one and the later fractions of peak two were saved.
7.7.6 Storage. Depending on the intended use of the protein, storage conditions may
vary. For enzymatic assays, it was often done by adding glycerol (33% final
concentration) to the protein solution. The mixture was aliquoted into 50 µL samples and quick freeze with liquid nitrogen. The frozen samples were stored at -80 ◦C. For other
purposes (e.g. mass spectrometry, ICP analysis) it may be necessary to concentrate the
protein using a Centriprep 30 for desalting and buffer exchange. The Pharmacia
Desalting HR column was used for this purpose.
7.7.7 Protein Quantitation. Protein concentrations were determined by measuring the
UV absorbance at 280 nm, assuming 0.93 mg/mL per OD280 (164). The concentrations
were also confirmed by Bradford method using bovine serum albumin (BSA) as standard.
For E. coli PDF, the concentration obtained from Bradford method was readjusted by a
factor of 0.56 (actual concentration = Bradford x 0.56).
7.8 Purification of Substituted Zn(II)-, Fe(II)-, and Co(II)-PDF. This procedure
was used to purify the non-histagged PDF.
7.8.1 Cell Growth. BL21(DE3) E. coli cells were transformed with pET22b-def DNA
construct. All media and buffers were prepared using metal free water (Millipore
system). Cell culture was carried out in minimal medium containing 2.5 g/L D-(+)-
glucose (Sigma), 5 g/L casamino acids (Difco), 10.8 g/L K2HPO4, 5.5 g/L KH2PO4, 1.0
154 g/L ammonium sulfate, 2 mg/L thiamine (Sigma), 1 mg/L (+)-biotin (Aldrich) and
. . supplemented with metals 35 mg/L MgSO4 7H2O, 10 mg/L CaCl2 2H2O, 4.7 mg/L
. . MnCl2 4H2O, 4 mg/L H3BO3, 1.5 mg/L (NH4)6Mo7O24 4H2O, 0.45 mg/L CuSO4, and 75 mg/mL ampicillin. Growth was performed at 37ºC until OD600 reached ~0.9. Induction
of protein expression was carried out at 30ºC by adding 100 µM isopropyl-β-D-
. thiogalactopyranoside and 100 µM (ZnCl2, CoCl2, or Fe(NH4)2(SO4)2 6H2O). Cells were
harvested after 12-16 hours.
7.8.2 Buffers: Buffer A: 20 mM Mes, pH 6.0, 10 mM NaCl. Buffer B: 20 mM Mes,
pH 6.0, 1.0 M NaCl. Buffer C: 20 mM Tris pH 8.0, 10 mM NaCl. Buffer D: 20 mM Tris,
pH 8.0, 1.0 M NaCl. Buffer E: 20 mM potassium phosphate, pH 7.0, 10 mM NaCl, 1.7 M
ammonium sulfate. Buffer G: 20 mM potassium phosphate, pH 7.0, 10 mM NaCl.
7.8.3 Protein Purification. Cell pellet was lysed in buffer A containing 1% (v/v)
Triton X-100, 0.5% (w/v) protamine sulfate, 2 mg trypsin inhibitor, 5 mg PMSF, and 5 mg chicken egg white lysozyme. The suspension was incubated for 30 minutes at 4ºC and sonicated 5 x 10 s pulses. The solution mixture was centrifuged at 15,000 rpm, Sorval
SS-34 rotor. The supernatant was loaded onto SP-Sepharose (Sigma) column (8.0 x 3.0 cm), preequilibrated in buffer A. Elution was performed with linear gradient from 10-
1000 mM NaCl with buffer B. Fractions containing deformylase activity was loaded onto Q-Sepharose (Pharmacia) column (8.0 x 3.0 cm), preequilibrated in buffer C.
Elution was performed with linear gradient from 10-1000mM NaCl with buffer D.
Deformylase active fractions were pooled and adjusted with solid ammonium sulfate to a
155 final concentration of 1.7 M and loaded onto phenyl-Sepharose (Pharmacia) column
(HiLoad 16/10), preequilibrated in buffer E. Protein elution was performed linearly in reversed salt concentration from 1.7-0 M ammonium sulfate with buffer F. Peptide deformylase fractions were pooled and concentrated. Protein concentration was determined by Bradford assay using bovine serum albumin as standard and corrected by a factor of 0.56 (actual concentration = Bradford x 0.56). Metal content was determined by
ICP-mass spectrometer.
7.8.4 Purification of His-tagged PDF. Very often the deformylase protein was expressed as PDF-H6. The H6 denoted six histidine residues tagged at the C-terminus of
PDF. The tag had been shown to not interfere with PDF activity. The cell growth culture and protein expression were carried similarly as mentioned previously.
Buffer A: 25 mM Tris.HCl, 500 mM NaCl, 5 mM imidazole, pH 7.9.
Buffer B: 25 mM Tris.HCl, 500 mM NaCl, 60 mM imidazole, pH 7.9.
Lysis buffer: 180 mL buffer A, 20 mL 10% Triton X-100, 1.0 g protamine sulfate, and
stir to dissolve completely. 200 µL of 10 mg/mL trypsin inhibitor, 200 µL of 10
mg/mL leupeptin, 2 mL of 1 mg/mL pepstatin A, 5 mg phenyl-methyl-sulfonyl
fluoride, and 20 mg of chicken egg white lysozyme were added.
The cell pellet (from 6-L culture, as previously described) was resuspended in
100 mL of Lysis buffer. Cell lysis was carried out as mentioned above. The supernatant was loaded onto 10 mL Talon resin (Clontech) preequilibrated in buffer A. The column was washed with 100 mL of the same buffer. Protein elution was carried by adding 100 mL of buffer B. The PDF-H6 often eluted within 50-60 mL. For the Co-PDF-H6, the
156 protein was pinkish. Zinc- and iron-PDF were colorless. The purification of the Fe-PDF-
H6 required 0.5 mM TCEP in all of the buffers, making sure that the buffers were degassed by reduced pressure. The eluted protein often was very homogenous. The imidazole in the buffer was removed by either a fast desalting column (Pharmacia) or dialysis overnight in 4-L of 20 mM Hepes, 150 mM NaCl, pH 7.0. The identity and ratio of the metal content were determined by ICP-mass spectrometry at the Universtiy of
Georgia. The protein was stored as 100-µL aliquots in 33% glycerol and without glycerol (i.e. for metal analysis and mass spectrometry). The 33% glycerol protein cannot be used for metal analysis due to a high metal ion contamination found in the glycerol stock.
157
APPENDIX
Escheria Coli Peptide Deformylase DNA and Protein Sequence: atgtcagttttgcaagtgttacatattccggacgagcggcttcgcaaagttgctaaaccg M S V L Q V L H I P D E R L R K V A K P 20 gtagaagaagtgaatgcagaaattcagcgtatcgtcgatgatatgttcgagacgatgtac V E E V N A E I Q R I V D D M F E T M Y 40 gcagaagaaggtattggcctggcggcaacccaggttgatatccatcaacgtatcattgtt A E E G I G L A A T Q V D I H Q R I I V 60 attgatgtttcggaaaaccgtgacgaacggctggtgttaatcaatccggagcttttagaa I D V S E N R D E R L V L I N P E L L E 80 aaaagcggcgaaacaggcattgaagaaggttgcctgtcgatccctgaacaacgtgcttta K S G E T G I E E G C L S I P E Q R A L 100 gtgccacgcgcagagaaagttaaaattcgcgccctggaccgcgacggtaaaccatttgaa V P R A E K V K I R A L D R D G K P F E 120 ctggaggcagatggtctgttagccatctgtattcagcatgagatggatcacctggtcggc L E A D G L L A I C I Q H E M D H L V G 140 aaactgtttatggattatctgtcaccgctgaaacaacaacgtattcgtcagaaagttgaa K L F M D Y L S P L K Q Q R I R Q K V E 160 aaactggatcgtctgaaagcccgggcttaa K L D R L K A R A -
Plasmodium falciparum Peptide Deformylase DNA and Protein Sequence: atgttgatgtattattcacttttcctttttaatttaataatatgttgtaatgttacaagt M L M Y Y S L F L F N L I I C C N V T S 20 atttatggatatatacacaatgttagatcacttgaaccatatataaaaaatgatcagata I Y G Y I H N V R S L E P Y I K N D Q I 40 aaaaattatagtagtaatataaaacaaaagagaaaaggctctttatatttattaaaaaat K N Y S S N I K Q K R K G S L Y L L K N 60 gaaaaggatgagataaaaatcgtcaagtacccggaccctatattaaggcgacgaagtgaa E K D E I K I V K Y P D P I L R R R S E 80 gaagtcacaaattttgatgataatttgaagagagttgtgagaaaaatgtttgatattatg E V T N F D D N L K R V V R K M F D I M 100 tacgagagcaaaggtattggtttgtctgcaccacaagtaaatataagcaaacgaattatt Y E S K G I G L S A P Q V N I S K R I I 120 gtatggaatgcattatatgaaaaaagaaaagaagaaaatgaacgaatatttattaatccg V W N A L Y E K R K E E N E R I F I N P 140 tccatagtagaacagagtctagttaaattaaaattaatagaaggatgtttatcatttcct 158 S I V E Q S L V K L K L I E G C L S F P 160 ggaatagaaggaaaagttgaacgacctagtatagtatctatatcatattatgatattaat G I E G K V E R P S I V S I S Y Y D I N 180 ggatataaacatttaaaaattttgaaaggtatacattctagaatatttcaacatgaattt G Y K H L K I L K G I H S R I F Q H E F 200 gatcatcttaatggtacattatttattgataaaatgacacaagtcgataaaaaaaaagta D H L N G T L F I D K M T Q V D K K K V 220 agaccaaaacttaacgagctaattagggattataaggctactcactcagaagaaccagcc R P K L N E L I R D Y K A T H S E E P A 240 ctataa L -
Homo sapiens Peptide Deformylase DNA and Protein Sequence: atggcccggctgtggggcgcgctgagtctttggccactgtgggcggccgtgccgtggggc M A R L W G A L S L W P L W A A V P W G 20 ggggcggcagccgtcggtgtccgggcttgcagctccacggccgccccggacggcgtcgag G A A A V G V R A C S S T A A P D G V E 40 ggcccggcgctgcggcgctcctattggcgccacctgaggcgtctggtgctgggtcctccc G P A L R R S Y W R H L R R L V L G P P 60 gaaccgccgttctcgcacgtgtgccaagtcggggacccggtgctgcgcggcgtggcggcc E P P F S H V C Q V G D P V L R G V A A 80 ccggtggagcgggcgcagctaggcgggcccgagctgcagcggctgacgcaacggctggtc P V E R A Q L G G P E L Q R L T Q R L V 100 caggtgatgcggcggcggcgctgcgtgggcctaagcgcgccgcagctgggggtgccgcgg Q V M R R R R C V G L S A P Q L G V P R 120 caggtgctggcgctggagctccccgaggcgctgtgtcgggagtgcccgccccgccagcgc Q V L A L E L P E A L C R E C P P R Q R 140 gcgctccgccaaatggagcccttccccctgcgcgtgttcgtgaaccccagcctgcgagtg A L R Q M E P F P L R V F V N P S L R V 160 cttgacagccgcctggtcacctttcccgagggctgcgagagcgtcgccggcttcctggcc L D S R L V T F P E G C E S V A G F L A 180 tgcgtgccccgcttccaggcggtgcagatctcagggctggaccccaatggagaacaggtg C V P R F Q A V Q I S G L D P N G E Q V 200 gtgtggcaggcgagcgggtgggcagcccgcatcatccagcacgagatggaccacctgcag V W Q A S G W A A R I I Q H E M D H L Q 220 ggctgcctgtttattgacaaaatggacagcaggacgttcacaaacgtctattggatgaag G C L F I D K M D S R T F T N V Y W M K 240 gtgaatgactaa V N D -
159
BIBLIOGRAPHY
1. Clark, B. F. C., and Marcker, K.A. (1966) J. Mol. Biol. 17, 394-406.
2. Marcker, K., and Sanger, K. (1964) J. Mol. Biol. 8, 835-840.
3. Dev, I. K., and Harvey, R.J. (1978) J. Biol. Chem 253, 4242-4244.
4. Petersen, H. U., Roll, T., Grunberg-Manago, M., and Clark, B.F.C. (1979) Biochem. Biophys. Res. Commun. 91, 1068-1074.
5. Sundari, R., Stringer, L., Schulman, L., and Maitra, U. (1976) J. Biol. Chem. 251, 3338-3345.
6. Fry, K. T., and Lamborg, M.R. (1967) J. Mol. Biol. 28, 423-433.
7. Adams, J. M. (1968) J. Mol. Biol. 33, 571-589.
8. Ben-Bassart, A., Baur, K., Chang, S.Y., Myambo, K., Boosman, A., and Chang, S. (1987) J. Bacteriol. 169, 751-757.
9. Chang, S. Y. P., McGary, E.C., and Chang, S. (1989) J. Bacteriol. 171, 4071- 4072.
10. Hirel, P. H., Schmitter, J.M., Dessen, P., Fayat, G., and Blanquet, S. (1989) Proc. Natl. Acad. Sci. USA 86, 8247-8251.
11. Mazel, D., Pochet, S., and Marliere, P. (1994) EMBO J. 13, 914-923.
12. Meinnel, T., and Blanguet, S. (1993) J. Bacteriol. 175, 7737-7740.
13. Newton, D. T., Creuzenet, C., and Mangroo, D. (1999) J. Biol. Chem. 274, 22143- 22146.
14. Guillon, J.-M., Mechulam, Y., Schmitter, J.M., Balnquet, S., and Fayat, G. (1992) J. Bacteriol. 174, 4294-4301.
15. Giglione, C., Pierre, M., and Meinnel, T. (2000) Mol. Microbiol. 36, 1197-1205.
16. Meinnel, T., and Blanquet, S. (1994) J. Bacteriol. 176, 7387-7390.
160 17. Margolis, P. S., Hackbarth, C.J., Young, D.C., Wang, W., Chen, D., Yuan, Z., White, R., and Trias, J. (2000) Antimicrob. Agents Chemother. 44, 1825-1831.
18. Meinnel, T. (2000) Parasitol. Today 16, 165-168.
19. Bianchetti, R., Lucchini, G., and Sartirana, M.L. (1971) Biochem. Biophys. Res. Comm. 42, 97-102.
20. Feldman, F., and Mahler, H.R. (1974) J. Biol. Chem. 249, 3702-3709.
21. Mahler, H. R., Dawidowicz, K., and Feldman, F. (1972) J. Biol. Chem. 247, 7439- 7442.
22. Mannhaupt, G., Beyreuther, K., and Michaelis, G. (1985) Eur. J. Biochem. 150, 435-439.
23. Velours, J., Esparza, M., Hoppe, J., Sebald, W., and Guerin, B. (1984) EMBO J. 3, 207-212.
24. Sebald, W., and Wachter, E. (1978) Energy Conservation in Biological Membranes.
25. Weiss, H. (1976) Biochim. Biophys. Acta 456, 291-313.
26. Tuschen, G., Sackman, U., Nehls, U., Haiker, H., Buse, G., and Weiss, H. (1990) J. Mol. Biol. 213, 845-857.
27. Polz, G., and Kreil, G. (1970) Biochem. Biophys. Res. Comm. 39, 516-521.
28. von Jagow, G., Engel, W. D., Schagger, H., Machleidt, W., and Machleidt, I. (1981) (al., P. e., Ed.) pp 149-161, Elsevier, Amsterdam.
29. Steffens, G. J., and Buse, G. (1976) Hoppe-Seyler's Z. Physiol. Chem. 357, 1125- 1137.
30. Steffens, G. J., and Buse, G. (1979) Hoppe-Seylaer's Z. Physiol. Chem. 360, 613- 619.
31. Fearnley, I. M., and Walker, J.F. (1986) EMBO J. 5, 2003-2008.
32. Yagi, T., and Hatefi, Y. (1988) J. Biol. Chem. 263, 16150-16155.
33. Hauska, G., Nitschke, W., and Herrmann, R.G. (1988) J. Bioenerg. Biomemb. 20, 211-228.
34. Schmidt, J., Herfuth, E., and Subramanian, A.R. (1992) Plant Mol. Biol. 20, 459- 465.
161 35. Shanklin, J., DeWitt, N.D., and Flanagan, J.M. (1995) Plant Cell 7, 1713-1722.
36. Giglione, C., Serero, A., Pierre, M., Boisson, B., and Meinnel, T. (2000) EMBO J. 19, 5916-5929.
37. Meinnel, T., and Blanquet, S. (1995) J. Bacteriol. 177, 1883-1887.
38. Rajagopalan, P. T. R., Datta, A., and Pei, D. (1997) Biochemistry 36, 13910- 13918.
39. Rajagopalan, P. T. R., Yu, X.C., and Pei, D. (1997) J. Am. Chem. Soc. 119, 12418-12419.
40. Groche, D., Becker, A., Schlichting, I., Kabsch, W., Schultz, S., and Wagner, A.F.V. (1998) Biochem. Biophys. Res. Comm. 246, 342-346.
41. Rajagopalan, P. T. R., and Pei, D. (1998) J. Biol. Chem. 273, 22305-22310.
42. Rajagopalan, P. T. R., Grimme, S., and Pei, D. (2000) Biochemistry 39, 779-790.
43. Becker, A., Schlichting, I., Kabsch, W., Schultz, S., and Wagner, A. F. V. (1998) J. Biol. Chem. 273, 11413-11416.
44. Becker, A., Schlichting, I., Kabsch, W., Groche, D., Schultz, S., and Wagner, A.F. (1998) Nat. Struct. Biol. 5, 1053-1058.
45. Chan, M. K., Gong, W., Rajagopalan, P.T.R., Hao, B., Tsai, C.M., and Pei, D. (1997) Biochemistry 36, 13904-13909.
46. Dardel, F., Ragusa, S., Lazennec, C., Blanquet, S., and Meinnel, T. (1998) J. Mol. Biol. 280, 501-513.
47. Meinnel, T., Blanquet, S., and Dardel, F. (1996) J. Mol. Biol. 262, 375-386.
48. Hao, B., Gong, W., Rajagopalan, P.T.R., Zhou, Y., Pei, D., and Chan, M.K. (1999) Biochemistry 38, 4712-4719.
49. Neu, H. C. (1992) Science 257, 1064-1073.
50. Levy, S. B. (2001) Clin. Infect. Dis. 33, S124–S129.
51. Olarte, J. (1983) APUA Newsletter 1, 3ff.
52. Watanabe, T. (1963) Bacteriol. Rev. 27, 87–115.
53. Weinstein, R. A. (2001) Emerg. Infect. Dis. 7, 188–192.
54. Anonymous. (2002) EARSS, Annual Report. 162 55. Cosgrove, S. E., Sakoulas, G., Perencevich, E., Schwaber, M., Karchmer, A., and Carmeli, Y. (2003) Clin. Infect. Dis. 36, 53–59.
56. Fridkin, S. K. (2001) Clin. Infect. Dis. 32, 108–115.
57. Hiramatsu, K. (1998) Drug Resist. Updat. 1, 135–150.
58. Baldwin, E., Harris, M. Yem, A., Wolfe, C., Vosters, A., Curry, K., Murray, R., Bock, J., Marshall, V., Cialdella, J., Merchant, M., Choi, G., and Deibel, M., Jr. (2002) J. Biol. Chem. 277, 31163-31171.
59. Bracchi-Ricard, V., Nguyen, K.T., Zhou, Y., Rajagopalan, P.T.R., Chakrabarti, D., and Pei, D. (2001) Arch. Biochem. Biophys. 396, 162-170.
60. Park, J., Moon, J., Kim, Jae, H., and Kim, E. (2005) Acta Crystallographica, Section F: Structural Biology and Crystallization Communications 61, 150-152.
61. Guilloteau, J.-P., Mathieu, M., Giglione, C., Blanc, V., Dupuy, A., Chevrier, M., Gil, P., Famechon, A., Meinnel, T., and Mikol, V. (2002) J. Mol. Biol. 320, 951- 962.
62. Zhou, Z., Song, X., Li, Y., and Gong, W. (2004) J. Mol. Biol. 339, 207-215.
63. Mazel, D., Coic, E., Blanchard, S., Saurin, W., and Marliere, P. (1997) J. Mol. Biol. 266, 939-949.
64. Apfel, C., Banner, D.W., Bur, D., Dietz, M., Hirata, T., Hubschwerlen, C., Locher, H., Page, M.G.P., Pirson, W., Rosse, G., and Specklin, J.-L. (2000) J. Med. Chem. 43, 2324-2331.
65. Chen, D. Z., Patel, D.V., Hackbarth, C.J., Wang, W., Dreyer, G., Young, D.C., Margolis, P.S., Wu, C., Ni, Z.-J., Trias, J., White, R.J., and Yuan, Z. (2000) Biochemistry 39, 1256-1262.
66. Broughton, B. J., Cahplen, P., Freeman, W.A., Warren, P.J., Wooldridge, K.R.H., and Wright, D.E. (1975) J. Chem. Soc., Perkin Trans. 1, 857.
67. Clements, J. M., Beckett, R.P., Brown, A., Catlin, G., Lobell, M., Palan, S., Thomas, W., Whittaker, M., Wood, S., Salama, S., Baker, P.J., Rodgers, H.F., Barynin, V., Rice, D.W., and Hunter, M.G. (2001) Antimicrob. Agents Chemother. 45, 563-570.
68. Chen, D. Z., Patel, D.V., Hackbarth, C.J., Wang, W., Dreyer, G., Young, D., Margolis, P.S., Wu, C., Ni, Z.-J., Trias, J., White, R., and Yuan, Z. (2000) 40th ICAAC, Toronto, Canada Abstract No. 2175.
163 69. Hu, Y.-J., Wei, Y., Zhou, Y., Rajagopalan, P.T.R., and Pei, D. (1999) Biochemistry 38, 643-650.
70. Lazennec, C., and Meinnel, T. (1997) Anal. Biochem. 244, 180-182.
71. Wei, Y., and Pei, D. (1997) Anal. Biochem. 250, 29-34.
72. Prescott, J. M., and Wilkes, S.H. (1976) Methods Enzylmol. 45, 530.
73. Guo, X.-C., Rajagopalan, P.T.R., and Pei, D. (1999) Anal. Biochem. 273, 298- 304.
74. Ishidoh, K., Muno, D., Sato, N., and Kominami, E. (1991) J. Biol. Chem. 266, 16312-16317.
75. Tran, T. V., Ellis, K.A., Kam, C.-M., Hudig, D., and Powers, J.C. (2002) Arch. Biochem. Biophys. 403, 160-170.
76. Ragusa, S., Mouchet, P., Lazennec, C., Dive, V., and Meinnel, M. (1999) J. Mol. Biol. 289, 1445-1457.
77. Arabaci, G., Guo, X.-C., Beebe, K.D., Coggeshall, K.M., and Pei, D. (1999) J. Am. Chem. Soc. 121, 5085-5086.
78. Kaiser, E. T., and Furlanetto, R.W. (1970) J. Am. Chem. Soc. 92, 6980-6982.
79. Nguyen, K. T., Hu, X., Colton, C., Chakratarti, R., Zhu, M.X., and Pei, D. (2003) Biochemistry 42, 9952-9958.
80. Bracchi-Ricard, V., Barik, S., DelVecchio, C., Doerig, C., Chakrabarti, R., and Chakrabarti, D. (2000) Biochem. J. 347, 255-263.
81. Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Lab. Press, Plainview, NY.
82. Altschul, S. F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. (1997) Nucleic Acids Res. 25, 3389-3402.
83. Nielsen, H., Engel, B., Brunak, S., and von Heijne, G. (1997) Protein Eng. 10, 1- 6.
84. Livingston, D. M., and Leder, P. (1968) Biochemistry 8, 435-443.
85. Takeda, M., and Webster, R. E. (1968) Proc. Natl. Acad. Sci. USA 60, 1487-1494.
86. Ragusa, S., Blanquet, S., and Meinnel, T. (1998) J. Mol. Biol. 280, 515-523.
164 87. Huntington, K. M., Yi, T., Wei, Y., and Pei, D. (2000) Biochemistry 39, 4543- 4551.
88. Inhibitor Synthesis was carried out by Dr. Xubo Hu.
89. Wiesner, J., Sanderbrand, S., Altincicek, B., Beck, E., and Jomaa, H. (2001) Trends Parasitol. 17, 7.
90. Kumar, A., Nguyen, K.T., Srivathsan, S., Ornstein, B., Turley, S., Hirsh, I., Pei, D., and Hol, W.G.J. (2002) Structure 10, 357-367.
91. McFadden, G. I., Reith, M., Munholland, J., and Lang-Unnasch, N. (1996) Nature 381, 482.
92. Wilson, R. J., Denny, P.W., Preiser, P.R., Roberts, K., Roy, A., Whyte, A., Strath, M., Moore, D.J., and Williamson, D.H. (1997) J. Mol. Biol. 261, 155-172.
93. Bodyl, A. (1999) Acta Protozool. 38, 31-37.
94. Waller, R. F., Reed, M. B., Cowman, A. F., and McFadden, G. I. (2000) EMBO J. 19, 1794-1802.
95. Varshavsky, A. (1996) Proc. Natl. Acad. Sci. U.S.A 93, 12142-12149.
96. Nicholls, A., Sharp, K.A., and Honig, B. (1991) Proteins 11, 281–296.
97. Giglione, C., Vallon, O., and Meinnel, T. (2003) EMBO J. 22, 13-23.
98. Meinnel, T., Mechulam, Y., and Blanquet, S. (1993) Biochimie 75, 1061-1075.
99. Pei, D. (2001) Emerging Ther. Targets 5, 23-40.
100. Giglione, C., and Meinnel, T. (2001) Emerging Ther. Targets 5, 41-57.
101. Chenchik, A., Diachenko, L., Moqadam, F., Tarabykin, V., Lukyanov, S., and Siebert, P.D. (1996) BioTechniques 21, 526- 534.
102. Anderson, S., Bankier, A.T., Barrell, B.G., de Bruijn, M.H., Coulson, A.R., Drouin, J., Eperon, I.C., Nierlich, D.P., Roe, B.A., Sanger, F., Schreier, P.H., Smith, A.J., Staden, R., and Young, I.G. (1981) Nature 290, 457-465.
103. Nielsen, H., Brunak, S., and von Heijne, G. (1999) Protein Eng. 12, 3-9.
104. Hauschild-Rogat, P. (1968) Mol. Gen. Genet. 102, 95-101.
105. Marasco, W. A., Phan, S.H., Krutzsch, H., Showell, H.J., Feltner, D.E., Nairn, R., Becker, E.L., and Ward, P.A. (1984) J. Biol. Chem. 259, 5430-5439.
165 106. Milligan, D. L., and Koshland, D.E., Jr. (1990) J. Biol. Chem 265, 4455-60.
107. Meinnel, T., Patiny, L., Ragasu, S., and Blanquet, S. (1999) Biochemistry 38, 4287-4295.
108. Wei, Y., and Pei, D. (2000) Bioorg. Med. Chem. Lett. 10, 1073-1076.
109. Jayasekera, M., Kendall, A. Shammas, R., Dermyer, M., Tomala, M., Shapiro, M., and Holler, T. (2000) Archives of Biochem. and Biophys. 381, 313-316.
110. Thorarensen, A., Douglas, M.R., Jr., Rohrer, D.C., Vosters, A.F., Yem, A.W., Marshall, V.D., Lynn, J.C., Bohanon, M.J., Tomich, P.K., Zurenko, G.E., Sweeney, M.T., Jensen, R.M., Nielsen, J.W., Seest, E.P., and Dolak, L.A. (2001) Bioorg. Med. Chem. Lett. 11, 1355-1358.
111. Roblin, P. M., and Hammerschlag, M.R. (2003) Antimicrob. Agents Chemother. 47, 1447-1448.
112. Coats, R. A., Lee, S.-L., Davis, K.A., Patel, K.M., Rhoads, E.K., and Howard, M.H. (2004) J. Org. Chem. 69, 1734-1737.
113. Gross, M., Clements, J., Beckett, R.P., Thomas, W., Taylor, S., Lofland, D., Ramanathan-Girish, S., Garcia, M., Difuntorum, S., Hoch, U., Chen, H., and Johnson, K.W. (2004) J. Antimicrob. Chemother. 53, 487-493.
114. Xue, C.-B., He, X., Roderick, J., DeGrado, W., Cherney, R., Hardman K., Nelson, D., Copeland, R., Jaffee, B., and Decicco, C. (1998) J. Med. Chem. 41, 1745- 1748.
115. Wei, C.-Q., Gao, Y., Lee, K., Guo, R., Li, B., Zhang, M., Yang, D., and Burke, T., Jr. (2003) J. Med. Chem. 46, 244-254.
116. Nguyen, K. T., Hu, X., and Pei, D. (2004) Bioorg. Chem. 32, 178-191.
117. Bertini, I., and Luchinat, C. (1994) in In Bioinorganic Chemistry (Bertini, I., Gray, H. B., Lippard, S. J., and Valentine, J. S., Ed.) pp 37-106, University Science Books, Sausalito, CA.
118. Morrison, J. F., and Walsh, C.T. (1988) Adv. Enzymol. 61, 201-301.
119. Wise, R., Andrews, J.M., and Ashby, J. (2002) J. Antimicrob. Agents Chemother. 46, 1117-1118.
120. Jones, R. N., and Rhomberg, P.R. (2003) J. Antimicrob. Chemother. 51, 157-161.
121. Murphy, P. M. (1994) Annu. Rev. Immunol. 12, 593-633.
122. Ye, R. D., and Boulay, F. (1997) Avd. Pharmacol. 39, 221-289. 166 123. Schiffmann, E., Corcoran, B.A., and Wahl, S.M. (1975) Proc. Natl. Acad. Sci. U.S.A. 72, 1059-1062.
124. Schiffmann, E., Showell, H.V., Corcoran, B.A., Ward, P.A., Smith, E., and Becker, E.L. (1975) J. Immunolo. 114, 1831-1837.
125. Carp, H. (1982) J. Exp. Med. 155, 264-275.
126. Greaves, D. R., and Channon, D.M. (2002) Trends in Immunology 23, 541-548.
127. Chadwick, V. S., Mellor, D.M., Myers, D.B., Selden, A.C., Keshavarzian, A., Broom, M.F., and Hobson, C.H. (1988) Scand. J. Gastroenterol. 23, 121-128.
128. Hobson, C. H., Roberts, E.C., Broom, M.F., Mellor, D.M., Sherriff, R.M., and Chadwick, V.S. (1990) J. Gastroenterol. Hepatol. 5, 32-37.
129. Chester, J. F., Ross, J.S., Malt, R.A., and Weitzman, S.A. (1985) Am. J. Path. 121, 284-290.
130. Nast, C. C., and LeDuc, L.E. (1988) Dig. Dis. Sci. 33, 50S-57S.
131. Sherriff, R. M., Broom, M.F., and Chadwick, V.S. (1992) Biochem. Biophys. Acta 1119, 275-280.
132. Broom, M. F., Sherriff, R.M., Tate, W.P., Collings, J., and Chadwick, V.S. (1989) Biochem. J. 257, 51-56.
133. Witheiler, J., and Wilson, D.B. (1972) J. Biol. Chem. 247, 2217-2221.
134. Yoshida, A., and Lin, M. (1972) J. Biol. Chem. 247, 952-957.
135. Jones, W. M., and Manning, J.M. (1988) Biochimica et Biophysica acta 953, 357- 360.
136. Tsunasawa, S., Narita, K, and Ogata, K. (1975) J. Biochem. 77, 89-102.
137. Termignoni, C., Freitas, J.O., Jr., and Guimaraes, J.A. (1986) Biochem. J. 234, 469-473.
138. Kobayashi, K., and Smith, J.A. (1987) J. Biol. Chem. 262, 11435-11445.
139. Radhakrishna, G., and Wold, F. (1986) J. Biol. Chem. 261, 9572-9575.
140. Ackerman, S. K., and Douglas, S.D. (1979) Biochem. J. 182, 885-887.
141. Aronson, J. H., and Lugay, J.C. (1969) Biochem. Biophys. Res. Comm. 34, 311- 314.
167 142. Fu, H., Dahlgren, C., and Bylund, J. (2003) Antimicrob. Agents Chemother. 47, 2545-2550.
143. Kobayashi, K., Lin, L.W., Yeadon, J.E., Klickstein, L.B., and Smith, J.A. (1989) J. Biol. Chem. 264, 8892 - 8899.
144. Polgar, L., and Patthy, A. (1992) Biochemistry 31, 10769-10773.
145. Sharma, K. K., and Ortwerth, B.J. (1993) Eur. J. Biochem. 216, 631-637.
146. Gade, W., and Brown, J.L. (1978) J. Biol. Chem. 253, 5012-5018.
147. Mitta, M., Miyagi, M., Kato, I., and Tsunasawa, S. (1998) J. Biochem. (Tokyo) 123, 924-931.
148. Unger, T., Nagelschmidt, M., and Struck, H. (1979) Eur. J. Biohem. 97, 205-211.
149. Pittelkow, S., Lindner, H., and Rohm, K.-H. (1998) Protein Expression and Purification 12, 269-276.
150. Giardina, T., Perrier, J., and Puigserver, A. (2000) Eur. J. Biochem. 267, 6249- 6255.
151. Grisolia, S., Reglero, A., and Rivas, J. (1977) Biochem. Biophys. Res. Comm. 77, 237-244.
152. Perrier, J., Durand, A., Giardina, T., and Puigserver, A. (2004) Comp. Biochem. and Phys., Part B: Biochem. and Mol. Biol. 138B, 277-283.
153. Schuchman, D. H., Jackson, C.E., and Desnick, R.J. (1990) Genomics 6, 149-154.
154. Cook, R. M., Burke, B.J., Buchhagen, D.L., Minna, J.D., and Miller, Y.E. (1993) J. Biol. Chem. 268, 17010-17017.
155. Mitta, M., Ohnogi, H., Yamamoto, A., Kato, I., Sakiyama, F., and Tsunasawa, S. (1992) J. Biochem. 112, 737-742.
156. Miyagi, M., Sakiyama, F., Kato, I., and Tsunasawa, S. (1995) J. Biochem. 118, 771-779.
157. Erlandsson, R., Boldog, F., Persson, B., Zabarovsky, E.R., Allikmets, R.L., and Jornvall, H. (1991) Oncogene 6, 1293-1295.
158. Jones, W. M., Scaloni, A., Bossa, F., Popowicz, A., Schneewind, O., and Manning, J.M. (1991) Proc. Natl. Sci. U.S.A. 88, 2194-2198.
159. Hu, X., Nguyen, K.T., Verlinde, C.L.M.J., Hol, W.G.J., and Pei, D. (2003) J. Med. Chem. 46, 3771-3774. 168 160. Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989) Cold Spring Harbor Laboratory, Cold Spring Harbor, NY Sections B.4 and B.5.
161. Laemmli, U. (1970) Nature 227, 680.
162. Holmes, M. D., and Quigley, M. (1981) Anal. Biochem. 114, 193.
163. Sanger, R., Nicklen, S., and Coulson, A. (1977) Proc. Natl. Sci. USA. 74, 5463.
164. Cuatrecasas, P., Fuchs, S., and Anfinsin, C.J. (1967) J. Biol. Chem 242, 1541.
169