FOLDING STUDIES OF OVINE PRION PROTEINS By Kai-Chun Chen A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirement for the degree of DOCTOR OF PHILOSOPHY Physiology 2010 ABSTRACT FOLDING STUDIES OF OVINE PRION PROTEINS By Kai-Chun Chen Several studies on scrapie in sheep have shown that the propensity for the conversion of a naturally occurring protein PrP from its normal cellular form (PrPC) to a virulent scrapie Sc form (PrP ), both in vitro and in vivo, correlates with susceptibility to the disease. Since Sc scrapie results from the PrPC-to-PrP conversion, the PrP Sc precursor occurring in PrP folding should play a key role in modulating disease occurrence. Susceptibility to classical scrapie correlates strongly with specific polymorphisms at positions 136, 154, and 171 in ovine prion proteins (ovPrPs). Therefore, I hypothesized that these polymorphisms affect scrapie susceptibility by modulating the structure and population of the PrP Sc precursor. This hypothesis led to two sub-hypotheses: First, folding and unfolding of ovPrPs proceed through an intermediate. Second, this intermediate is the Sc precursor of PrP . In this dissertation, I examined whether an intermediate state occurs in ovPrP (un)folding using a continuous-flow mixing method. Four PrP variants, comprising residues 94-233 of full-length ovPrP and correlated with differing susceptibilities to classical scrapie in sheep, were studied. An initial lag phase in refolding and unfolding kinetics indicates the presence of a native-like intermediate. I found that the relative populations and structural stability of the folding intermediates in these variants correlate with their propensities for classical scrapie. Variants susceptible to classical scrapie appear to have a larger population and higher structural stability in their intermediate states than do resistant variants. This could give susceptible variants more Sc opportunities to undergo the PrPC-to-PrP conversion and oligomerize. Therefore, I Sc argue that the observed folding intermediate is the precursor of PrP . A model for amyloid formation is proposed: Conformational folding of PrPC: N ↔ I ↔ U Assembly of PrP Sc Sc Sc oligomers: (PrP )n + I → (PrP )n+1 where N, I, and U represent native, intermediate, and unfolded states, respectively. Contrary to earlier studies on human PrP, I was unable to resolve any kinetic intermediate under refolding conditions within the dead-time of our instrument (~90 µs). Thus, I postulate that the folding kinetics of ovPrPs differs from the kinetics of human PrPs, which could reflect structural differences in their respective intermediates. Residues (136, 154, and 171) involved in genetic modulation are distant in the primary structure. Since ovPrP variants exhibit differing population and structural stability in their intermediate state, the polymorphism may modulate structural conversion through longrange interactions on the intermediate species. Consistent with this idea, a peptide model that is not involved in genetic modulation shows that local interactions in the vicinity of the disulfide bond do not suffice for the formation of a folding intermediate. To Bill and My Family iv ACKNOWLEDGEMTS First and foremost, I would like to thank my advisor Dr. William Wedemeyer, who brought me into this exciting field, and provided me with his total trust. I would also like to thank my collaborator Dr. Heinrich Roder, who is my life-saver. Without their continuous scientific and spiritual support, this work would never be done. I want to acknowledge my committee members, including Drs. Kathleen Gallo, David Kreulen, Vilma Yuzbasiyan-Gurkan, and Birgit Zipser for their guidance and encouragement. I will always remember the time they spent on my second preliminary exam. Thanks for making me talk and connect to the outside world. I appreciate the help from the MSU Physiology department, especially from Richard Brandt, Kim Crain, and Dr. Arthur Weber. They always tried to solve problems for me even before I recognized them. I acknowledge Terry Ball, Yi-Heng Sen, and Drs. Daniel Jones, Jon Kaguni, Lisa Lapidus, Min-Hao Kuo, Xue Li, Denise Mills, Jose-Luis Gallegos-Perez, Dean Shooltz, John Wang, Ming Xu, and Honggao Yan for their scientific support. I also thank supports from The Lim Pen-Yuan Cultural and Educational Foundation and my previous educators in Taiwan. v I want to express many thanks to my friends for going through the stress and depression with me along the whole PhD program. Ahlen, Chin-Mei, Chun-I, Feng, Guillermo, HuiWen, Joyce, Shuyi, Weining, Yi, Yvette, and many other people, I hope that absorbing those negative emotions has not caused permanent damage to your mental health. Last but not least, I want to send bunches of appreciation to my family. I thank my parents for their continuous loving nags. I acknowledge the warmth transmitted by the daily phone calls from my lovely sisters. I also appreciate my brother in-law Hari for his long-standing support. vi TABLE OF CONTENTS LIST OF TABLES………………………………………………………………………...x LIST OF FIGURES……………………………………………………………………....xi LIST OF ABBREVIATIONS…………………………………………………………...xiv CHAPTER 1. INTRODUCTION TO PRION DISEASES………………………………………………1 1.1. Protein-only hypothesis…………………………………….……………………...3 1.2. PrP27-30…………………………………………………………………………...6 Sc 1.3. Conformations of PrPC and PrP ………………………………………………....7 1.4. Theoretical models for structural conversion…………………………………….11 1.5. Prion strain and species barrier…………………………………………………...13 1.6. Biogenesis and trafficking of PrPC………………………………………….……15 1.7. Proposed functions of PrPC……………………………………………………….18 1.8. Scrapie in sheep…………………………………………………………………..19 1.9. Genetic modulation in sheep scrapie……………………………………………..20 1.10. Genetic modulation in human prion diseases…………………………………...21 1.11. Prion diseases and other protein misfolding disorders………………………….25 CHAPTER 2. INTRODUCTION TO PROTEIN FOLDING…………………………………….……..28 2.1. Equilibrium unfolding…………………………………………………………….28 2.2. Kinetic modeling of protein folding……………………………………………...30 2.3. Conformational states…………………………………………………………….31 2.3.1. The unfolded state………………………………………………………….33 2.3.2. The native state…………………………………………………………….34 2.3.3. The intermediate state……………………………………………………...35 2.3.4. The transition state…………………………………………………………36 2.4. Kinetic models of protein folding………………………………………………...36 2.4.1. Formation of native disulfide bonds……………………………………….36 2.4.2. Proline isomerization………………………………………………………37 2.4.3. Experimentally observed kinetic models for protein folding……………...38 2.5. Experimental approaches to study protein folding kinetics………………………40 2.5.1. Initiation of protein folding………………………………………………...41 2.5.2. Optical probes for folding detection……………………………………….45 2.5.3. Probe interactions formation along the folding pathway…………………..47 2.6. Theoretical aspects of protein folding…………………………………………….52 vii 2.6.1. Initiation of protein folding………………………………………………...52 2.6.2. Roles of intermediates in protein folding…………………………………..53 2.6.3. Theoretical models for protein folding…………………………………….54 2.6.4. Visualization of protein folding………………………………………...….56 CHAPTER 3. EXPERIMENTAL DESIGNS…………………………………………………………...62 3.1. Specific aim one: Assay for an ovPrP folding intermediate……………………...62 3.1.1. The major gap in knowledge……………………………………………….62 3.1.2. Hypothesis………………………………………………………………….63 3.1.3. Methods…………………………………………………………………….63 3.2. Specific aim two: Establish the relationship between (un)folding and susceptibility of ovPrP variants to classical scrapie……………………………………………..64 3.2.1. Major gap in knowledge…………………………………………………...64 3.2.2. Hypothesis………………………………………………………………….67 3.2.3. Methods…………………………………………………………………….68 3.3. Role of the disulfide bond in stabilizing the folding intermediate………………..68 3.3.1. Major gap in knowledge…………………………………………………...68 3.3.2. Hypothesis………………………………………………………………….68 3.3.3. Methods…………………………………………………………………….71 CHAPTER 4. EFFECTS OF POLYMORPHISMS ASSOCIATED WITH SCRAPIE SUSCEPTIBILITY ON THE FOLDING KINETICS OF OVINE PRION PROTEINS..74 4.1. Introduction……………………………………………………………………….74 4.2. Experimental procedure…………………………………………………………..77 4.2.1. Protein preparation and purification……………………………………….77 4.2.2. Tryptophan fluorescence reporter………………………………………….82 4.2.3. Continuous-flow measurements……………………………………………87 4.2.4. Exponential fitting…………………………………………………………95 4.2.5. Data analysis……………………………………………………………….96 4.3. Results…………………………………………………………………………...101 4.3.1. Unfolding of ovPrPs……………………………………………………...101 4.3.2. Refolding of ovPrPs………………………………………………………103 4.4. Discussion……………………………………………………………………….115 Sc 4.4.1. Kinetic intermediate is a plausible PrP precursor………………………115 4.4.2. Intermediates observed under refolding conditions………………………117 Supplemental materials………………………………………………………………119 CHAPTER 5. ROLE OF THE DISULFIDE BOND IN STABILIZING THE FOLDING INTERMEDIATE………………………………………………………………………131 5.1. Introduction……………………………………………………………………...131 5.2. Materials and methods…………………………………………………………..134 5.2.1. Materials………………………………………………………………….134 5.2.2. Construction of peptide models…………………………………………..134 viii 5.2.3. Preparation of peptide models……………………………………………134 5.2.4. Hydrogen/deuterium exchange…………………………………………...137 5.2.5. Circular dichroism (CD) spectroscopy…………………………………...140 5.3. Results…………………………………………………………………………...140 5.4. Discussion…………………………………………………………………….…146 5.5. Future directions………………………………………………………………...147 5.5.1. Preparation of GST-tagged peptides...……………………………………148 5.5.2. Future work……………………………………………………………….150 CONCLUSIONS AND FUTURE DIRECTIONS……………………………………..151 APPENDIX……………………………………………………………………………..158 REFERENCES…………………………………………………………………………160 ix LIST OF TABLES Table 1.1. Prion diseases in humans and animals…………………………………………2 Table 1.2. A list of protein misfolding diseases………………………………………….27 Table 4.1. Kinetic parameters for the folding of ovPrPs……………………………….113 x LIST OF FIGURES Figure 1.1. Prion amplification by PMCA………………………………………………...5 Figure 1.2. PrP isoforms…………………………………………………………………..7 Figure 1.3. Ribbon representation of the structure of ovPrP (residue 126-233)…………..8 Sc Figure 1.4. Comparison of structural models for PrP and amyloid……………………10 Sc Figure 1.5. Two models for conformational conversion of PrPC to PrP ……………….13 C Figure 1.6. Biosynthesis and trafficking of PrP ………………………………………...17 C Figure 1.7. Human PrP and the mutations linked to familial prion diseases…………...23 Figure 2.1. Example for a simple chemical reaction involving formation of an intermediate………………………………………...…………………………………….32 Figure 2.2. Schematic illustration of cooperativity between two interactions in a protein.…………………………………………………………………………………...35 Figure 2.3. Isomeric states of proline peptide bond and its isomerization reaction……...38 Figure 2.4. Three possible folding schemes account for an experimentally observed folding intermediate……………………………………………………………………...40 Figure 2.5. Mixing methods for stopped- and continuous-flow experiments……………44 Figure 2.6. Far-UV CD spectra associated with various types of secondary structure….47 Figure 2.7. Schematic illustration for pulsed hydrogen exchange method……………...50 Figure 2.8. Free energy diagram for Ф-value analysis of transition and intermediate states……………………………………………………………………………………...52 Figure 2.9. Illustration of three classical folding mechanisms…………………………..56 Figure 2.10. Schematic representation of the free energy surface calculated for the folding of a 27-mer………………………………………………………………………58 xi Figure 2.11. Simplified schematic diagram for illustrating different types of free energy surface……………………………………………………………………………………60 Sc Figure 3.1. Conversion from PrPC to PrP through disulfide polymerization in the domain swapping model…………………………………………………………………66 Figure 3.2. Positions of the sole disulfide bond and model peptides in ovPrP…………..70 Figure 4.1. Ribbon representation of the structure of ovPrP (residue 126-233)…………75 Figure 4.2. SDS-PAGE and Western blotting show the expression of ovPrP in E. coli...80 Figure 4.3. Monomeric ovPrP(ALRQ) eluted from Ni-NTA column…………………...80 Figure 4.4. MALDI mass spectrum of the ALRQ variant……………………………….81 Figure 4.5. Concentration profiles of sedimentation equilibrium………………………..82 Figure 4.6. Primary structure of the wild-type ALRQ variant…………………………...84 Figure 4.7. Far-UV circular dichroism spectra of the wild-type ovPrP and the ovPrP(W102F/Y221W)………………………………………………………………….85 Figure 4.8. Representative equilibrium transition curve of ovPrP(W102F/Y221W) monitored by tryptophan fluorescence…………………………………………………...86 Figure 4.9. Fluorescence emission of native and unfolded ovPrP(W102F/Y221W)……86 Figure 4.10. Sample delivery and detection in the continuous-flow method……………88 Figure 4.11. Time course of the quenching reactions between NATA and NBS of various concentrations……………………………………………………………………………92 Figure 4.12. Combination of ALRQ refolding kinetic traces taken at flow rates of 0.9 and 1.8 ml/s…………………………………………………………………………………...95 Figure 4.13. Denaturant dependence of the folding and unfolding rates for a two-state (un)folding protein……………………………………………………………………….97 Figure 4.14. Denaturant-dependence of the folding and unfolding rates for a three-state (un)folding protein……………………………………………………………………….98 Figure 4.15. Unfolding kinetic traces taken for ovPrPs in the presence of various concentrations of GuHCl at pH 7, 15oC………………………………………………...102 xii Figure 4.16. Kinetic trace taken for ALRQ variant in the presence of 5.2 M GuHCl at pH 7 and 15oC………………………………………………………………………………103 Figure 4.17. Refolding kinetic trace taken for ALRQ variant in the presence of 1.45 M GuHCl at pH 5 and 15oC……………………………………………………………….105 Figure 4.18. Refolding kinetic traces taken for ovPrPs in the presence of various concentrations of GuHCl at pH 7, 15oC………………………………………………...106 Figure 4.19. Refolding kinetic traces taken for ovPrPs in the presence of various concentrations of GuHCl at pH 5, 15oC…………………………………………...........107 Figure 4.20. Refolding kinetic traces taken for ovPrPs in the presence of various concentrations of urea or GuHCl……………………………………………………….109 Figure 4.21. Chevron plots of ALRQ, ALHQ, ALRR, and VLRQ variants at pH 7, 15oC…………………………………………………………………………………….110 Figure S1. Amplitudes and rate constants from the best fit to refolding kinetic trace………………………………………………….………………………………….119 Figure S2. Amplitudes and rate constants from the best fit to refolding kinetic traces……………………………………………………………………………………128 Figure 5.1. Positions of the disulfide bond and the selected model peptides…………..132 Figure 5.2. Identification of P1 and P2…………………………………………………136 Figure 5.3. Mechanism of HDX reaction and its reaction rates plotted as a function of pH for polyalanine………………………………………………………………………….139 Figure 5.4. CD spectra of the five peptide models……………………………………..142 Figure 5.5. HDX reactions for the model peptides……………………………………..144 Figure 5.6. DNA encoding P1 and P2 were cloned into PGEX-4T-1………………….149 Figure 5.7. SDS-PAGE and Western blotting show the purified GST- P1 and GSTP2……………………………………………………………………………………….150 Figure C1. Schematic illustration for quenching reaction…...…………………………154 Figure C2. The positions of quenchers, Trp, and residues involved in sheep polymorphisms………………………………………………………………………….155 Figure C3. Quenching reactions of the Trp/Cys pair…………………………………...156 xiii LIST OF ABBREVIATIONS AD: Alzheimer’s disease Ala (A): Alanine ALS: Amyotrophic lateral sclerosis ANS: 8-Anilinonaphthalene-1-sulfonic acid Arg (R): Arginine Asp (D): Aspartic acid BSE: Bovine spongiform encephalopathy (mad cow disease) CD: Circular dichroism CI2: Chymotrypsin inhibitor 2 CJD: Creutzfeldt-Jacob disease CNS: Central nervous system CWD: Chronic wasting disorder Cys: Cystine (disulfide bond) ER: Endoplasmic reticulum fCJD: Familial CJD FFI: Fatal familial insomnia FSE: Feline spongiform encephalopathy Gln (Q): Glutamine GSS: Gerstmann-Straussler-Scheinker diseases GuHCl: Guanidine hydrochloride xiv HD: Huntington’s disease HDX: Hydrogen/deuterium exchange His: Histidine Iα: Intermediate state IPTG: Isopropyl β-D-1-thiogalactopyranoside Leu (L): Leucine LRS: Lymphoreticular system MALDI-TOF-MS: spectrometry Matrix-assisted laser-desorption/ionization time-of-flight mass Met (M): Methionine N: Native folded state NATA: N-acetyl tryptophan amide NBS: N-bromo-succinamide Ni-NTA: Nickel-nitrilotriacetic acid ORF: Open reading frame OvPrP: Ovine prion protein PD: Parkinson’s disease Phe (F): Phenylalanine PMD: Protein misfolding disorder Prion: The infectious agent that is believed to cause TSEs. It denotes to ‘small proteinaceous infectious particle’ Pro (P): Proline PrP: Prion protein C PrP : The normally occurring cellular form of a PrP xv Sc PrP : An abnormal form of a PrP that is involved in prion disease. Its conformation differs from PrPC, and its aggregate is associated with infectivity in prion diseases RP-HPLC: Reverse-phase high-performance liquid chromatography TCEP: Tris-(2-carboxyethyl) phosphine hydrochloride TMT: Transmissible mink encephalopathy Trp (W): Tryptophan TS: Transition state TSE: Transition state ensemble TSE: Transmissible spongiform encephalopathy Tyr (Y): Tyrosine Phe (F): Phenylalanine Pro (P): Proline U: Unfolded state UV: Ultraviolet Val (V): Valine xvi Chapter 1. Introduction to Prion Diseases Prion diseases, also termed transmissible spongiform encephalopathies (TSEs), are a group of fatal neurodegenerative diseases including scrapie in sheep and goats, bovine spongiform encephalopathy in cattle (mad cow disease), chronic wasting disorder (CWD) in deer and elk, fatal familial insomnia (FFI), Gerstmann-Straussler-Scheinker diseases (GSS), as well as Creutzfeldt-Jacob disease (CJD) in humans (Table 1). Based on the etiologies, TSE can be classified as sporadic, inherited, and acquired. The pathologic changes include vacuolation, neuronal death, and astrocytosis in the central nervous system (CNS)(1). Despite intensive studies, the means of treatment and diagnosis are still limited by our current knowledge of its pathogenesis. 1 Table 1.1. Prion diseases in humans and animals(2-3) Disease Host Etiology Kuru humans infection through cannibalism Iatrogenic CJD humans transmitted through neurosurgical procedures or administration of pituitary tissue extracts Variant CJD humans infection from bovine prions Familial CJD humans germline mutations FFI humans germline mutations Sporadic CJD humans somatic mutations or C spontaneous PrP -to-PrP Sc conversion (uncertain) Classical Scrapie sheep infection in genetically susceptible sheep Atypical Scrapie sheep sporadic (uncertain) BSE cattle infection with contaminated (bovine spongiform encephalopathy) meat and bone meal (MBM) TMT infection with prions from mink (transmissible mink encephalopathy) sheep or cattle FSE infection with contaminated cats (feline spongiform encephalopathy) CWD bovine tissues or MBM deer, elk (chronic wasting disease) 2 unknown 1.1. Protein-only hypothesis. The identity of TSE infectious agent had been puzzling for decades. Based on its resistance to ultraviolet (UV) irradiation, Alper and Griffith suggested that this agent is solely composed of proteins and might be able to self-replicate(4-5). However, further evidence did not appear until twenty years later. In the 1980s, Prusiner et al. isolated this agent with high purity, and reported three observations supporting the ‘protein-only hypothesis’: First, this agent is resistant to procedures that attack nucleic acids. Second, treatments that modify proteins can diminish scrapie infectivity. Third, this agent is smaller than a virus, a spiroplasma-like organism, or a parasite. The name ‘prion’ was coined to denote this agent as ‘a small proteinaceous infectious particle’(6). Purified prions were used to identify the cDNA clone encoding the entire open reading frame (ORF) of a prion protein (PrP)(7). A chromosomal gene encoding PrP was subsequently identified(8). The entire ORF of Prnp genes is contained within a single exon, suggesting that alternative splicing seems unlikely to explain the two PrP C isoforms(7-8). A later study suggested that the normal cellular form (PrP ) and the Sc virulent scrapie form (PrP ) share the same primary structure; therefore, the pathogenic conversion of PrP seems to involve either posttranslational modifications or conformational changes(9). Although earlier studies suggested that posttranslational modifications are not involved in the pathogenic conversion(10), emerging evidence indicates that covalent differences might exist in these two isoforms(11-12). By contrast, C Sc dramatic structural changes have been observed upon PrP to PrP conversion(13-15). 3 Substantial evidence supports the protein-only hypothesis. First, prion infectivity is reduced by antibody against PrP(16-18). Second, prions from inherited diseases are infectious to experimental animals(19). Third, mice lacking PrP C do not support Sc conversion to PrP (20). Fourth, the scrapie agent is able to induce propagation of infectivity in neuroblastoma cells(21-22). Since the protein-only hypothesis suggests that prion diseases result from the structural C Sc conversion of PrP to PrP (6), its final proof can be archived only if conversion of a pure PrPC to a form that causes prion diseases can be carried out in vitro. Caughey et al. have developed the first successful in vitro conversion assay, termed as cell-free Sc conversion methods. Although it generates misfolded proteins similar to PrP , these preparations are not infectious(23-24). Another important in vitro conversion system, termed as protein-misfolding cyclic amplification assay (PMCA), has been developed by Soto et al.. In this method, PrP Sc amplification is carried out via cycles of incubation and Sc has been proved to be infectious Sc sources; nevertheless, cofactors sonication (Figure 1.1)(25). The de novo generated PrP when brain homogenates were used as PrPC and PrP have been suggested to involve in this conversion process(25-27). A recent advance in this field is that structural conversion has been carried out using recombinant PrPC purified from E. coli. Wang et. al. have shown that prions with relatively high infectivity can be generated using recombinant PrPC in the presence of lipid and RNA in PMCA method(27). A very recent report provides compelling evidence for the protein-only hypothesis. Kim et al. have observed that PrP 4 Sc infectivity can be propagated using either the full-length or the C-terminal domain of a recombinant PrP C in the absence of cofactors in PMCA method(28). However, the de novo generated PrP Sc exhibits much lower infectivity, as compared to the previous preparation in the presence of lipid and RNA, suggesting that these cofactors facilitate, instead of determine, the structural conversion process(28). PrPC PrPSc Growing of oligomeric PrPSc sonication PrPSc PrPC PrPSc Growing of oligomeric PrPSc C Figure 1.1. Prion amplification by PMCA. PrP is first incubated with minute amount of PrP Sc to induce formation of PrP Sc polymers. Sonication is then applied to break the Sc newly formed PrP polymers to generate new growth ‘seeds’ for subsequent conversion. 5 Although PrP Sc is associated with infectivity(29), and aggregates thereof have been suggested to be neurotoxic(30-32), growing evidence has challenged the latter idea. First, 0/0 inoculation of prion does not cause neuronal death in PrP null (Prnp ) mice, indicating C that PrP expression is a prerequisite for prion-induced neurotoxicity(20, 33-34). Second, PrP Sc is barely or not detected in some prion diseases, and the pathological changes in the brain region are not always associated with PrP Sc accumulation(35-38). Third, PrP Sc accumulation can occur without developing neurotoxicity even in the presence of C PrP (39-40). 1.2. PrP27-30. PrP Sc exhibits partial resistance to proteinase K digestion. The proteinase-resistant core is composed of ~142 residues and has an apparent molecular Sc C mass of ~27-30 kDa; thus, it is designated as PrP27-30. Unlike PrP , PrP is sensitive to PK digestion(41) (Figure 1.2). 6 PRNP ORF N-linked oligosaccharide GPI anchor S S Octapeptide repeats Disulfide bond PHGGGWGQ PrPC (~209 amino acids) PrPSc (~209 amino acids) Proteinase K digestion PrP27-30 (~142 amino acids) Figure 1.2. PrP isoforms. 1.3. C Sc C Conformations of PrP and PrP . PrP is composed of an unstructured N- terminal domain and a globular C-terminal domain (residues ~120-233) of known structure, comprising three α-helices, a short two-stranded antiparallel β-sheet, and a disulfide bond (Cys182-Cys217) linking helices 2 and 3 (Figure 1.3)(42-46). Relatively Sc little is known about the structure of PrP , except that it appears to have significantly more β-sheet and slightly less α-helix (17-30% α-helix; 43-54% β-sheet) compared to PrPC (47% α-helix; 3% β-sheet)(13-15). 7 N’ 171R/Q Helix-2 Helix-1 C’ 136A/V Helix-3 154R/H 141L/F Figure 1.3. Ribbon representation of the structure of ovPrP (residue 126-233). Helix 1 is shown in green; helix 2 is shown in mauve; helix 3 is shown in blue. The antiparallel β-sheet is shown in yellow; the disulfide bridge is shown in gold. The residues involved in genetic modulation in sheep scrapie are shown in red. Sheep numbering is used here. Figure was drawn with MOLMOL(47). For interpretation of the references to color in this and all other figures, the reader is referred to the electronic version of this dissertation. Three structural models for PrP Sc have been proposed. (i) β-helical model (Figure 1.4.A). Based on electron crystallography and molecular modeling, Govaerts et al. proposed the trimeric β-helical model of PrP27-30(48). By refolding residues 89-174 into a left-handed β-helical fold(Figure 1.4.A1), PrP form a trimer as the fundamental unit of a fibril with helices 2 and 3 mostly preserved (Figure 1.4.A2)(48). To initiate a fibril, PrP 8 Sc trimers stack on one another through hydrogen bonds in the direction perpendicular to the βstrands, thus constructing a cross-β architecture (Figure 1.4.A3)(48). The C-terminal αC helices and the N-linked glycans provide an external coating similar to PrP (Figure Sc 1.4.A3), which might explain the absence of host immune response against PrP (48). Consistent with this model, an antibody study observed that secondary structure changes occur in α-helix 1, whereas α-helices 2 and 3 remain largely conserved during the PrPCto-PrP Sc conversion(49). (ii) Spiral model. (Figure 1.4.B) Based on molecular dynamics simulations, DeMarco et al. proposed that the core of the β-structure in the prion protofibril is composed of a three-stranded sheet E1-E3 (E1: 116-119; E2: 129-132; E3: 160-164), and an isolated strand E4 (135-140), with retention of all three native αhelices(50) (Figure 1.4.B1). Oligomerization occurs by docking PrP Sc monomers through E4 to E1 (Figure 1.4.B2); protofibrils grow in a spiral-like manner (Figure 1.4.B3)(50). (iii) Parallel and in-register β-structure model. (Figure 1.4.C) Contrast to the above two C models, Surewicz et al. proposed that PrP to PrP Sc conversion involves refolding of the entire α-helical domain (Figure 1.4.C1) based on the analyses of hydrogen/deuterium exchange and site-directed spin labeling(51). In this model, the β-sheet core is composed of residues ~160-220, which forms a single-molecule layer that stack on top of one another with parallel, in-register alignment of β-strands to form amyloid(51) (Figure 1.4.C2). 9 Sc Figure 1.4. Comparison of structural models for PrP and amyloid. (A) β-helical model. A1: Model for monomeric PrP27-30. Structural conversion involves residues 89C 174, whereas C-terminal helices of monomeric PrP remain intact. A2: Trimeric PrP2730 forms the fundamental unit for building up a fibril. A3: Two trimeric PrP27-30 are linked through hydrogen bonds. Glycans and C-terminal α-helices extend away from the center of the structure. Figure was adopted and modified from Govaerts, C. et al. (2004) Sc Proc. Natl. Acad. Sci. U S A. 101, 8342-8347. (B) Spiral model. B1: Monomeric PrP . All three native α-helices remain intact during structural conversion. B2: Propagation of a protofibril occurs by docking PrP monomers through E4 to E1. B3: A protofibril grows in a spiral-like manner. Figure was adopted and modified from DeMarco, M. et al. (2004) Proc. Natl. Acad. Sci. U S A. 101, 2293-2298. (C) Parallel and in-register β-structure Sc model. C1: Monomeric PrP . Structural conversion involves a major part of helices of a C monomeric PrP . C2: Individual monomers stack on top of one another to form amyloid. Figure was adopted and modified from Cobb, N.J. et al (2007) Proc. Natl. Acad. Sci. U S A. 104, 18946-18951. (A1) H3 (A3) Fibril growing direction (A2) H2 Residues 89-174 fold into β-helical structure H2 and 3 Glycans 10 Figure 1.4. (cont’d) (B1) (B2) (B3) Docking the monomers through E1 to E4 (oligomerization site) Long axis of the fibril (C1) (C2) Native disulfide Bond Monomer 1.4. Theoretical models for structural conversion. Two theoretical models have C been proposed for the PrP -to-PrP Sc conversion(1, 52). The refolding model suggests that C * the PrP is in equilibrium with a partially folded intermediate PrP (Figure 1.5). A high C Sc activation energy barrier prevents spontaneous conversion of PrP to PrP . When PrP Sc * Sc interacts with exogenously introduced PrP , it refolds under the effect of PrP , and adopts the PrP Sc conformation. In this model, the driving force for structural conversion Sc is monomeric PrP . However, this model has a chicken-and-egg problem. For example, it cannot explain the existence of PrP Sc found in patients who have never been exposed to 11 Sc exogenous PrP , as in inherited prion diseases. Proponents of the refolding model suggest that the concentration of PrP* for the mutant PrP is higher than that for the wildtype PrP, and is sufficient to initiate aggregation without a preexisting PrP Sc Sc seed, leading * to de novo formation of PrP . For illustration, the reaction may occur as PrP + PrP Sc → (PrP )2. In contrast to the refolding model, the nucleation model proposes that PrP Sc C itself is the precursor of PrP . PrP and PrP strongly favored by free energy. PrP PrP Sc Sc Sc are in a reversible equilibrium, with PrP * C C monomer is stable only when it is added onto a aggregate (Figure 1.5). In other words, the structural conversion is driven by Sc oligomeric PrP . This model suggests that infectivity is encoded in the highly ordered PrP Sc Sc aggregate, instead of monomeric PrP , since the latter exists ubiquitously(52). The key differences between these two models are: (i) whether a kinetic folding intermediate * Sc PrP is formed, (ii) the identity of the precursor to PrP , and (iii) the driving force for structural conversion. 12 Nucleation model (PrPSc)m (PrPSc)m+1 (PrPSc)m PrPSc (PrPSc)m+1 + PrPC PrP* + Refolding model C Sc Figure 1.5. Two models for conformational conversion of PrP to PrP . The refolding model (upper panel) indicates that a partially folded intermediate PrP* is the Sc precursor of PrP . This conversion process can be expressed as (PrP Sc Sc C ↔ PrP*) + C (PrP )m → (PrP )m+1. The nucleation model (lower panel) indicates that PrP itself is Sc the precursor of PrP . This conversion process can be expressed as (PrP Sc C Sc PrP ) Sc + (PrP )m → (PrP )m+1. 1.5. Prion strain and species barrier. One of the most puzzling phenomena is that prions exist in different strains that exhibit distinct neuropathology, incubation time, and specific neuronal target areas(52). Conformational stability(53), glycosylation patterns(54), and cleavage sites for proteinase K digestion(55-57) have been shown to differentiate strain types, suggesting that strain-specific information is encoded in PrP structure. 13 Sc Transmission of prions across species often delay the onset of disease, or prevent it altogether(1, 58-59). Such effects are usually reduced and finally stabilized in further passage(60). This phenomenon has been referred as the ‘species barrier’. Early studies argue that the species barrier is determined by the degree of the differences in the primary structure of donor and recipient PrPs. However, growing evidence suggests that the conformation of the donor prion strain and the primary structure of the recipient PrP are both important in determining the species barrier(58, 60). To explain the role of the primary structure of the host PrP in prion transmission, the ‘conformational selection hypothesis’ was proposed. In this model, the transmission barrier is determined by the conformation of the donor prion strain and the conformational constraints imposed by the primary structure of recipient PrP(58, 61). If the conformation of the donor PrP Sc is within the spectrum of the conformations allowed by the recipient primary structure, a low transmission barrier is expected(61-62). Conversely, if the primary structure of the recipient PrP does not favor to form the conformation of the donor prion, a high transmission barrier is expected. The passage can be stabilized only after a new strain showing structural compatibility has emerged(61-62). It is conceivable that cellular chaperone and environment could set different conformational constraints on the recipient PrPC, thus modulating the range of preferred conformation. Two theoretical models have been proposed to explain the emergence of a new strain. First, a new strain might be generated de novo during the intermolecular interaction 14 C Sc between the host PrP and the donor PrP (61-62). Second, the concept of ‘substrain’ has been proposed. A substrain is a collective of interconvertible prion conformations(63). A strain comprises a set of substrains, with one dominant PrP Sc conformation coexisting with other minor conformational states(63). When a prion inoculum composed of a diverse substrain population is introduced into a recipient animal, recipient PrPC may preferentially interact with a specific substrain and causes the dominant conformation to C shift to the PrP -favored substrain(61-62). A recent report has shown that endogenous cofactors can lead to the emergence of a new strain(63). Consistent with the substrain concept, strain selection has been observed to occur within a population of diverse substrains to adapt different cell types(63). 1.6. Biogenesis and trafficking of PrPC. The mammalian PrP gene (Prnp) encodes a protein of ~ 250 amino acids that contains an amino-terminal signal peptide (residues 122), several octapeptide repeats, a highly conserved central hydrophobic region (residues 111-134), and a C-terminal signal peptide (residues 231-254) for addition of a glycosylphosphatidylinositol (GPI) anchor(2). The primary translation product of Prnp is subject to cleavage of amino- and carboxylterminal signal peptides, addition of N-linked oligosaccharide chain at two positions (Asn181 and Asn197), formation of a disulfide bond, and attachment of a GPI anchor. The mature PrPC is then transported to the cell surface(2). 15 C It has been reported that PrP cycles continuously between the plasma membrane and an endocytic compartment through clathrin-dependent mechanism(64-65). A small percentage (~ 1-5%) of the endocytosed molecules undergoes proteolytic cleavage near residue 110, after which the amino- and carboxyl-terminal cleavage products are C externalized(66). Most of the endocytosed PrP returns to the cell surface, but a small fraction is degraded by lysosome(65). Some of the endocytosed molecules are subject to cleavage within the GPI anchor, and then released into the extracellular medium(64, 66). 16 Plasma membrane (lipid bilayer) (F) (C) (D) (B) Endosome PrPC with cleaved GPI anchor (E) NH2-terminal cleavage product Catabolism Intact PrPC with GPI anchor (A) ER Lumen COOH-terminal cleavage product Figure 1.6. Golgi C Biosynthesis and trafficking of PrP . (A): In the ER. After being synthesized, a nascent PrPC polypeptide is subject to removal of the N-terminal signal peptide. A GPI anchor is added after the C-terminal signal peptide is removed. Glycosylation is initiated. Disulfide bond formation also occurs in the endoplasmic reticulum (ER). (B): In the Golgi apparatus. The N-linked oligosaccharide chains are C modified in the Golgi to yield complex-type glycans. (C): Mature PrP is transported to C the cell surface, where it is attached via its GPI anchor. (D): PrP is internalized through clathrin-mediated endocytosis. Most of molecules remain intact, but a small portion undergoes proteolytic cleavage along the endocytic pathway. (E): A small number of endocytosed PrP C is degraded in lysosome. (F): PrP C is externalized. Most of the molecules remain intact and return to the cell membrane, but some of them are released into the extracellular medium. 17 1.7. C C Proposed functions of PrP . PrP is a copper-binding cell-surface glycoprotein associated with detergent-resistant membranes, also known as lipid rafts. It is most abundant in the CNS, but also exist in other non-neuronal tissues, including spleen, C lymph node, lung, heart, kidney, intestine, and skeletal muscle(67-68). Neither PrP nor its mRNA has been detected in the liver(67). In the brain, PrPC is predominantly localized to the synaptic membranes(69-70). However, later studies challenged this observation and proposed that PrP C distribution on the plasma membrane of neurons has no preference for synapses(71-72). Mice devoid of PrP exhibit normal development and reproduction with a number of subtle abnormalities(73-75). However, given its highly conserved primary sequences, C PrP seems likely to have an important physiological functions. It has been postulated C that the function of PrP is involved only in certain conditions. Therefore, ablation of C C PrP does not affect the unchallenged experimental mice(76). Deletion of PrP may also C initiate a mechanism that compensates for PrP deficiency(76). C Although the exact roles of PrP remain unknown, several potential functions have been C proposed. First, PrP has been suggested to regulate copper metabolism(77-79). It has been proposed that PrPC maintains the level of copper in the pre-synaptic cytosol and C protects synaptic membrane from copper toxicity(70, 80-82). Second, PrP has been suggested to promote neuroprotection through its anti-Bax activity, antioxidant activity, and other signaling pathways(76). These functions require signaling at cell surface and/or 18 C an undefined mechanism associated with octapeptide repeats(76). Third, PrP has been suggested to support the self-renewal of hematopoietic stem cells and to regulate neurogenesis and neural development(83-84). 1.8. Scrapie in sheep. Scrapie, a fatal neurological disorder occurring in sheep and goat, is a member of TSE. Classical scrapie was first reported in Europe in the 1700s, and was the first identified TSE. The neurological signs in classical scrapie include abnormalities in behavior, sensation, and movement. In 1998, an atypical type of scrapie were diagnosed in Norway. Based on PrP Sc conformation and its glycosylation pattern, a new strain (Nor98) is designated for these five cases(85). The major clinical sign in atypical scrapie is ataxia; anxiety and loss of body condition were also observed(85-86). The primary structure affected in classical scrapie, the dorsal motor nucleus of the vagus, has never been involved in atypical scrapie(85-86). These two types of scrapie have different epidemiology. Classical scrapie has been identified as an infectious form of TSE, and can spread through the following routes: First, horizontal spread can occur between two unrelated animals(87). Second, a ewe can transmit prion to her offspring through parturition and fluids(87). Third, parents can spread prion to offspring via germplasm(87). By contrast, although the origin of atypical scrapie remains elusive, two observations suggest that this disease could occur sporadically. First, transmission between sheep in the same flock has not been identified(85-86, 88). There are only two reported cases 19 where more than one affected animals was found in the same flock(89-90). Second, the geographical distribution for atypical scrapie occurrence is more homogeneous than expected for an infectious disease(85-86, 88). In most cases of classical scrapie and many other acquired TSE, prion spreads from peripheral to the central nervous system (CNS) via the lymphoreticular system (LRS). However, PrP Sc is barely found in the peripheral lymphoid tissues in atypical scrapie(91-92). Some authors suggest a possibility that PrP to PrP Sc C conversion occurs sporadically in the brain(85-86, 88), thus LRS is not necessarily needed for prion invasion. Nevertheless, this observation may reflect that atypical scrapie has alternative routes for transporting prion from the peripheral tissues to the CNS. A similar situation has been observed for prion neuroinvasion in different strains of the transmissible mink encephalopathy (TME) agent(85-86, 88, 93). 1.9. Genetic modulation in sheep scrapie. Polymorphisms of ovine PrPs (ovPrPs) have been associated with differing susceptibility to classical and atypical scrapie, particularly at positions 136 (A/V), 141 (L/F), 154 (H/R), and 171 (Q/R). According to surveys conducted in 15 different countries, the average allelic frequencies are 56% for A136L141R154Q171 (ALRQ), 30% for A136L141R154R171 (ALRR), 6% for A136L141H154Q171 (ALHQ), 5% for V136L141R154Q171 (VLRQ), 3% for A136L141R154H171 (ALRH), and 11.6% for A136F141R154Q171 (AFRQ)(88). In classical scrapie, the variations at positions 136 (A/V), 154 (R/H) and 171 (Q/R) are involved in the genetic modulation(88-89, 94-96). The combination of these 20 polymorphisms forms five alleles: ALRQ, VLRQ, ALHQ, ALRR, and ALRH. The VLRQ allele is linked to the highest susceptibility, whereas the ALRR allele confers resistance(88-89, 94-96). The other three variants, ALRQ, ALRH, and ALHQ, are associated with medium to low susceptibilities. However, this scale is not absolute and exhibits prion strain-dependence(88, 97-99). In atypical scrapie, the susceptibility is mainly modulated by the polymorphisms at residues 141 (L/F) and 154 (L/H). The highest susceptibility is associated with AFRQ allele. Other two haplotypes, ALHQ and ALRR, are also linked to atypical scrapie. Surprisingly, VLRQ, which renders the highest risk to classical scrapie, seems to confer resistance to atypical scrapie(85-86, 88-89, 100-101). 1.10. Genetic modulation in human prion diseases. Approximately 10-15% human prion diseases occurs in the familial form(102). Most mutations associated with human familiar prion diseases in Prnp gene involve the following three regions (Figure 1.7): First, human Prnp normally carries five octapeptide repeats between residues 51 to 91. Insertions of extra repeats have been linked to familial prion diseases(103-106). Although deletions of single repeat do not cause prion diseases(2), a case with two-octapeptide repeat deletion has been reported(103). Second, many point mutations have been found in the region between helices 2 and 3(105). Third, premature stop codons caused by point mutations have been reported to induce prion diseases(105, 107). 21 Polymorphisms at residue 129 between Met or Val have been reported to modulate disease susceptibility and pathogenesis in sporadic, acquired, and familial prion diseases(108-112). Homozygosity (129MM or 120VV) at this position appears to predispose individuals to sporadic prion diseases(108-109). In the cases of variant CJD, only patients with 129 MM genotype have been reported(54, 108). The two allelic forms also determine the phenotypes in the familial diseases caused by the pathogenic mutation at residue 178 (D178N)(111-112). Whereas D178N-129MM genotype is linked to FFI, D178N-129MV is associated with familial CJD (fCJD). It has been suggested that this modulation is not absolute. Several cases of fCJD have been reported in patients with D178N-129MM genotype(113). An explanation is that FFI and fCJD represent two extreme phenotypes of a diseases rather than two separate diseases entities(113). 22 Figure 1.7. Mutations Octarepeat causing insertion GSS P102L129M P105L129V A117V129V G131V129M G145*129M H187R- Q217R129V 129M F189S-129V D202N-129V Q212P D178N- E208H M232R 129V V180I E200K T188K V210I Mutations Octarepeat causing insertion gCJD T188R-129V E196KE211Q V203I Mutations causing FFI D178N-129M 23 Figure 1.7. (cont’d) Human PrPC and the mutations linked to familial prion diseases. OR denotes the octapeptide repeats. H1, H2, and H3 denote helices 1, 2, and 3, respectively. Two forks on top of the protein represent potential sites for glycosylation. The sole disulfide bond (S-S) and GPI anchor are also shown in the figure. The associated polymorphisms at residue 129 are indicated. The asterisk indicates a stop codon. Figure was adopted and modified from Aguzzi, A. et al. (2008) Annu. Rev. Neurosci. 31, 439-477. 24 1.11. Prion diseases and other protein misfolding disorders. Several neurodegenerative diseases, including prion diseases, Alzheimer’s disease (AD), Huntington’s disease (HD), Parkinson’s disease (PD), and amyotrophic lateral sclerosis (ALS), involve protein misfolding, and thus are collectively called protein misfolding disorders (PMD). These diseases share some common features in their pathogenesis and usually involve the formation of protein aggregate in the form of fibrillar amyloid(29). It remains to be established whether there is a unifying mechanism to explain neuronal death occurring in these diseases. A ‘seeding nucleation’ model has been proposed to explain amyloid formation(114). In this theory, the rate-determining step is the slow process where nucleation of monomeric misfolded proteins occurs. Once nuclei are formed, further addition of monomers becomes very fast, resulting in a rapid polymerization phase. Although this self-seeding and propagation process seems to be common for PMDs, transmissibility has been observed only in prion diseases. Several studies have shown prion-like transmission in animal models of AD(115-119); nevertheless, many of these results are conflicting(52, 118, 120). It has been argued that the recipient animals used in some of the studies are highly susceptible to amyloidosis, and spontaneous amyloidosis could occur under the experimental conditions(52, 119, 121). Thus, it remains to be established whether inoculation of amyloid in these animals initiates amyloidosis thus reflecting amyloid transmissibility, or simply facilitates an ongoing pathogenesis(122). 25 Another mystery in these diseases is the toxic entity. Earlier studies proposed that protein aggregation account for the neurotoxicity in PMD based on the following observations. First, protein aggregates have been observed to co-localize with brain lesions in many cases(29, 123). Second, genes involved in the familial form of these diseases have been found to encode the aggregated proteins(29). Third, overexpression of the abnormal proteins in animal models induces protein aggregation and causes neurodegeneration(124-125). Fourth, most pathogenic mutations involved in PMD promote in vitro and in vivo protein aggregation(126). However, accumulating evidence suggests the possibility that formation of protein aggregates play a neuroprotective, instead of neurotoxic, role in these diseases. As mentioned in the previous section, neuronal death is not always associated with PrP Sc aggregates in prion diseases. In some cases, diseases develop without detectable PrP Sc aggregates(35-38). A similar situation has been observed in the animal models of polyglutamine diseases(127). Soluble oligomers or diffuse deposits of the misfolded proteins have been shown to correlate with neuronal death in the animal models of AD(128-131). In addition, it has been shown that antibodies against soluble Aβ42 oligomers block the neurotoxicity(132). Taken together, these studies suggest that precursors of amyloid fibers, such as monomeric or soluble oligomeric forms of misfolded proteins, might account for neuronal death. If it is the case, formation of protein aggregate may protect neurons by sequestering the toxic species. 26 Table 1.2. A list of protein misfolding diseases. Table was adopted and modified from Caughey, B. et al. (2003) Annu. Rev. Neurosci. 26, 267-298. Disease Alzheimer's disease Proteinaceous deposit, abnormal protein Extracellular amyloid plaques, fibrillar Aβ Product of abnormal gene Amyloid precursor protein α-synuclein Parkinson's disease Lewy body, fibrillar α-synuclein Amyotrophic lateral sclerosis Superoxide dismutase-1 Transmissible spongiform encephalopathies Fronto-temporal dementia Intraneuronal inclusions, insoluble SOD1 Extracellular plaques, protease-resistant PrP (PrP-res) Neurofibillary tangles, fibrillar tau Diffuse Lewy body disease Cortical Lewy bodies, fibrillar α-synuclein Huntington's disease Intranuclear neuronal inclusions, fibrillar Htn None (sporadic only) Huntingtin Spinocerebellar ataxias Intranuclear neuronal inclusions Prion protein (PrP) Tau Ataxin 27 Phenotype of mutations (including other genes) FAD mutations generally increase production of Aβ42 FPD mutations seem to cause α-synuclein aggregation or Accumulation FALS mutations promote aggregation, but not fibrillization Mutant PrP becomes protease-resistant in vivo Mutant variants of tau have altered affinity for microtubules and a tendency to Aggregate No mutations have been linked Expansion of naturally occurring polyglutamine repeat Expansion of naturally occurring polyglutamine repeat Chapter 2. Introduction to Protein Folding Much of this thesis is focused on kinetic models of protein folding, and their structural interpretation in terms of statistical ensembles of protein conformations (conformational ensembles). This chapter is intended as an introduction to such modeling, and protein folding in general. This chapter will focus on the in vitro folding process, which is less complex than in vivo folding. In vitro folding refers to a process in which protein molecules fold independently of one another and any cellular components. For comparison, in in vivo folding, the protein's conformation is shaped by chaperones and other cellular proteins, not to mention the protein's changing cellular environment as it is synthesized and transported to different compartments. By eliminating the cellular contributions, in vitro folding studies reveal the interplay between intermolecular and intramolecular interactions that are intrinsic to the polypeptide molecule. Thus, the in vitro folding studies pertain to in vivo folding; however, they must be integrated with in vivo folding studies to understand the physiological processes. 2.1 Equilibrium unfolding. Most globular proteins have two conformational states, the unfolded state and the folded state (also known as the native state). The folded state is that revealed by X-ray crystallography, in which most atoms of the protein molecule have a stable position, being held in place by a network of many noncovalent interactions, such as hydrogen bonding, steric packing, van der Waals attractions, and hydrophobic interactions. The 28 relatively stable structure and many interactions give the folded state relatively low entropy and enthalpy. By contrast, the unfolded state describes the high-entropy, highenthalpy state, in which the protein has no stable structure and is rapidly interconverting between conformations, most of them open conformations with few interactions. In naturally occurring proteins under physiological conditions, both the folded and unfolded states are low in free energy, typically differing only by a few kcal/mol. (For comparison, the thermal agitation energy at room temperature is roughly 0.6 kcal/mol.) Relatively small changes in the solution conditions (such as temperature, pH, salt concentrations, pressure, etc.) can alter whether the folded state or the unfolded state is preferred. The very gradual alteration of solution conditions to cause a folded protein to unfold (e.g., by gradually altering the temperature) is known as equilibrium unfolding, because it is carried out under quasi-equilibrium conditions at every step. Such experiments allow us to measure the free-energy difference ∆GNU between the folded and unfolded states. In addition to the folded and unfolded states, other distinct conformational states may be populated significantly under certain solution conditions. For example, these may be partially folded states, or states where the folded conformation is roughly the same but more labile. Generically, these states are known as intermediate states, because they are often populated at intermediate solution conditions between strongly unfolding and strongly folding solution conditions. These three types of states will be discussed in more detail below; for convenience, I denote the folded (native), unfolded and a generic intermediate states by N, U, and I, respectively. 29 2.2. Kinetic modeling of protein folding. The relative concentrations of folded and unfolded conformations under given solution conditions is a function of the relative rates of the underlying basic reactions, folding and unfolding. Since the folded and unfolded states are stable states (free-energy basins), the folding U->N and unfolding reactions N->U are activated processes, meaning that their rate constants vary exponentially with the height of the free-energy barrier, the transition state, between the two states. (This corresponds to Arrhenius' law.) This is true even when there are intermediate states; in general, the rate constants for the transitions among any two conformational states (say, kIN for the transition I->N) is governed by an Arrhenius-like equation. Because protein-folding reactions are unimolecular (e.g., I->U), the ensemble of protein conformations will relax to equilibrium as a sum of exponentials with decay constants λ. If there are p conformational states (N, U and the various intermediates I) then mathematically, there will be p-1 decay constants. These observed decay constants are not the same as the fundamental rate constants, but may approach them very closely. For illustration, in two-state folding N <=> U where p=2, a single exponential phase is observed with the decay constant λ equals kUN + kNU. Under strongly folding conditions, kNU << kUN and hence λ is approximately kUN. In the three-state folding model N <=> I <=> U with p=3, two decay rates will be observed, λfaster and λslower, such that λfaster + λslower equals the sum of the four fundamental rate constants, kNI, kIN, kIU, and kUI. In all cases, given the fundamental rate constants under given solution conditions, it is possible 30 to predict the decay constants λ of protein folding and the corresponding amplitudes of these exponential phases. Conversely, by carefully observing the decay constants λ under various solution conditions, it is possible to determine the number of intermediate states and estimate the fundamental rate constants of the kinetic model, under the assumption that the Arrhenius law holds. That is the procedure followed in this dissertation. The rate constants and their dependence on solution conditions, in turn, provide structural information about the various states, particularly the intermediates, which are difficult to characterize by other means. Here, I have observed a single intermediate in PrP folding kinetics, and from its deduced properties in variants susceptible to scrapie in different degrees, I argue that it is Sc a plausible precursor for the scrapie isoform PrP . This model suggests new scientific hypotheses and potential treatments by targeting this intermediate. In the remainder of this chapter, I summarize the current understanding of conformational states and protein folding, as well as the experimental methods used to characterize its kinetics. 2.3. Conformational states. A simple chemical reaction proceeds from its reactant to its product along a well-defined pathway, i.e. through a sequence of specific structures (Figure 2.1). The point of highest free energy along the reaction coordinate is called the transition state (Figure 2.1). After reaching this point, reactant molecules will always go on to form products. A reaction 31 sometimes involves formation of short-live intermediates on the direct pathway between the reactants and the products (Figure 2.1). Ts1 Intermediate state Reactant H Ea2 Free energy Ea1 Ts2 X+ H Product X H+ X + Reaction coordinate Figure 2.1. An example for a simple chemical reaction involving formation of an intermediate. Ea and Ts are denoted to the activation energy barrier and the transition state, respectively. Protein folding also involves the conversion of the reactant (an unfolded state) to the product (a native folded state). However, since a polypeptide chain can adopt a larger number of possible conformations (i.e., it possesses a much greater conformational freedom) than small molecules in a simple chemical reaction, the unfolded, intermediate, and transition states in protein folding should be viewed as statistical ensembles of 32 conformations rather than any particular conformation. These states occurring in protein folding are also called kinetic species throughout this dissertation. 2.3.1. The unfolded state. The term unfolded state of a polymer has usually been thought to be a random coil with the restriction imposed by steric repulsion between atoms and the excluded volume effect(133-134). However, considerable residual structure in unfolded proteins has been observed(135-137). Such residual structures are only marginally stable; they form rapidly and dissolve rapidly. Nevertheless, this reduces the accessible conformations of a polypeptide chain. The conformation of the unfolded state is determined by the balance between interactions of the protein with itself and those of the protein with the surrounding solvent. When interactions within a protein are more energetically favorable than those with the solvent, a protein tends to be more structured. Conversely, when its interactions with solvent are more favorable than those within itself, a protein shows higher degree of randomness(133-134). Since the chemical properties of amino acids are diverse, intermolecular interactions with the solvent are not homogeneous along a polypeptide chain(133-134). Furthermore, most solvents include more than one component, and a non-uniform distribution of solvent components in a solution has been observed(138-140). Thus, the surrounding solvent for each part of a protein may differ from the mean solvent composition(133-134). Besides the energetic balance between intermolecular and intramolecular interactions, disulfide bonds also play an important role in determining the residual structure of an unfolded protein. 33 2.3.2. The native state. A native folded state of a protein in a conformational ensemble centered on a single well-defined conformation, in which a protein performs its biological function. The three major stabilizing interactions in the native state are hydrogen bonds, van der Waals interactions, and hydrophobic interactions(134). Each individual interaction is weak; however, many of them form a ‘cooperative system’ to stabilize a protein(134). Figure 2.2 represents a simplified cooperative system in protein folding. One interacting pair will bring the other potentially interacting pair into proximity and a more favorable orientation, and decrease the entropic cost for forming the second interacting pair(134). As consequence, the contribution of a cooperative system to protein stabilization is much greater than the sum of individual interaction. The interactions involved in cooperativity are primarily long-range(134). 34 B A B A Native state Unfolded state B A Figure 2.2. Schematic illustration of cooperativity between two interactions in a protein. An oversimplified polypeptide chain with two potential interacting pairs is shown here to explain the formation of a cooperative system. Formation of interacting pair A facilitates and decreases the entropic costs for formation of interacting pair B. 2.3.3. The intermediate states. The driving force for intermediate formation is hydrophobic collapse(141). In contrast to random collapsed conformations predicted for a hetropolymer, hydrophobic collapse in a protein shows structural preference and specific interactions(141). It has been reported that, on average, about 60% of the surface exposed in the unfolded state is buried in the intermediate state(141). In other words, a protein has 35 exhibited high degree of compactness at this stage. Such compact structure is characterized by approximately correct tertiary folds and native-like but fluctuating secondary structure(141). 2.3.4. The transition state. The concept of folding nucleus has been introduced (142-143) to explain the initiation of protein folding. In the nonspecific nucleus model, formation of transition state ensemble (TSE) depends on forming a sufficient number of native interactions without considering which specific contacts are involved(142). In the specific nucleus model(143-147), folding nucleus represents a specific set of interactions which is shared by all the conformations in TSE. Optional interactions also occur in each conformation; however, their number and location vary from one conformation to another(142). Specific and nonspecific nucleus models represent the extreme cases of defining folding nucleus; the real scenario may vary between these two extremes. 2.4. Kinetic models of protein folding. Initiation of protein folding depends on the diffusion-driven(148) contact formations between different regions of an unfolded polypeptide chain. Disulfide bonds impose conformational constraints on unfolded proteins, thus modulating the initiation of folding. By contrast, cis–trans isomerism of proline residues usually accounts for the slowest process in protein conformational folding. 2.4.1. Formation of native disulfide bonds. Oxidative folding is defined as a process consisting of disulfide bond formation and forming the native structure of a protein 36 (conformation folding)(149-150). Several characteristics have been observed for such a coupling process. First, disulfide bonds bring residues distant in the primary structure into proximity, thus affect interactions between residues by altering the entropic cost(134, 149). Second, correctly formed disulfide bonds strengthen native local interactions, which evolve concurrently with the formation of native global folds(149). Third, only the evolution of native tertiary structure has been observed in oxidative folding(149). Fourth, tertiary structures are stable only when native disulfide bonds are formed(149). Fifth, native disulfide bonds limit the conformational freedom of their neighboring residues; therefore, folding nucleus are likely to form in the vicinity of disulfide bonds(149). Such structures have been found in several studies(151-154). In short, disulfide bonds impose bias for conformational folding. 2.4.2. Proline isomerization. Unlike other peptide bonds, which generally favor the trans isomer, those between proline and its preceding amino acid (X-Pro bond) usually exist as an equilibrium mixture in an unfolded ensemble (Figure 2.3)(155). However, when an unfolded protein folds into its native structure, X-Pro exists exclusively in either cis or trans isomer(155). 37 Cα Cα C Cα+1 N C Cα+1 O N O trans cis Figure 2.3. Isomeric states of proline peptide bond and its isomerization reaction. In an unfolded protein ensemble, molecules with native set of X-Pro fold more rapidly than those with at least one non-native X-Pro(155). Nevertheless, non-native X-Pro need not block folding progress, and its re-isomerization may not be required for other folding steps(156). Re-isomerization of non-native X-Pro is a slow process with a time constant of around 10 to 100 sec at 25oC. This reaction can occur either concurrently with, or after, other faster folding steps. This reaction shows a strong dependence on temperature, but not on denaturant concentrations(157-158). 2.4.3. Experimentally observed kinetic models for protein folding. Several small singledomain protein have been found to fold according to a two-state scheme(159-160): N↔U where N and U represent the native and unfolded states, respectively. In this scheme, only the fully unfolded and the fully folded protein are populated; this all-or-none character is due to the cooperative stabilization. The following factors could account for the apparent two-state folding(133): First, folding proceeds without passing through an intermediate state. Second, the population of the intermediate is too small to be detected. 38 Third, the spectroscopic property of the intermediate is indistinguishable from that of the unfolded or native folded protein; therefore, U → I or I → N cannot be detected directly. Fourth, the low steady-state concentration of an intermediate required for the sequential folding step is established too rapid to be detected. Because an unfolded ensemble is composed of a highly heterogeneous population, interconversion between each unfolded conformations has to occur very rapidly relative to the main folding step, so that all the molecules can fold into the native state through the same single step(133). In most cases, proteins exhibit more than one folding phase, suggesting that folding involves the formation of intermediates. Nevertheless, such observations are sometimes difficult to be interpreted due to the heterogeneity in unfolded protein ensemble (Figure 2.4). By contrast, unfolding kinetics usually can be described as a single first-order reaction, since the native protein population is highly homogeneous(133). 39 Scheme 1 (on-pathway) U↔I↔N Scheme 2 (off-pathway) I↔U↔N Scheme 3 (parallel pathway) I ↕ ↕ U↔N Figure 2.4. Three possible folding schemes account for an experimentally observed folding intermediate. In scheme 1, I is on the direct pathway between U and N. In scheme 2, I is a misfolded state that must unfold before folding to N. In scheme 3, conformational interconversion can occur directly among U, N, and I. U, N, and I represent the unfolded, native, and intermediate states, respectively. A long-standing problem in biochemistry and biophysics is the protein-folding problem: How does a protein reach its native folded structure from its unfolded conformational ensemble? This question leads to several sub-questions: Is folding process hierarchical? Does hydrophobic collapse occur concurrently with, or prior to, structure formation? Are there folding nuclei? What is the role of folding intermediates in folding? 2.5. Experimental approaches to study protein folding kinetics. Protein folding can be initiated using rapid mixing techniques, and followed by optical probes, such as ultraviolet-visible (UV) absorption, circular dichroism, X-ray light 40 scattering, and fluorescence. The observed exponential responses represent the barriercrossing events in protein folding; therefore, the number of kinetic species can be deduced from the number of exponential responses. In generally, a kinetic model with n kinetic species should produce a signal of n-1 exponential phases, although not all phases may be detected. A short-lived intermediate state is usually observed only in kinetic studies, although it is sometimes populated to a significant amount that can be detected under certain conditions in equilibrium studies. Some properties of intermediates can be inferred from kinetic data; however, due to the transient nature of intermediates, detailed structural information usually can not be obtained. Detection of the interaction formations along the folding progress can be achieved by applying techniques such as hydrogen/deuterium exchange and mutagenesis combined with kinetic approaches. 2.5.1. Initiation of protein folding. (1) Protein denaturation. One of the most commonly methods to unfold a protein is using chaotropic agents, such as guanidine hydrochloride (GuHCl) and urea. These two denaturants unfold a protein mainly though disrupting hydrophobic interactions; GuHCl also diminishes ionic interactions(161). Although several models suggest that urea and GuHCl increase the solubility of unfolded proteins through hydrogen bonding, a recent report shows that GuHCl does not hydrogen bond to peptide groups(162). Other methods, such as pressure, temperature, and pH denaturations, are also routinely used to unfold proteins. High pressure unfolds a protein mainly through changing its packing and hydration properties(163-164). Cold denaturation causes reduced hydrophobic interactions thus unfolding a protein(165). Heat denaturation disrupts protein structures 41 through breaking its hydrogen bond and other enthalpically favorable interactions(165). The problems associated with this method is that “heat-denatured” proteins are usually partially structured and aggregated, and covalent side reactions sometimes occur as temperature increases. pH denaturation induces protein unfolding by breaking ionic interactions and salt bridges. Similar to heat denaturation, structured proteins are often seen in the denaturing conditions in this method. (2) Changing external conditions to initiate folding reactions. Protein refolding and unfolding can be initiated by changing external conditions; such process has to be carried out rapidly to be studied kinetically. One way to induce protein folding is to mix a solution containing chemical-denatured proteins with a buffer that favors folding using a stopped- or continuous-flow mixing method. Folding kinetics is then followed by an optical probe, such as UV absorption, circular dichroism, X-ray light scattering, and fluorescence. The time resolution of a mixing device is determined by the dead-time of the instrument, which depends on the time required to complete mixing (mixing time), the flow velocity, and the volume between the mixing area and the point of observation(166). Figure 2.5 represents the principles of flow-kinetics. Solutions containing reactants are delivered into the mixer through two syringes; mixing are completed by either turbulent or laminar flow. The main difference between stopped- and continuous-flow methods is as follows: In the stopped-flow method, reactions occurring in an observation cell are monitored after the flow comes to a complete stop(Figure 2.5.A). However, in the 42 continuous-flow method, mixed solution is allowed to flow continuously without being stopped, and signal is recorded along the flow direction from the downstream of the mixing area (Figure 2.5.B). Since the measurements in a stopped-flow method are made after flow stops completely, its dead-time is limited to ~ 2 ms, and artifacts due to vibration and pressure are sometimes induced(166). A much shorter dead-time (10-90 µs) is achieved in a continuous-flow method(166-167). 43 A Syringe 1 Unfolded protein Laser Stop syringe Mixing area Refolding buffer Observation cell Syringe 2 Detector Stop bar Flow direction B Unfolded protein Laser Jet of mixed solution Mixing area Observation channel Refolding buffer Detector Flow direction Figure 2.5. Mixing methods for stopped- (A) and continuous-flow (B) experiments. (A) In stopped-flow kinetics, the flow is stopped after a certain volume of solution has passed through the mixer. The detector then starts to record the reaction in the observation cell. (B) In continuous-flow kinetics, optical signal is recorded along the whole observation channel without stopping the flow(168). 44 2.5.2. Optical probes for folding detection. (1) Protein fluorescence. Absorption of UV radiation by a molecule excites it to an excited singlet state. Fluorescence occurs when this molecule returns to the ground state by emitting a photon. The efficiency of the fluorescence process can be defined as Φ (quantum yield) = (number of photons emitted)/(number of photons absorbed) Three naturally occurring aromatic residues (Phe, Tyr, and Trp) contribute significantly to protein fluorescence. As compared to Phe and Tyr, Trp has much greater quantum yield and absorbance at the wavelength of excitation. Also, fluorescence emission of Phe and Tyr are easily quenched due to the resonance energy transfer(133). Therefore, Trp contributes to most of the observed fluorescence emissions of a protein(133). The three aromatic residues have distinct emission and absorption properties. Although the fluorescence of a protein is a mixture of individual aromatic residues, detection can be made exclusively for Trp by exciting proteins at ~ 295 nm and/or collecting fluorescence above 350 nm(133). Both Trp and Tyr fluorescence are sensitive to their local environment. However, an advantage of Trp over Tyr is that Trp usually occurs rarely in a protein; thus, changes in Trp fluorescence reflects conformational changes in a specific region of a protein. Although the fluorescence intensity can be either stronger or weaker as folding 45 progresses, the emission maximum of Trp shifts to a shorter wavelength (blue-shift) upon forming native structure(133). (2) Fluorescence dye. Interaction with fluorescence dye 8-Anilinonaphthalene-1-sulfonic acid (ANS) and its derivative is a commonly used technique to monitor protein folding, since ANS fluorescence emission greatly increases upon binding to a partially unfolded folding intermediate(169-171). A high binding affinity of ANS to solvent-accessible hydrophobic regions has been proposed to explain this phenomena(170); nevertheless, such binding behavior is not observed in unfolded proteins. Although several alternative theories have been suggested(170-172), the exact binding nature remains elusive. In addition, structural perturbation upon binding of ANS to proteins have been reported(171, 173). Both factors could complicate data interpretations. (3) Circular dichroism. Circular dichroism (CD) refers to the differential absorption of left and right circularly polarized light. In the far-UV region (wavelength 240 nm and below), CD signals of a protein arise mainly from its peptide bonds. The characteristic spectra exhibited by different types of secondary structure have been observed(174) (Figure 2.6), and routinely used to probe secondary-structure composition of a protein. 46 80000 [θ] (deg*cm2*dmol-1) 60000 40000 20000 0 -20000 -40000 160 180 200 220 240 Wavelength (nm) 260 Figure 2.6. Far-UV CD spectra associated with various types of secondary structure. Solid line, α-helix; long dashed line, anti-parallel β-sheet; short dashed line, irregular structure. Figure was adopted with modification from Kelly, S. M. et al. (2005) Biochim. Biophys. Acta. 1751, 119-139. 2.5.3. Probe interactions formation along the folding pathway. Interactions formation in protein folding can be assessed at single residue level by applying techniques such as hydrogen/deuterium exchange and mutagenesis combined with the kinetic approaches mentioned above. 47 (1) Hydrogen/deuterium exchange. Exchange of protons between proteins and surrounding solvent occurs spontaneously. Such reaction barely happens on the hydrogen attached to the backbone carbon atom, and proceeds too fast to be measured for protons located on protein side chains (-OH, -SH, -NH2, and –COOH)(175). Only the exchange reactions occur on amide protons can be measured easily(175). Hydrogen bonding and solvent accessibility determine the exchange reactions. Amide protons involved in hydrogen-bonded structure exhibit slow exchange; nevertheless, those free from hydrogen bonding exchange more rapidly, even when they are not exposed to solvent(134, 176). In other words, hydrogen bonding is more important than solvent accessibility in determining exchange reactions(134). In an unfolded protein or polypeptide chain, hydrogen exchange is a fast process; however, exchange for amide protons in a native form is much slower. Since the exchange rate for an individual amide proton depends on its involvement in hydrogen bonding and its solvent accessibility, isotopic exchange of amide proton is an excellent probe for structure formation. Hydrogen exchange labeling has been coupled with flow-kinetics to obtain detailed structural information for early folding events occurring on millisecond time-scale. As shown in Figure 2.6, unfolded proteins are first equilibrated with D2O in denaturing solution, so that all the liable backbone and side-chain amide protons are exchanged with deuterium (Figure 2.6.A)(134). The well-equilibrated solution (in Syringe 1) is then mixed with refolding solution in D2O (in syringe 2) to initiate refolding reaction (Figure 2.6.B). Proteins are allowed to refold for a chosen time interval (T1-T0) until being mixed 48 with a pulse buffer containing H2O buffer solution at high pH (Syringe 3)(134). Refolding at this time point (T0) is stopped. Amides that already form a stable hydrogen bond at this time point remain deuterated (ND), while those exposed amides become fully protonated (NH) (Figure 2.6.C)(134). After a pulse time (T2-T1), the solution is mixed with H2O at low pH to quench deuterium to hydrogen exchange, and refolding is allowed to complete (Figure 2.6.D). The fully folded protein is then collected and analyzed using NMR. By choosing different time interval (T1-T0) to cover the refolding reaction, the time course of hydrogen bond formation during protein folding can be monitored. 49 Mixer 2 S3 S4 Mixer 3 S2 S1 Mixer 1 T0 T1 T2 (A) D (B) H (C) H D D (D) H O D D H D O Figure 2.7. Schematic illustration for pulsed hydrogen exchange method. Instrumental setting (quenched-flow technique) is on the top. Sequence of reactions (bottom) is illustrated using an oversimplified polypeptide chain. S is denoted to syringe. (2) Transition-state analysis. A comprehensive mapping of protein folding process can be achieved by protein engineering. To assess the contribution of individual amino acid to folding progress, side chains of a protein are modified by site-directed mutagenesis to delete or alter existing weak interactions without perturbing the overall structure(134). A pairwise interaction can be identified by removing the amino acids involving in the 50 interaction separately and together(177). Folding kinetics and equilibrium properties are measured for the wild-type and mutant proteins. The free energies of the native, intermediate, and transition states can be calculated from equilibrium and rate constants. After the energy profile is established, the free energy difference between the wild-type and the mutant protein in each state is compared. ФTs and ФI are introduced to analyze interaction formation at each residue involved in the transition and intermediate states (Figure 2.7): ФTs = ∆∆GU-Ts/∆∆GUN ФI = ∆∆GUI/∆∆GUN where ∆∆GU-Ts, ∆∆GUI, and ∆∆GUN are the apparent energy differences between the wild-type and mutant proteins in the transition, intermediate, and native states relative to the unfolded state, respectively. A Ф value of 1 indicates that the interaction affected by the mutation is formed as completely as in the native protein, whereas a Ф of 0 suggests that the interaction at this site is not formed, as in the unfolded state(134, 177-179). 51 TS’ ∆∆GU-Ts Free energy TS U and U’ I’ ∆∆GUI I N’ ∆∆GUN N Reaction coordinate Figure 2.8. Free energy diagram for Ф-value analysis of transition and intermediate states. N, TS, and I represent the native, transition, and intermediate states, respectively. Prime is denoted to mutant. 2.6. Theoretical aspects of protein folding. 2.6.1. Initiation of protein folding. Contact order of a protein provides a measure of the average sequence separation of the contacting residues in the folded state(180). The first step for protein folding is to form contacts between different regions of an unfolded protein. As in a simple chemical reaction, the free energy landscape of protein folding is determined by the trade-off between the loss of entropy and the gain of favorable native interactions. Entropy cost increases with an increasing distance between the contacting residues in the primary structure, since a larger portion of a polypeptide chain has to be 52 restricted in order to form native contacts. Conceivably, proteins with lower contact order fold faster than those with higher contact order(180). Such simple correlation has been observed for several proteins(181). However, recent evidence indicate that primary structure is also important in determining folding speed by affecting the relative free energies of ordering different part of a polypeptide chain(180-185). When there are several routes along the free energy surface, which involve forming interactions between residues equally distant in the primary structure, folding reactions will favor those with the lowest local free energy of ordering(180-181). 2.6.2. Roles of intermediates in protein folding. The role of intermediates in protein folding is still under debate. Fersht(186) suggested that a stable unfolded state may increase the free energy barrier for conversion to the native folded state under certain conditions, thus is unbeneficial for folding reactions. Consistent with this idea, a gap theory has been proposed. In this model, folding rates show positive correlation with the energy difference between the native state and the state closest to it(142, 187). Statistic evidence has shown that hydrophobic residues in a naturally occurring protein sequence tends to be further apart from each other than expected for a random distribution(188), suggesting that nature selection tends to weaken local contacts in order to destabilize the unfolded state. The same idea has been applied to intermediate formation. Shakhnovich et al. and Fersht et al. proposed that folding is generally faster without passing through an intermediate state(186, 189-190), since the free energy of an intermediate usually lays between the unfolded and the native states. Taken together, these studies suggested that accumulation of an intermediate is a kinetic trap, instead of a facilitator, in protein 53 folding. On the other hand, formation of an intermediates does decrease the entropic costs for structural rearrangements in the sequential folding step, and thus is advantageous for efficient folding(142). Contrary to the studies mentioned above, conditions stabilizing intermediates have been observed to accelerate folding reactions(141, 191-195), suggesting that intermediates play a productive role in protein folding. 2.6.3. Theoretical models for protein folding. Since the 1950s, three classical models have been put forward to visualize protein folding. In the framework model (also called the hierarchical model) (Figure 2.8.A)(134, 196-197), secondary structure forms first, followed by packing these pre-existing structural elements into the native tertiary structure. This model predicts that secondary structure elements themselves are stable in an unfolded protein; the most difficult step in folding is docking these structural elements. However, experiments have shown that the isolated secondary structure is rarely stable in solution(198). Although different amino acids confer different secondary structure propensities to polypeptide chains(199-200), most isolated peptides that form regular secondary structure in a protein are disordered in solution(198). Hydrophobic interactions and long-range contacts within a protein are required to stabilize secondary structure elements(145). In the nucleation-growth model (Figure 2.8.B)(201), small nucleus encompassing localized secondary-structure elements occur randomly all over the unfolded protein(134, 202). These nuclei are not stable, but serve as templates for rapid propagation of tertiary structure. The nuclei involved in the folding process are not specific(134). This model predicts the absence of folding intermediates, thus became 54 unfavorable after intermediate states were observed experimentally(202). In the hydrophobic collapse model(203), hydrophobic interactions collapse a protein first, then folding occurs in a confined volume(134). Conformational search is thus facilitated by decreasing the conformational freedom of a polypeptide chain(134). The problem in this model is that the excess non-specific hydrophobic interactions may interfere subsequent structural rearrangements(204); it also contradicts to experimental evidence where secondary structure is formed concurrently with collapsing(205). In 1995, Fersht and his colleagues proposed the nucleation-condensation model based on their studies on chymotrypsin inhibitor 2 (CI2) protein(145). This model states that tertiary interactions, including hydrophobic and other long-range interactions, form in concert with secondary interactions in the transition state. As the tertiary interactions stabilize otherwise unstable secondary structures, folding becomes more hierarchical and eventually follows the framework model(204, 206). Although this model is in good agreement with several experimental studies(204, 206-209), it remains to be determined whether protein folding follows such a mechanism. 55 Unfolded state Secondary structure formation A Hydrophobic collapse Nucleation B C Growth Diffusion-collision Native state Figure 2.9. Illustration of three classical folding mechanisms. (A) Framework model. (B) Nucleation-growth model. (C) Hydrophobic-collapse model. 2.6.4. Visualization of protein folding. In 1973, Christian Anfinsen proposed the famous thermodynamics hypothesis, stating that the native structure of a protein corresponds to the global minima of free energy. In other words, formation of the native folded structure does not depend on the kinetic routes(210-211). However, Cyrus Levinthal noticed that most folding reactions complete on the millisecond time-scale. By the given large conformational space of a polypeptide chain, random searching of the native state seems implausible for an unfolded protein to find its native structure within such a short time(212). Hence, the search is not fully random, but determined by a trade-off in entropy 56 and enthalpy. This is also evident experimentally, in the small free-energy barriers between the native and unfolded states. To help in visualizing this trade-off, the concept of a free energy landscape for protein folding was introduced(213). Free-energy landscapes have a funnel-like shape(213-214), represents the conformational space as a function of intramolecular-plus-solvation free energy of a given protein (Figure 2.9)(212, 215). The key component in this model is the number of possible conformations at each energy level(212). A protein molecule does not need to follow a specific folding route; instead, a number of paths are available. The conformational space decreases as folding progresses; a series of local optimization occurs during this process. It finally leads to the bottom of the funnel, which corresponds to the global energy minima, i.e. the native structure. 57 (unfolded state) Possible starting configurations (1016) Compact configuration (1010) Free en ergy Transition state (103) Native state Figure 2.10. Schematic representation of the free energy surface calculated for the folding of a 27-mer. This free-energy surface was constructed using Monte Carlo simulation. The possible configuration decreases as the folding progresses down to the bottom of the funnel. Figure was adopted and modified from Dinner, A. R. et al. (2000) Trends. Biochem. Sci. 25, 331-339. Several local energy minima may exist and form shallow valleys on the free energy surface (Figure 2.10.A). Energetic variations within the same energy level are due to fluctuations of local structures, such as rotation of an individual side-chain. The local energy minima correspond to the most stable conformations at each energy level(214). 58 Interconversion between these low-energy species in the local minima usually occurs rapidly(212, 214). Protein folding is determined by the smoothness of its free energy landscape. Levinthal’s argument does not hold because it assumes the same possibility for a protein to travel through each configuration without considering the variations in the free energy of each path(214). It leads to a free energy landscape with a shape of a flat golf course (Figure 2.10.B): There is only one minimum on the free energy surface, which corresponds to the global energy minima. In other words, local energy minima does not exist(214). In the real situation, however, the paths have different likelihood, based on their free energies. The probability for a protein molecule passing through conformations with lower free energies is higher than through those with higher free energies(214). 59 Local energy minima (small valley) B Free energy A Global energy minima (native state in the bottom of funnel) Figure 2.11. Simplified schematic diagram for illustrating different types of free energy surface. Each point on the free energy surface represents each possible conformation. (A) shows a free energy surface with several small valleys, which correspond to local energy minima. (B) shows a free energy surface with equally likely conformations without considering the energy differences in each configuration (golf course-like landscape). In this model, folding is through random searching. Figure was adopted with modification from Dill, K. A. et al. (2008) Annu. Rev. Biophys. 37, 289316. Both thermodynamics and kinetics play important roles in determining folding paths. If the structure with global energy minima is not kinetically accessible for other local energy minima within a reasonable time (i.e., the energy barrier between the small valleys to the bottom of the funnel is too high), proteins will not fold to the structure corresponding to the lowest free energy(214). To satisfy both energetic elements, Gō proposed that kinetic pathway of folding is consistent with, and leads to, the state of 60 minima free energy(216). Such situation can be archived by evolutionarily ordered folding pathway, in which accessible conformational space decreases rapidly to reach the lowest energy state when folding proceeds through defined transition and intermediate states(133). From the view of free energy landscape, the transition state (TS) in protein folding should represent the lowest energy barrier on the way to form the native structure(142). To avoid TS conformational ensemble (TSE) falling back to the unfolded state, it should resemble the native state but be distinct from the unfolded state(142). Since folding reactions descent rapidly to the native state after passing through TS, TSE should contain native interactions(142). In an unfolded polypeptide chain, local interactions are the dominate force, some of which are present in TSE and the native state(142). To be dissimilar to the unfolded state, some native non-local interactions should form in TSE(142). Therefore, formation of TSE is a process where the local interactions fluctuate until proteins form certain native non-local interactions; such conformational ensemble can be viewed as a distorted form of the native structure(142). 61 Chapter 3. Experimental Designs The current development of methods for preventing, detecting, and treating prion diseases is limited by our knowledge of its pathogenesis. Although the role of PrP Sc aggregates in prion diseases is unclear, all the proposed pathogenic models involve a C Sc structural conversion from PrP to PrP (52, 217); for this dissertation, I denote this conversion as ‘misfolding’. To understand the initiation of PrP misfolding, the normal folding pathway has to be characterized. Sheep susceptibility to classical scrapie is governed by the polymorphisms at positions 136, 154, and 171 in ovine PrPs (ovPrPs)(88, 94-96). Since the key event in prion C diseases is the PrP -to-PrP effects on PrP Sc Sc conversion, I hypothesized that polymorphisms exert their precursors, i.e., they affect the disease susceptibility by modulating the structural conversion. I aim to examine whether folding and unfolding of ovPrPs proceed through an intermediate. If such an intermediate exists, I aim to study its role in PrP misfolding by examining whether its population and/or structure correlate with scrapie susceptibilities of ovPrP variants, and to elucidate the role of the sole disulfide bond in its formation. Thus, three specific aims have been proposed: 3.1. Specific aim one: Assay for an ovPrP folding intermediate. 3.1.1. The major gap in knowledge. 62 Previously observed kinetic intermediates in normal PrP folding. An early study suggested that murine PrP (muPrP) folds with a half-life of ~ 170 µs without populating an intermediate(218). However, since these measurements were taken with 2000 µs time resolution, they do not seem reliable(219-220). A later study of human PrP (huPrP) was undertaken with 100 µs time resolution, and observed the accumulation of an early intermediate with a time constant of ~ 50 µs, followed by a rate-limiting folding step with a time constant of ~ 700 µs(219). It is unclear whether such an intermediate exists in other PrP species, and if so, whether the properties of this intermediate are consistent across species. 3.1.2. Hypothesis. In Specific Aim One, I hypothesize that folding and unfolding of ovPrPs proceed through one or more intermediate state(s). This would be consistent with huPrP, which has a very similar primary sequence to ovPrP. 3.1.3. Methods. This project is in collaboration with Dr. Heinrich Roder at Fox Chase Cancer Center in Philadelphia. Folding and unfolding kinetics were followed by tryptophan fluorescence using a continuous-flow capillary mixing method. This method offers two advantages: First, tryptophan fluorescence is an excellent probe of the folding process because of its high sensitivity to environmental changes. In contrast to commonly used fluorescent dyes, such as 8-anilino-1-naphthalenesulfonate (ANS), the intrinsic fluorescence probe Y221W perturbs the protein conformation minimally. Second, the high time resolution (90 µs) of 63 the continuous-flow system(166) allows folding and unfolding events occurring on microsecond time-scale to be detected directly. Continuous-flow systems are at present the most reliable method for recording such events. Such a high time resolution is necessary for following the folding kinetics of PrPs, since this reaction has been reported to occur on the microsecond time-scale(218-221). Third, such high time resolution is required for direct observation of the folding intermediates of many proteins. On the other hand, two disadvantages are associated with this mixing technique. First, this method requires large amounts of protein sample. Second, the maximum observation time of this method is limited to ~ 1000 to 2000 µs. In other words, reactions occurring more slowly than 1000 µs cannot be observed with this technique. 3.2. Specific aim two: Establish the relationship between (un)folding kinetics and susceptibility of ovPrP variants to classical scrapie. 3.2.1. Major gap in knowledge. (1) PrP Sc precursor occurring on the normal folding pathway. An important step in designing therapeutic targets for prion diseases is to identify the PrP precursor of PrP Sc Sc precursor. The occurring under physiological cellular conditions, denoted here as PrP*Cell, has not yet been identified, although a precursor occurring under nonphysiological oligomer-forming and fibrillization conditions (denoted here as PrP*Fib) has been observed(222-227). Presumably, the structure of PrP*Cell should be ready for Sc conversion to PrP*Fib or PrP . In other words, the conformation of PrP*Cell should either Sc be (1) similar to PrP*Fib or PrP , or (2) flexible enough to undergo structural conversion. 64 If the PrP*Cell of both resistant and susceptible variants are poised to form PrP*Fib or PrP Sc (i.e., if the structures of the precursor in these allelic variants do not differ significantly), the genetic modulation should be reflected in the population of the precursor. (2) Previously proposed mechanisms for genetic modulation in classical sheep scrapie. Several mechanisms have been proposed to explain the genetic modulation observed in classical scrapie. Glycosylation patterns have been suggested to differentiate scrapie susceptibilities by modulating the accessibility to residue A/V171(228). Increased susceptibility to classical scrapie displayed by homozygous VLRQ sheep correlates with higher level of plasma PrPC(229). OvPrP variants susceptible to classical scrapie appear to have longer cellular half-lives, when compared to the resistant ones(230). The propensities of ovPrPs for structural conversion, both in vitro and in vivo, correlate with their susceptibilities to classical scrapie(231-235). In Kelly’s model, any mutation destabilizing the native state is predicted to enhance amyloid formation(236). However, this model is in apparent contradiction with previous observations. In classical scrapie, susceptible variants are more stable than the resistant ones, as measured by the free energy difference between the native and unfolded states, ∆GNU(233, 237-238). Different in vivo clearance has been proposed to explain this paradox(230, 233, 237-238); nevertheless, it cannot account for the increased in vitro propensities to misfolding observed in susceptible variants. 65 Native PrPC has been suggested to be PrP Sc precursor. Zhang et al. found that Syrian hamster PrPC possesses a relatively open conformation based on circular dichroism and NMR measurements, and proposed that the flexible region of native PrPC is susceptible to structural conversion(46). Welker et al. suggested a domain swapping model, where oligomerization proceeds through a thiol/disulfide exchange reaction between two native PrPs(239) (Figure 3.1). Such dimeric structure has been observed by Knaus and his colleagues(43). Nevertheless, a later report on in vitro conversion study seems to contradict to this model(240). S SH S S S PrPSc PrPC PrPSc PrPC S S SH Sc Figure 3.1. Conversion from PrPC to PrP PrPSc PrPSc through disulfide polymerization in the domain swapping model. Attack of the terminal thiolate of PrP of PrP C C initiates the PrP -to-PrP Sc S S SH S Sc on the disulfide bond conversion. Association with PrP Sc weakens the C tertiary structure protection of the disulfide bond in PrP , thus facilitating polymerization reactions(239). Several studies have been carried out to assess the relationship between the native state and the susceptibility to prion diseases. Many of the mutations corresponding to human inherited prion diseases do not alter the thermodynamic stability in the native state 66 relative to the wild-type huPrP(241-242). In the case of classical scrapie, susceptible alleles confer even higher structure stability in their native forms, as compared to the resistant ones(233, 237-238). Moreover, a recent study shows that muPrPs carrying pathogenic or protective mutations in human or sheep prion diseases exhibit structures and backbone dynamics similar to their wild-type counterpart in the native state(243). Taken together, these studies suggest that genetic modulations associated with prion diseases do not exert their effects on the native form. Therefore, if they are associated with folding whatsoever, they must exert their effects on a folding intermediate. If so, a Sc folding intermediate seems to be a potential PrP precursor. Kuwata et al. have identified the intermediate states of human and hamster PrPs by high pressure NMR(244-246), but their correlation with the disease occurrence has not been established. As mentioned earlier, Apetri et al. observed a kinetic intermediate in huPrP refolding(219-221). The mutations linked to familial prion diseases result in a pronounced increase in the population and the hydrophobicity of this intermediate(219, 221). However, such an intermediate was not observed for murine PrP(218). It remains to be established whether genetic variations in different mammalian species modulate disease susceptibility in the same way as observed in human inherited prion disease. 3.2.2. Hypothesis. To explain the paradox mentioned above, I hypothesize that the folding intermediate observed in Specific Aim One is the PrP Sc precursor, and genetic modulation in classical scrapie exerts its effects on this kinetic species, and not on the native form. In other 67 words, I hypothesize that the structure and population of this intermediate are critical in determining the susceptibility of ovPrP allelic variants. 3.2.3. Methods. This hypothesis will be tested by comparing the folding and unfolding kinetics of four ovPrP variants using the method described in Specific Aim One. The relationship between the susceptibilities and the intermediates associated with four allelic variants will be established, then the plausible role of the intermediate in classical scrapie will be inferred. 3.3.Role of the disulfide bond in stabilizing the folding intermediate. 3.3.1. Major gap in knowledge. The role of the sole disulfide bond (Figure 3.2) of PrPs in prion diseases remains controversial. Some studies suggest that an intact native disulfide bond is required for the C PrP -to-PrP Sc conversion (247-249); however, others indicate that this disulfide bond has to be reduced to initiate misfolding(239, 250-253). Welker et al. suggest that an Sc intermolecular disulfide bond does not bridge monomeric PrP , and, in contradiction to the previous studies, temporary reduction of the intramolecular disulfide bond is not Sc necessary for converting to PrP (240). A very recent report has shown that the disulfide bond restricts the motion of the putative transmembrane domain (residues 111-135) of Syrian hamster PrP, thus affecting the initiation of misfolding by regulating membrane binding(254). 3.3.2. Hypothesis. 68 I hypothesize that the disulfide bond in PrP suffices to stabilize the formation of the previously observed folding intermediate (see Chapter Four). Although the role of the disulfide bond in prion diseases has not been well established, the importance of disulfide bonds in protein conformational folding(149) suggests that the disulfide bond may be involved in the PrP misfolding. A native disulfide bond imposes conformational constraints on its neighboring residues, stabilizing its native local structure; this property permits the formation of folding intermediates in some proteins(149, 151-154). Therefore, the disulfide bond in PrPs may exert its effects by stabilizing a folding nucleus for the intermediate. (Folding nucleus is defined here as a specific subset of native interactions required for stabilizing an intermediate state.) If so, and if PrP misfolding proceeds through a folding intermediate, the local structure around the disulfide bond may serve as C Sc a folding nucleus for directing structural conversion to both PrP and PrP . 69 Figure 3.2. Positions of the sole disulfide bond and model peptides in ovPrP. (A) Ribbon representation of the structure of ovPrP (PDB entry: 1UW3). The disulfide bond is shown in gold. P1 and P2 are shown in blue and red, respectively. (B) Sequence alignment of homologous PrP. α and β are denoted to α-helix and β-sheet in PrPs. (A) Helix 1 Helix 2 P1 P2 Disulfide bond Helix 3 70 Figure 3.2. (cont’d) (B) βA Sheep Human Cat Mink Bovine Goat Deer Elk YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDI K I MERVVEQ YSNQNNFVHDCVNITI KQHTVTTTTKGENFTETDVKMMERVVEQ YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDMK IMERVVEQ YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDMKI MERVVEQ YSNQNNFVHDCVNITVKEHTVTTTTKGENFTETDI KMMERVVEQ YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDI K I MERVVEQ YNNQNTFVHDCVNITVKQHTVTTTTKGENFTETDI KMMERVVEQ YNNQNTFVHDCVNITVKQHTVTTTTKGENFTETDI KMMERVVEQ α1 βB ********************* Sheep MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDR Human MLGSAMSRPI IHFGSDYEDRYYRENMHRYPNQVYYRPMDE Cat MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ Mink MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYKPVDQ Bovine MLGSAMSRPLIHFGSDYEDRYYRENMHRYPNQVYYRPVDQ Goat MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ Deer MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ Elk MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ α3 α2 ********************************************** ************************* P2 P1 α3 ************************ Sheep Human Cat Mink Bovine Goat Deer Elk MCITQYQRESQAYYQRGASVI LFSSPPVILLISFLIFLIVG MCITQYERESQAYYQRGSSMVLFSSPPVILLISFLIFLIVG MCVTQYQKESEAYYQRRASAI LFSSPPVILLISFLIFLIVG MCVTQYQRESEAYYQRGASAI LFSPPPVILLISLLILLIVG MCITQYQRESQAYYQRGASVI LFSSPPVILLISFLIFLIVG MCITQYQRESQAYYQRGASVI LFSPPPVILLISFLIFLIVG MCITQYQRESEAYYQRGASVI LFSSPPVILLISFLIFLIVG MCITQYQRESEAYYQRGASVI LFSSPPVILLISFLIFLIVG P2 3.3.3. Methods. To examine this hypothesis, a peptide model flanking the sole disulfide bond will be used to measure the local folding of PrP through circular dichroism (CD) and hydrogen/deuterium exchange (HDX). 71 (1) Selection of peptide models. I chose to use a peptide model, instead of the full-length protein, because it allows me to sort out the contributions of local and global interactions in forming the folding intermediate. The peptides are selected based on the conservation of the residues around the disulfide bond without considering their involvement in the secondary-structure elements. This allows me to assess the role of the disulfide bond, instead of the intrinsic secondary-structure propensity of the peptides, in the local structure formation. Two peptides are selected: (P1)YSNQNNFVHDCVNITVKQH (residues 169-187), and (P2) MERVVEQMCITQ YQRE (residues 206-221). P1 involves the N-terminus of helix 2 (residues 175-197), whereas P2 involves the central region of helix 3 (residues 202-230). Both peptides do not cover the whole of their helices (Figure 3.2). (2) Hydrogen/deuterium exchange and circular dichroism measurements. The hydrogen/deuterium exchange (HDX) method allows me to probe the hydrogen bonding and solvent accessibility of individual residues in the peptides. If interactions around the intramolecular disulfide bond suffice to form the PrP folding intermediate, the P1P2 model peptide should adopt its native structure (α-helix) in solution. Therefore, its HDX rate will be significantly decreased compared to its monomeric components (P1 and P2). HDX is able to probe individual amides that are involved in stable secondary structure and stable hydrogen bonding, and also gives insight into the formation of tertiary structure. However, marginally stable secondary structure and other interactions that are not involved in stable hydrogen bonding are not detectable by this method(134). This limitation can be overcome by monitoring the folding reaction with far-UV circular 72 dichroism (CD). This technique measures the average secondary-structure content, including the structure elements that are only marginally stable(134). Mass spectrometry, instead of NMR, is chosen to monitor HDX reactions in this study because of its low sample requirements. 73 Chapter 4. Effects of Polymorphisms Associated with Scrapie Susceptibility on the Folding Kinetics of Ovine Prion Proteins 4.1. Introduction. Prion diseases are a class of neurodegenerative diseases that includes scrapie in sheep and goats, bovine spongiform encephalopathy in cattle (mad cow disease), as well as fatal familial insomnia, Gerstmann-Straussler-Scheinker syndrome, and Creutzfeldt-Jacob disease in humans. According to the prevailing hypothesis(255), prion diseases result from the misfolding of a naturally occurring prion protein (PrP) from its normal cellular C Sc form (PrP ) to a virulent scrapie form (PrP ). This abnormal isoform PrP Sc is able to catalyze the conversion of PrPC to itself(255), and its aggregation has been proposed to explain the neuronal death in prion diseases(30-31, 255). Although some studies suggest that PrP Sc aggregates are not neurotoxic(20, 34, 256), all models of pathogenesis involve the structural conversion of PrP(52, 217). PrP C is a copper-binding cell-surface glycoprotein composed of an unstructured N-terminal domain and a globular C-terminal domain (residues ~120-233) of known structure, comprising three α-helices, a short twostranded antiparallel β-sheet, and a disulfide bond (Cys182-Cys217) linking helices 2 and Sc 3 (Figure 4.1)(42-45, 257-258). Relatively little is known about the structure of PrP , 74 except that it appears to have significantly more β-sheet and slightly less α-helix (17-30% α-helix; 43-54% β-sheet) compared to PrPC (47% α-helix; 3% β-sheet)(13-15). 171R/Q N’ reporter group (Y221W) C’ 136A/V 154R/H 141L/F Figure 4.1. Ribbon representation of the structure of ovPrP (residue 126-233). Helix 1 is shown in green; helix 2 is shown in mauve; helix 3 is shown in blue. The antiparallel β-sheet is shown in yellow; the disulfide bridge is shown in gold. The residues involved in genetic modulation in sheep scrapie are shown in red; the introduced fluorescence reporter is shown in green. Figure was drawn with MOLMOL(47). Polymorphisms of ovine PrPs (ovPrPs) have been associated with differing susceptibility to classical and atypical scrapie, particularly at positions 136 (A/V), 141 (L/F), 154 (H/R), and 171 (Q/R) (Figure 4.1)(88). In classical scrapie, the A136L141R154R171 (ALRR) and A136L141H154Q171 (ALHQ) alleles confer resistance, whereas the V136L141R154Q171 (VLRQ) and A136L141R154Q171 (ALRQ) genotypes are correlated with susceptibility(88, 75 94-96). Several factors, alone or in combination, might account for this genetic modulation(228-229, 231, 233-235, 237-238, 259). Variations in glycosylation patterns between allelic variants have been suggested to modulate scrapie susceptibilities by altering solvent accessibility(228, 259). PrPC variants susceptible to classical scrapie appear to have longer cellular half-lives and higher plasma concentrations, when compared to resistant variants, giving them more opportunities to convert and Sc oligomerize(49, 229, 233, 238, 260). Finally, the propensity for PrPC-to-PrP conversion, both in vitro and in vivo, also correlates with susceptibility to classical scrapie(231, 234-235, 237). Since this structural conversion requires at least partial unfolding, it is plausible to hypothesize that more susceptible variants are less stable structurally(236). However, exactly the opposite is observed for classical scrapie: variants with lower structural stability (as measured by the free energy difference ∆GNU between the folded and unfolded states) are less prone to the disease than those with higher structural stability(233, 237-238). Different in vivo clearance has been proposed to explain this inverse correlation; nevertheless, it cannot explain the higher in vitro conversion propensities displayed by susceptible variants(231, 234). To account for this apparent paradox, I made the working assumption that polymorphisms affect the susceptibility by modulating the population and/or structure of the PrP Sc precursor, which I took to be a folding intermediate. This hypothesis led me to examine two questions: First, do the folding and unfolding of ovPrPs proceed through an intermediate? Second, if so, is this folding intermediate the PrP Sc precursor? 76 C To study these questions and the folding of PrP in general, I measured the microsecond folding and unfolding kinetics of these four allelic variants under various solution conditions, including two temperatures (5 and 15 °C), two pH values (5 and 7), and two denaturants [guanidine hydrochloride (GuHCl) and urea]. The unfolding kinetics of ALRQ, ALRR, and VLRQ variants exhibited two exponentials at pH 7: an initial lag phase followed by a slower exponential decay. The double exponential indicates the presence of at least one intermediate species besides the native (N) and unfolded (U) species. Biophysically, the most plausible kinetic model is N→Iα→U, in which the native species N unfolds quickly to a native-like intermediate Iα before unfolding completely to U. This intermediate is denoted as Iα because, being similar to the native species N, its secondary structure seems likely to be largely α-helical. Such α-helix-rich species have been observed on oligomerization pathways(222-225, 261). Adopting this kinetic model, I find that the population and structural stability of Iα in the variants correlate with their differing propensities towards classical scrapie. Variants with higher classical scrapie propensity are characterized by higher populations and more stable tertiary structure in Iα. Hence, I conjecture that the formation of this partially unfolded, native-like intermediate Sc Iα is an early step in the formation of PrP . 4.2. Experimental procedure. 4.2.1. Protein preparation and purification. The plasmid encoding ovPrP94-233, ALRQ allele with an N-terminal (His)6 tag was a generous gift from Dr. P. M. Bayley. Based on 77 this parental plasmid, ALRR, ALHQ, VLRQ, and AFRQ plasmids were constructed by site-directed mutagenesis using QuikChange kit (Stratagene) and the following primers: For A134 → V134 mutation: 5’-GGC TAC ATG CTG GGA AGT GTG ATG AGC AGG CCT CTT ATA C-3’ and 5’-GTA TAA GAG GCC TGC TCA TCA CAC TTC CCA GCA TGT AGC C-3’ For L141 → F141 mutation: 5’-GGA AGT GCC ATG AGC AGG CCT TTT ATA CAT TTT GGC AAT GAC-3’ and 5’-GTC ATT GCC AAA ATG TAT AAA AGG CCT GCT CAT GGC ACT TCC-3’ For R154 → H154 mutation: 5’-GAG GAC CGT TAC TAT CAT GAA AAC ATG TAC CGT TAC CCC-3’ and 5’-GGG GTA ACG GTA CAT GTT TTC ATG ATA GTA ACG GTC CTC-3’ For Q171 → R171 mutation: 5’-CTA CAG ACC AGT GGA TCG CTA TAG TAA CCA GAA C-3’ and 5’-TTC TGG TTA CTA TAG CGA TCC ACT GGT CTG TAG-3’ All sequences were confirmed by DNA sequencing. These plasmids were transformed into competent E. coli BL21 (DE3) cells. To express recombinant proteins, E. coli cells containing the desired plasmid were grown in LB medium containing 100 µg/ml ampicillin with shaking at 250 rpm at 37 oC. Based on 78 induction studies, once the optical density reaches 0.6 - 1.0, expression is induced by adding isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM. The culture was then incubated overnight. After the cells were harvested, sonication was used to disrupt the cells. Soluble proteins and inclusion bodies in the cell lysate were separated by centrifugation at 12,000 rpm. Since the expressed ovPrPs accumulate in inclusion bodies (Figure 4.2), inclusion bodies were collected and incubated in 8 M urea with gentle stirring at 4oC for a few hours to solubilize the proteins. Purification was carried out by metal-affinity chromatography using nickel-nitrilotriacetic acid (Ni-NTA) resin (Qiagen). OvPrPs were eluted in a elution buffer containing 50 mM NaH2PO4, 300 mM NaCl, 0.5 M imidazole, and 5 mM β-mercaptoethanol, pH 8.0. The purified proteins were then immediately dialyzed into a buffer containing either 50 mM sodium acetate (pH 5) or 50 mM imidazole (pH 7) using Millipore protein concentrator. This step has to be completed in half an hour; otherwise, extensive aggregation occurs. The yield of pure protein is about 1 mg/L of culture. Protein concentration was determined by the optical density at 280 nm using an extinction coefficient of 22015 M-1cm-1 predicted from the amino-acid composition. The identity of the purified protein was confirmed by SDS-PAGE, Western blotting (Figure 4.3), and mass spectrometry (Figure 4.4). Sedimentation equilibrium experiments were carried out on a Beckman XLI analytical ultracentrifuge at 15000 rpm, 25 oC. After 6 days, the traces converged to a classic 79 Boltzmann distribution with an estimated molecular weight of ~17 kDa, in close agreement with molecular weight of ovPrP94-233 (16.9 kDa), suggesting that the purified ovPrP is monomeric (Figure 4.5). Figure 4.2. SDS-PAGE and Western blotting show the expression of ovPrP in E. coli. Left panel, SDS-PAGE (12%) analysis. Right panel, Western blotting analysis using antipoly(histidine) as the primary antibody. Lanes 1 & 3: Cytosolic fraction of cell lysate corresponding to 25 ◦C and 30 ◦C growth temperatures, respectively. Lanes 2 & 4: Inclusion body of cell lysate corresponding to 25 ◦C and 30 ◦C growth temperatures, respectively. M: Invitrogen protein ladder. M P P Monomeric PrP Figure 4.3. Monomeric ovPrP(ALRQ) eluted from Ni-NTA column. Left panel, SDSPAGE (12%) analysis. Right panel, Western blotting analysis using anti-poly(histidine) as the primary antibody. P: ovPrP. M: Invitrogen protein ladder. 80 16821.76 Relative intensity 100 80 60 40 20 0 15199.4 16399.8 17600.2 m/z Figure 4.4. MALDI mass spectrum of the ALRQ variant. 81 18800.5 20001.0 Optical Signal 1 1 M GuHCl 1 0.5 0.5 0 0 6.2 Optical Signal 2 1.5 0 M GuHCl 6.4 6.6 6.8 6.2 7 1.5 2 0.5 M GuHCl 1 6.4 6.6 6.8 7 5.5 M GuHCl 1.5 1 0.5 0.5 0 0 6.2 6.4 6.6 6.8 7 6.2 6.4 6.6 6.8 7 Radius, cm Radius, cm Figure 4.5. Concentration profiles of sedimentation equilibrium. As shown in the figure, the concentration profiles assume a Boltzmann distribution, indicating that ovPrP was monomeric under these four different solution conditions. Measurements were carried out in 50 mM sodium acetate with various concentrations of GuHCl, pH 5. 4.2.2. Tryptophan fluorescence reporter. The local environment of a tryptophan changes as a protein undergoes conformational changes. Since tryptophan fluorescence is sensitive to environmental changes, it was used to probe folding process in this study. The only tryptophan (Trp102) in wild-type ovPrP94-233 is located in the unstructured Nterminal segment and thus is a poor probe of conformational folding (Figure 4.6). The mutation W102F was made to eliminate the signal from the native tryptophan, and the 82 mutation Y221W was made to introduce a fluorescent reporter (Figure 4.1). A corresponding mutation was made in an earlier kinetic study of human PrP(219-221). Site-directed mutagenesis was carried out for all the variants as described above using the following primers: For W102 → F102 mutation: 5’-GGT AGC CAC AGT CAG TTC AAC AAG CCC AGT AAG CC-3’ and 5’-GGC TTA CTG GGC TTG TTG AAC TGA CTG TGG CTA CC-3’ For Y221 → W221 mutation: 5’-GCA AAT GTG CAT CAC CCA GTG GCA GAG AGA ATC CCA GGC-3’ and 5’-GCC TGG GAT TCT CTC TGC CAC ATT TGC-3’ The secondary structure of native ovPrP and its double mutant W102F/Y221W was assessed by circular dichroism (CD); both spectra exhibit a double minimum at around 208 and 222 nm, which is typical for an α-helical protein (Figure 4.7). Equilibrium unfolding experiments monitored by far-UV CD and fluorescence indicate that the double mutant has native-like stability (Figure 4.8), and that its fluorescence shows a significant increase in the quantum yield (approximately 4- to 6-fold) upon unfolding (Figure 4.9). 83 Native Trp ¤ ¤¤ 182 ¤ ¤ ¤ & ¤ 205 & & 165 & ¤ 125 102 94 (His)66tag (His) tag ¤ & 233 221 217 Cysteines involved in the disulfide bond ¤¤ Fluorescence reporter was introduced here Figure 4.6. Primary structure of the wild-type ALRQ variant. Positions of the native tryptophan, fluorescence reporter, and cysteines involved in the sole disulfide bond are indicated. The native prolines and tyrosines are labeled with the symbols & and ¤, respectively. 84 [ ]MRW mdeg dmol-1cm2 θ 0 -5 -4 -10 -8 A -12 200 220 240 B -15 260 wavelength (nm) 200 220 240 260 wavelength (nm) Figure 4.7. Far-UV circular dichroism spectra of the wild-type ovPrP and the ovPrP(W102F/Y221W). The spectrum of ovPrP(W102F/Y221W) (A) exhibits similar secondary structure as observed in wild-type ovPrP (B). Experiments were carried out in 50 mM sodium acetate, pH 5. The unit of Y-axis is in mean residue molar ellipticity (x 103). 85 Unfolded zone Transition zone Native zone Relative Fluorescence Emission [GuHCl] M Figure 4.8. Representative equilibrium transition curve of ovPrP(W102F/Y221W) monitored by tryptophan fluorescence. Excitation wavelength was 290 nm; fluorescence emission between 300 to 400 nm was collected. All the signal was normalized by the signal of unfolded protein (taken at 6.2 M GuHCl). Only the transition curve monitored at 360 nm is shown here. Raw Fluorescence Emission unfolded PrP native PrP Wavelength (nm) Figure 4.9. Fluorescence emission of native and unfolded ovPrP(W102F/Y221W). 86 4.2.3. Continuous-flow measurements. (1) Instrumentation. Figure 4.10 illustrates the continuous-flow capillary mixing apparatus. Reactants are delivered into the mixing region by the driven syringes through the two coaxial capillaries. Both solutions were forced through the narrow gaps between the platinum sphere and the outer capillary, where mixing occurs rapidly under extremely turbulent flow. The completely mixed solutions then enter the observation channel, still under highly turbulent flow. Reactions are followed by recording the fluorescence emission along the observation channel; distance along the channel is converted into time axis according to the flow rate in use. (2) Experimental procedure. The folding and unfolding kinetics of ovPrPs were followed by tryptophan fluorescence of Trp221 using a continuous-flow capillary mixing method described previously(166). The excitation wavelength was 296.7 nm. Only fluorescence above 324 nm was collected to minimize the signals from the tyrosines in the protein. For most experiments, mixing was carried out at 15 oC at a flow rate of 1.8 ml/s, with a final protein concentration of ~11 µM. The refolding (resp. unfolding) reactions were initiated by diluting the unfolded (resp. native) protein at 1:5 ratio into the same buffer containing various denaturant concentrations. The buffer consisted of either GuHCl or urea in either 50 mM sodium acetate, pH 5 or 50 mM imidazole, pH 7. The concentration of denaturants was determined from their refractive index. 87 Figure 4.10. Sample delivery and detection in the continuous-flow method. (A) Schematic diagram of the capillary mixing apparatus. Figure was adopted and modified from Roder, H. et al. (2006) Chem. Rev. 106, 1836-1861. (B) Illustration of mixing reaction. (C) Folding events occurring in the mixing apparatus. (A) (2) (3) (4) (3) (6) (1) (4) (7) Mixing area (5) (8) (5) (12) (10)(11) (9) Light source for exciting proteins (1): Syringe drive. (2): Syringes. (3): Inner capillary. (4): Outer capillary. (5): Mixer. (6): Pt sphere (200 µm). (7): Arc lamp. (8): Monochromator. (9): Observation channel. (10): Filter (250 µm)2. (11): Lens. (12): CCD detector for recording fluorescence emission. 88 Figure 4.10 (cont’d) (B) Syringes (1) and (2) are filled with refolding buffer and unfolded PrP, respectively. ↓ Refolding buffer and unfolded PrP are delivered to the mixing area through inner and outer capillaries, respectively. ↓ The completely mixed solution enters the observation channel. ↓ Protein fluorophore is excited by the light source set at 296.7 nm. ↓ Protein fluorescence passes through a 324 nm high-pass filter (i.e. only fluorescence above 324 nm can pass through the filter and be collected), and is recorded using the CCD camera. 89 Figure 4.10 (cont’d) (C) Unfolded protein (T = -100 µs) Turbulent flow ‘block’ Refolding buffer T = 0 µs T = 100 µs T = 600 µs Folding progresses Unfolded protein ensemble Native folded protein T = -100 µs: Refolding buffer and unfolded protein are in the inner and outer capillaries, respectively. T = 0 µs: The two solutions are mixed by extremely turbulent flow to establish the final solution conditions and thus initiate the folding reaction. T = 100 µs: The protein is on the way to forming its native state. T = 600 µs: The folding protein reaches its native state. 90 (3) Control experiments. To determine the mixing efficiency and assess possible artifacts, a series of control experiments was carried out as follows: Dead-time calibration. Dead-time of the continuous-flow instrument was determined as described previously(262). Briefly, quenching reactions between N-acetyl tryptophan amide (NATA) and N-bromo-succinamide (NBS) were carried out at several quencher concentrations. The kinetic traces are extrapolated to the time before the first observable point and intersect at one point, which we define as the absolute time zero. The fluorescence signal at this point is equal to one sixth of the signal produced by NATA before it passes through the mixing channel. The time interval between the intersection and the first observable point is the instrument dead-time (Figure 4.11). The dead-time of the capillary mixer was determined to be ~90 µs at 1.8 ml/s. Viscosity was taken into account by including appropriate amount of glycerol to match the viscosity caused by denaturants in the mixing experiments. The formulae for calculating B-viscosity coefficients are as follows(263): For urea: y = 0.0633x + 0.9679 (4.1) For GuHCl: y = 0.0643x + 0.9756 (4.2) For glycerol: y = 0.3047x + 0.9479 (4.3) 91 where x and y represent the concentration of cosolutes and the relative viscosity, respectively. T0 Relative fluorescence Tobs a b c d Time, µs Figure 4.11. Time course of the quenching reactions between NATA and NBS of various concentrations. All the traces were normalized by the signal of NATA in the absence of NBS. The concentrations of NATA was kept constant at 37.5 µM (after mixing). The solid line represents the best fit of the kinetic traces to one exponential. T0 and Tobs represent the absolute time zero (the intersection of traces) and the first observable time point. The dead-time of the instrument is defined as (Tobs - T0). Mixing artifacts. (i) Control experiments taken under the same initial and final conditions. These two experiments were designed to probe mixing artifacts that occur even when the solution 92 conditions are not changed upon mixing. In the unfolded control (resp. native control) experiments, proteins in either 5 M (resp. 1.8 M) GuHCl or 8 M (resp. 3 M) urea were diluted at 1:5 ratio into the same buffer with the same concentration of denaturant. Since the solution conditions remain constant, the protein conformational ensemble should not change; in particular, the protein fluorescence should remain constant over the whole experiment. NATA spectra were also taken under the same unfolded and native conditions. Since the fluorophore of NATA is fully exposed under every solution condition, the observed NATA fluorescence should not exhibit time-dependent changes. If changes in NATA fluorescence are observed, these changes should be attributed to artifacts resulting from the instrument and/or solution conditions, and are expected to occur in PrP control experiments as well. Another potential artifact is that proteins might undergo mechanical stretch (shear stress) when passing through the extremely turbulent region of the mixer. If such an artifact exists, protein conformational ensembles might be pulled out of equilibrium and may relax in the observation channel. Since NATA is a small molecule, it is immune to shear stress. Therefore, by comparing PrP and NATA spectra, one can assess the contribution of such mechanical artifacts for proteins. (ii) Control experiments taken under refolding and unfolding conditions. These experiments were designed to probe mixing artifacts that result from changing solution conditions. Except that NATA was used instead of PrPs, these conditions were the same as those used for our main PrP kinetic studies. In the refolding control (resp. unfolding control) experiments, NATA in either 5 M (resp. 1.8 M) GuHCl or 8 M (resp. 3 M) urea were diluted at 1:5 ratio into the same buffer with differing concentrations of denaturant. 93 Again, since the fluorophore of NATA is fully exposed under every solution condition, NATA fluorescence should remain constant over the whole experiment. By comparing spectra of NATA taken in such control experiments with those taken in the first pair of control experiments (i), one can probe artifacts resulting from mixing two solutions that contain different concentrations of denaturant. Estimate the effects from protein association. To determine whether the observed ovPrP (un)folding kinetics was affected by protein association, refolding measurements for the ALRR variant were carried out at six different concentrations (ranging from 3.2 µM to 13.5 µM for the final GuHCl concentration), pH 7. A systematical concentration dependence within this range was not found, suggesting that the observations mainly reflect the folding of monomeric ovPrP. Nevertheless, it cannot exclude the possibility that the concentration required for multimeric ovPrP dissociation is lower than 3.2 µM. Baseline for kinetic traces. Most experiments were taken at a flow rate of 1.8 ml/s, corresponding to a time window of ~0.6 ms. To get a better baseline for kinetic traces, the fluorescence emission of the flow-through from each refolding measurement was recorded. However, extensive protein aggregation in the flow-through was observed within half an hour; thus, the signal from the flow-through was not able to correct the baseline of kinetic traces. As an alternative method, traces were taken at a flow rate of 0.9 ml/s, which doubles the coverage of the time frame, to estimate the baseline correction. The traces taken at both 94 flow rates were combined; the combined traces exhibit exponential phases similar to those observed in the measurements taken at 1.8 ml/s flow rate (Figure 4.12), suggesting that the baseline in the former measurements is well-defined. Relative fluorescence 0.90 0.85 0.80 from trace taken at 1.8 ml/s 0.75 0.70 from trace taken at 0.9 ml/s 0.65 0.60 0.0 0.2 0.4 0.6 0.8 1.0 -3 Time, µs Figure 4.12. Combination of ALRQ refolding kinetic traces taken at flow rates of 0.9 and 1.8 ml/s. Traces were taken in 2 M GuHCl at pH 5 and 15 oC. The solid line represents the best fit to the combined trace. 4.2.4. Exponential fitting. The fluorescence signal was fit to a sum of exponentials using the Levenberg-Marquardt algorithm as implemented in IGOR Pro software (Wavemetrics Inc.). In every case, the residuals were examined to confirm the absence of systematic errors. At most two exponentials were required to fit the kinetic traces. 95 4.2.5. Data analysis. (1) Chevron plot analysis. The natural logarithm of an elementary refolding or unfolding reaction rate generally has a linear dependence on denaturant concentration(264): o ‡ ln kij=ln kij +(m ij/RT)[denaturant] (4.4) o where kij is the rate constant from species i to j in the absence of denaturant, T is the temperature in Kelvin, and R is the universal gas constant. The slope m‡ij, which describes the free-energy dependence on denaturant concentration, is approximately proportional to the change in the buried surface area that occurs between the reactant and the transition state(265). In a two-state (un)folding reaction (N ↔ U), where no intermediates are involved, the observed rate constant is λ = kNU + kUN, (4.5) where λ is the measured rate constant, kNU and kNU are the unfolding and refolding rate constants, respectively. Under refolding conditions, the folding rate constants kUN are significantly larger than their reverse unfolding rate constants kNU, and therefore λ ≈ kUN. By contrast, under unfolding conditions, the unfolding rate constants kNU are significantly larger than their reverse folding rate constants kUN, and therefore λ ≈ kNU. Thus, the plots of ln λ as a function of denaturant concentration (chevron plot) for a two-state (un)folding protein adopt a V-shape with a minimum at which ln kNU = ln kUN, i.e., the number of molecules in the native state is equal to that in the unfolded state (Figure 4.13). 96 → )U Unfolded Zone (1 Transition Zone 0 Ln λ kUN Folding Zone N←U N↔U N→U (1) U→N λ(refolding) = kUN (2) N→U N U N→ (2) λ(unfolding) = kNU kNU0 [N] = [U] [denaturant] M Figure 4.13. Denaturant dependence of the folding and unfolding rates for a twostate (un)folding protein. λ is denoted to the observed decay constant; k is denoted to the microscopic rate constant. By assuming (un)folding mechanisms are conserved in different denaturant concentrations, kUNo and kNUo can be derived from linear extrapolation. However, if a protein folds according to a three-state regime with an intermediate on the direct pathway between native and unfolded states, i.e. N ↔ I ↔ U, a deviation from the V shape is expected (Figure 4.14). 97 (1) I→U (2) U→ I (1) I→U ln λ (4) I→ N λfast (unfolding) = kIU λ (3 → )N I (2) U→I λfast (refolding) = kUI (3) N→I λslow (unfolding) = kNI (4) I→N [denaturant] M λslow (refolding) = kIN Figure 4.14. Denaturant-dependence of the folding and unfolding rates for a threestate (un)folding protein. λ is denoted to the observed decay constant; k is denoted to the microscopic rate constant. The points represent the observed decay constant (λ). The solid and dash lines represent the fitted dependence of decay constants. Figure 4.14 represents the chevron plot for a three-state (un)folding protein. Under strongly refolding conditions ([denaturant] ≤ 2 M in this case), the refolding intermediate is well-populated. Both kUI and kIN can be resolved from the kinetic data in this region. Under mildly refolding conditions (2 M ≤ [denaturant] ≤ 4 M in this case), the equilibrium between U and I (U ↔ I) shifts to U, and the intermediate is no longer populated(141). Although the reaction approaches a two-state mechanism, the small population of the intermediate still contributes to the observed rate constant (λ); therefore, 98 the microscopic rate constant kIN cannot be obtained from the decay constant λ in this region. (2) Quantitative analysis for the compactness of folding intermediates. Chemical denaturation exerts its effect on partially unfolded intermediates or fully unfolded states through changing their solvent-exposed surface area(141). Such effects ‡ are reflected in the m ‡ ij value in Equation 4.4. A m ij value represents the change in the solvation free energy induced by denaturant(266), thus provides a quantitative measure of the average compactness. In a three-state folding regime, the compactness of the intermediate state relative to the unfolded state can be represented by an α value: αUI = m‡UI - m‡IU / (m‡UI + m‡IN - m‡NI - m‡IU) (4.6) where αUI = 0 represents an intermediate state with the same surface-burial as the unfolded state, whereas a αUI = 1 represents an intermediate state which is as compact as the native state. (3) Kinetic modeling. The exponential responses correspond to transitions between protein conformational states in which a free-energy barrier is crossed; therefore, the number of kinetic species can be deduced from the number of exponential responses. In general, a kinetic model with n kinetic species should produce a signal of n-1 exponential phases, although not all phases may be detected; conversely, the detection of p exponential phases implies at least 99 p+1 kinetic states. By assuming the persistence of kinetic mechanisms in different denaturant concentrations, kinetics that exhibits two exponentials under unfolding conditions indicates at least three states exist in ovPrP unfolding and refolding: the native state N, the unfolded state U, and an intermediate state denoted as I. The most general three-species kinetic model has the form N ↔ I ↔ U ↔N in which each species can interconvert with the other two. If the protein is initially entirely in the native state N (as in the unfolding experiments), the initial time derivative of this fluorescence signal is given by -dF/dt = Afast λfast + Aslow λslow = kNI (QN – QI) + kNU (QN – QU) where QN, QI and QU represent the quantum yields of the native, intermediate and unfolded species, respectively. Experimentally, -dF/dt is approximately zero under unfolding conditions. Since the fluorescence signal of ovPrP shows 4- to 6-fold increase upon unfolding, (QN – QU ) is non-zero. Therefore, the zero initial slope implies that kNU is negligibly small and QI ≈ QN. Since the quantum yield is highly sensitive to conformational changes, the rough equality of QI and QN suggest that the tertiary structure of Iα is native-like, particularly in the vicinity of the reporting tryptophan (the third α-helix). More generally, Iα should be structurally closer to the native state N than 100 to the unfolded state U. By the given similarity of N and Iα in their conformations, and the negligible conversion rate between N and U, I argue that the most plausible model for this observation is N ↔ I ↔ U. 4.3. Results. 4.3.1. Unfolding of ovPrPs. The unfolding kinetics of variants ALRQ, VLRQ, ALHQ, and ALRR were measured in GuHCl at pH 7, 15oC (Figure 4.15). Figure 4.16 shows the representative kinetic trace of ALRQ variant taken in the presence of 5.6 M GuHCl, along with the best fit to one or two exponentials. As shown in the figure, the second exponential improves the quality of the fit significantly. Since the unfolding of ALRQ, ALRR, and VLRQ variants exhibits two exponential phases under strongly unfolding conditions, the kinetic model must have at least one intermediate. The two decay constants, λslow (unfolding) and λfast (unfolding), increase monotonically with increasing GuHCl concentration, and can be fit to straight lines on a semilogarithmic plot (Figure 4.21; Table 4.1). The amplitude of the slower decay is larger and negative, as expected from the increase in the quantum yield of ovPrPs observed in equilibrium unfolding. By contrast, the amplitude of the faster exponential is positive; the mutual cancellation of these amplitudes produces a lag phase. At lower denaturant concentrations (but still above the unfolding threshold), one exponential term suffices to fit the unfolding kinetic traces satisfactorily. Unfolding experiments were also carried out in urea; however, no useful information can be obtained due to the narrow unfolded zone of ovPrP under this condition. 101 Relative fluorescence (1) 1.7 ALRQ (2) 1.8 (2) (3) 1.6 (3) (4) 1.5 (5) 1.6 (4) (5) 1.4 (6) 1.4 (6) 1.3 1.2 1.2 0 200 -6 400 0 600 Time, µs Relative fluorescence (1) ALRR -6 400 600 Time, µs (2) ALHQ 1.6 200 (3) 1.5 (4) 1.4 (5) 1.3 (6) 1.2 0 200 400 600 Time, µs Figure 4.15. Unfolding kinetic traces taken for ovPrPs in the presence of various concentrations of GuHCl at pH 7, 15oC with the best fit to one or two exponentials. (1): 6 M GuHCl; (2): 5.6 M GuHCl; (3): 5.2 M GuHCl; (4): 4.8 M GuHCl; (5): 4.4 M GuHCl; and (6): 4 M GuHCl. Solid lines represent one- or two-exponential fit to the kinetic traces. Fluorescence signal was normalized by the fluorescence of native ovPrPs. 102 -3 x10 -3 x10 Relative fluorescence 30 -30 1.7 20 0 -20 1.7 1.6 1.6 1.5 1.5 1.4 1.4 A 1.3 0 200 400 600 -6 B 1.3 0 200 400 600 -6 Time, µs Time, µs Figure 4.16. Kinetic trace taken for ALRQ variant in the presence of 5.2 M GuHCl at pH 7 and 15oC with the best fit to one (A) or two (B) exponentials. Solid lines represent one- or two-exponential fit to the kinetic trace. Residuals of the fits are shown above the kinetic trace. Fluorescence signal was normalized by the fluorescence of native ovPrPs. 4.3.2. Refolding of ovPrPs. Refolding measurements were carried out in GuHCl at pH 5 or 7 at 15oC. Figure 4.17 shows the representative kinetic trace taken for ALRQ variant in the presence of 1.45 M GuHCl at pH 5 with the best fit to one and two exponentials. As shown in the figure, two exponential terms are required to fit the data completely. A similar fitting situation was encountered for refolding traces of all the allelic variants taken at different GuHCl concentrations and at different pH values (Figure 4.18 and 4.19). Based on the control experiments (see Methods), the first phase with a time constant of ~10 µs is attributed to the artifacts caused by initial changes in the refractive index; only the slower decay represents ovPrP refolding. These refolding rate constants are plotted in 103 Figure 4.21 on a semilogarithmic plot; the deduced kinetic parameters are listed in Table 4.1. Other fitting parameters are plotted in Figure S1 (see Supplemental Materials). Refolding experiments were repeated under several alternative conditions: (1) urea at pH 7, 5oC (for ALRQ), (2) urea at pH 5, 15oC (for ALRR), and (3) GuHCl at pH 7, 5oC (for ALRR) to determine if the folding behavior of ovPrP changes with solution conditions. Since ovPrPs exhibit only one refolding phase under these conditions (Figure 4.20 and Figure S2), I conclude that ovPrP variants refold without a detectable intermediate under refolding conditions. Several factors might account for this observation, including (1) the intermediate is poorly populated under refolding conditions; (2) the second refolding phase (I → N) occurs too fast to be discerned; and (3) the second refolding phase has a negligible amplitude. 104 -3 1.2 5 0 -5 1.2 1.1 1.1 1.0 1.0 0.9 0.9 0.8 0.8 0.7 0.7 x10 -3 x10 Relative Fluorescence 40 20 0 0.6 0.6 (A) 0.5 0 (B) 0.5 200 400 600 0 200 400 600 -6 Time, µs Time, µs Figure 4.17. Refolding kinetic trace taken for ALRQ variant in the presence of 1.45 M GuHCl at pH 5 and 15oC with the best fit to one (A) or two (B) exponentials. Residuals of the fits are shown above the kinetic traces. Solid lines represent one- or twoexponential fit to the kinetic traces. Kinetic traces were normalized by fluorescence signal of unfolded ovPrPs. 105 1.0 1.0 Relative Fluorescence ARR ALRR ARQ ALRQ 0.9 0.9 0.8 0.8 2.5 M 2.4 M 0.7 2.0 M 1.9 M 1.5 M 0.6 0.7 2.2 M 1.9 M 1.5 M 1.2 M 1.2 M 0.8 M 0.8 M 0 200 400 x10 0 600 200 -6 400 x10 1.0 1.0 AHQ ALHQ VRQ VLRQ Relative Fluorescence 600 -6 0.9 0.9 0.8 2.3 M 0.8 0.7 1.9 and 2M 0.7 1.9 M 1.5 M 1.2 M 0.6 0.8 M 0.6 0 200 400 600 1.5 M 1.2 M 0.8 M 0 200 400 600 -6 -6 Time, µs Time, µs Figure 4.18. Refolding kinetic traces taken for ovPrPs in the presence of various concentrations of GuHCl at pH 7, 15oC with the best fit to two exponentials. Data were taken in 50 mM imidazole, pH 7. The fluorescence signal was normalized by the fluorescence of unfolded ovPrPs. Solid lines represent two-exponential fits to the kinetic traces. Corresponding GuHCl concentrations are indicated next to the traces. For the accuracy of data analysis, I included the artifact phase in the exponential fitting; however, for clarity, I show here only the signal from ovPrP refolding. 106 1.00 1.0 Relative Fluorescence ARR ALRR ALRQ ALRQ 0.9 0.95 0.8 0.90 0.7 0.85 2.3 M 2.0 M 2.3 M 2.0 M 0.6 1.8 M 1.8 M 0.80 1.2 M 0.8 M 0.75 0 200 400 1.5 M 1.2 M 0.8 M 0.5 0 600 200 600 x10 Time, µs 1.0 1.00 VRQ VLRQ AHQ ALHQ 0.95 0.9 Relative Fluorescence 400 -6 -6 0.90 0.8 0.85 2.4 M 0.7 0.6 0 200 400 600 2.0 M 0.80 1.8 M 1.5 M 1.2 M 0.75 0.8 M 2.3 M 2.0 M 1.7 M 1.4 M 1.1 M 0 -6 Time, µs 200 400 -6 x10 600 Time, µs Figure 4.19. Refolding kinetic traces taken for ovPrPs in the presence of various concentrations of GuHCl at pH 5, 15oC with the best fit to two exponentials. Data were taken in 50 mM sodium acetate, pH 5. Fluorescence signal was normalized by the fluorescence of unfolded ovPrPs. Solid lines represent two-exponential fits to the kinetic traces. Corresponding GuHCl concentrations are indicated next to the traces. Similar refolding behavior was observed for AFRQ variant, which is involved only in atypical scrapie (Figure S2). For the accuracy of data analysis, I included the artifact phase in the 107 Figure 4.19 (cont’d) exponential fitting; however, for clarity, I show here only the signal from ovPrP refolding. 108 0.9 0.90 ALRR Relative Fluorescence ALRQ 0.8 0.85 0.7 0.80 3.8 M 3.5 M 0.6 2.9 M 3.4 M 0.75 3.0 M (A) 0 200 400 600 2.3 M (B) 2.6 M 0 200 400 600 -6 Time, µs Time, µs 0.90 ALRR Relative Fluorescence 0.85 0.80 2.4 M 0.75 1.9 M 0.70 1.6 M 0.65 1.2 M (C) 0.60 0 0.8 M 200 400 600 -6 Time, µs Figure 4.20. Refolding kinetic traces taken for ovPrPs in the presence of various concentrations of urea or GuHCl with the best fit to two exponentials. (A): urea at pH 7, 5oC. (B): urea at pH 5, 15oC. (C): GuHCl at pH 7, 5oC. Fluorescence signal was normalized by the fluorescence of unfolded ovPrPs. Solid lines represent two-exponential fits to the kinetic traces. Corresponding denaturant concentrations are indicated next to the traces. For the accuracy of data analysis, I included the artifact phase in the exponential fitting; however, for clarity, I show here only the signal from ovPrP refolding. 109 Figure 4.21. Chevron plots of ALRQ (A), ALHQ (B), ALRR (C), and VLRQ (D) variants at pH 7, 15 oC. λ is denoted to the observed decay constant; k is denoted to the microscopic rate constant (see Methods for detail explanation). The points represent the observed decay constant (λ), The solid and dash lines represent the fitted dependence of decay constants of the slow and fast phases, respectively. (A) kNI kIU N → I ←U ← → kIN kUI (3 ) (2) (1) I→U λslow (unfolding) = kIU (2) N→I λfast (unfolding) = kNI ( 1) ln λ (4) GuHCl (M) 110 (3) U→I λslow (refolding) = kUI (4) I→N λfast (refolding) = kIN Figure 4.21 (cont’d) (B) (1) (1 )λ ln λ g) din fol (re slo w λ slow (u nf old in g) (2) Observed rate for N → U λ(unfolding) (2) Observed rate for U → N λ(refolding) GuHCl (M) kNI (C) kIU N → I ←U ← → kIN kUI λslow (unfolding, [GuHCl]<5.2M) N (1) (2) I→U λslow (unfolding, [GuHCl]≥5.2M) = kIU (3) N→I λfast (unfolding) = kNI (2) I→U (4) U→I λslow (refolding) = kUI (5) I→N λfast (refolding) = kIN N→ I (4 ) → Observed rate for N → U U→ (3 ) ln λ (5 )I (1) I GuHCl (M) 111 Figure 4.21 (cont’d) (D) O er bs ) U or ng di ln λ f te l fo (re ra w d ve λ slo → N GuHCl (M) 112 Table 4.1. Kinetic parameters for the folding of ovPrPs. k and λ are in s-1, τ is in µs, mij is in kJ·mol-1·M-1, and ∆G is in kJ·mol-1. (1) at pH 7: U↔I↔N (k: microscopic rate constant) ALRQ ALRR kUI0 τUI kIU0 τIU kIN0 τIN 5973.6 56.3 17761.99 22314.7 167.40 5341.7 534.1 1872.31 96954.8 187.21 kNI0 τNI T (oC) denaturant 15 44.81 3506.6 285.18 GuHCl 146.4 6830.60 15 10.31 GuHCl ALRQ ALRR mUI‡ mIU‡ mIN‡ mNI‡ T (oC) denaturant 1.862 -0.477 0.437 15 -1.194 GuHCl -1.077 0.949 -0.925 1.834 15 GuHCl ALRQ ALRR ∆GUI KIN ∆GIN ∆GUN [U]:[I]:[N] αUI T (oC) denaturant KUI 106.1 -11.16 6.36 -4.42 -15.58 1:106:675 0.77 15 GuHCl 10 -5.51 662.26 -15.54 -21.05 1:10:6623 0.42 15 GuHCl (2) at pH 7: U↔(I)↔N (λ: observed rate constant) λUN0 τUN ALHQ§ 7064.9 141.54 ‡ VLRQ 3754.1 266.38 ALRR 3597.5 277.97 λNU0 303.9 τNU 3290.56 mUN‡ mNU‡ -1.368 1.038 T (oC) denaturant 15 GuHCl -0.521 15 GuHCl -0.296 5 GuHCl ALRQ 4336.8 230.58 -0.634 5 urea §: An intermediate state was not observed for ALHQ variant. However, the absence of I is not conclusive. ‡: An intermediate state was observed for VLRQ variant. However, the present data do not allow the kinetic parameters to be deduced. 113 Table 4.1. (cont’d) (3) at pH 5: U↔(I)↔N (λ: observed decay constant) λUN0 ALRQ VLRQ ALHQ ALRR AFRQ ALRR τUN mUN‡ T (oC) denaturant 7321.7 7084.0 136.58 141.16 -1.129 -1.137 15 15 GuHCl GuHCl 5322.5 187.88 -0.590 15 GuHCl 4902.0 9008.3 6537.4 204.00 111.01 152.97 -0.498 -1.404 -0.438 15 15 15 GuHCl GuHCl urea 114 4.4. Discussion. 4.4.1. Kinetic intermediate is a plausible PrP Sc precursor. Under unfolding conditions, the kinetic traces exhibit a lag phase, meaning that the initial rate of change of fluorescence is roughly zero. Thus, the quantum yields of the intermediate and native states are similar. The tertiary structure of this intermediate is native-like, particularly in the vicinity of the reporting tryptophan (the third α-helix), since quantum yield is highly sensitive to conformational changes. In other words, this intermediate is largely α-helical. Hence, this intermediate is denoted as Iα. The relative populations of Iα in the susceptible variant (ALRQ) is about 10 times higher than that in the resistant variant (ALRR). The surface exposure of this intermediate also differs these two variants: the ALRQ variant shows ~77% of surface-burial in its intermediate relative to its native state, in contrast with ~42% for the ALRR variant. By assuming Iα is on the direct pathway between native and unfolded states (see arguments in Methods), the differences in the surface exposure suggests that Iα in the susceptible variant (ALRQ) is structurally closer to its native state, as compared to the resistant variant (ALRR). This correlation indicates that the more stable native-like structure of Iα Sc in the susceptible variants correlates with the propensity to convert to PrP . Consistent with this idea, structural conversion to PrP Sc has been suggested to proceed through an α- helix-rich species under oligomer-forming conditions(222-225, 261). 115 Based on these correlations, I argue that Iα is the PrP Sc precursor, and that the genetic modulation found in classical scrapie exerts its effect on the population and structure of this kinetic species. Similar situations have been observed in human familial prion disease, where the pathogenic mutations result in a pronounced increase in the population and the hydrophobicity of the kinetic intermediate of human PrP (huPrP)(219, 221), although the structural properties of this kinetic species were not able to be discerned. Herein, a model is proposed to explain amyloid formation in classical scrapie: Conformational folding of PrPC: N ↔ Iα ↔ U Sc Sc Sc Assembly of PrP oligomers: (PrP )n + Iα → (PrP )n+1 In the susceptible variants, the higher population of Iα gives them more opportunities for oligomerization, and its more stable structure makes the conversion to PrP feasible. Exogenous PrP Sc Sc more is required for misfolding in this model, since naturally occurring allelic variants of ovPrP, including those confer high susceptibility to classical scrapie, cannot induce the disease spontaneously(267-269). Native PrPC has been suggested to be the PrP Sc precursor that occurs in normal PrP folding. Zhang et al. proposed that Syrian hamster PrPC possesses a relatively open conformation, which is ready for the PrPC-to-PrP Sc conversion(46). Welker et al. suggested that oligomerization proceeds through a thiol/disulfide exchange reaction 116 between two native PrPs(239). Although such dimeric structure has been observed(43), a later report on in vitro conversion study seems to contradict to this model(240). Several studies have been carried out to determine the role of the native PrPs in prion diseases. Many of the mutants linked to human prion diseases do not differ from the wildtype huPrP in the thermodynamic stability in their native state(241-242, 270). In the case of classical scrapie, susceptible alleles even exhibit higher structure stability in their native forms as compared to the resistant ones(233, 237-238). Moreover, pathogenic or protective mutations associated with human or sheep prion diseases do not alter structures and backbone dynamics in the native state of PrPs(243, 270). Taken together, these studies suggest that polymorphisms modulate prion diseases by exerting their effects on an intermediate species, instead of the native form. Thus, a folding intermediate seems to be a more plausible PrP Sc precursor. 4.4.2. Intermediates observed under refolding conditions. The current study shows that ovPrPs refold with a time constant of ~100-300 µs without a resolvable kinetic intermediate under various refolding conditions, including two temperatures (5 and 15 °C), two pH values (5 and 7) and two denaturants (GuHCl and urea). Although an intermediate state was not observed directly, ovPrP refolding should proceed through an intermediate state, assuming the conservation of kinetic mechanism in different concentrations of denaturant(177, 207, 271). Such conserved folding and unfolding pathways are standard in proteins(177). 117 The present experiments contrast with previous studies. A kinetic study of huPrP was undertaken with 100 µs time resolution(219). Accumulation of an early intermediate with a time constant of ~50 µs and a rate-determining folding step with a time constant of ~700 µs was observed in urea at 5oC, pH 4.8 and 7(219). Since I could not resolve such an intermediate under refolding conditions, the folding kinetics of ovPrPs differ from human PrPs, possibly reflecting structural differences in their respective intermediates. Such species differences have been observed for the intermediate state in human and Syrian hamster PrPs(244). Instrumentation should not be a concern when comparing the present data with the previous huPrP study, since the same capillary mixing system was used for both experiments, with improved detector sensitivity in the current study. 118 Supplemental materials. Figure S1. Amplitudes and rate constants from the best fit to refolding kinetic traces. Rate constants were plotted on semilogarithmic scale. Traces were taken in (A) - (E): 50 mM sodium acetate with various concentrations of GuHCl , at pH 5 and 15oC; (F) - (I): 50 mM imidazole with various concentrations of GuHCl, at pH 7 and 15oC. (A) ALRQ PrP refolding phase Rate (S-1) Amplitude Artifact phase [GuHCl] M [GuHCl] M 119 Figure S1. (cont’d) (B) ALHQ PrP refolding phase Rate (S-1) Amplitude Artifact phase [GuHCl] M [GuHCl] M 120 Figure S1. (cont’d) (C) VLRQ PrP refolding phase Rate (S-1) Amplitude Artifact phase 121 Figure S1. (cont’d) (D) ALRR PrP refolding phase [GuHCl] M [GuHCl] M Rate (S-1) x103 Amplitude Artifact phase 122 Figure S1. (cont’d) (E) AFRQ PrP refolding phase Rate (S-1) Amplitude Artifact phase [GuHCl] M [GuHCl] M 123 Figure S1. (cont’d) (F) ALRQ PrP refolding phase Rate (S-1) Amplitude Artifact phase [GuHCl] M [GuHCl] M 124 Figure S1. (cont’d) (G) ALHQ PrP refolding phase [GuHCl] M [GuHCl] M Rate (S-1) Amplitude Artifact phase 125 Figure S1. (cont’d) (H) VLRQ Artifact phase Rate (S-1) Amplitude PrP refolding phase [GuHCl] M [GuHCl] M 126 Figure S1. (cont’d) (I) ALRR PrP refolding phase [GuHCl] M [GuHCl] M Rate (S-1) Amplitude Artifact phase 127 Figure S2. Amplitudes and rate constants from the best fit to refolding kinetic traces. Rate constants were plotted on semilogarithmic scale. Traces were taken in (A): 50 mM imidazole with various concentrations of urea, at pH 7 and 5oC; (B): 50 mM sodium acetate with various concentrations of urea, at pH 5 and 15 oC; (C): 50 mM imidazole with various concentrations of GuHCl, at pH 7 and 5oC. (A) ALRQ PrP refolding phase [GuHCl] M [GuHCl] M Rate (S-1) Amplitude Artifact phase 128 Figure S2. (cont’d) (B) ALRR Artifact phase Rate (S-1) x103 Amplitude PrP refolding phase [GuHCl] M [GuHCl] M 129 Figure S2. (cont’d) (C) ALRR Artifact phase Rate (S-1) Amplitude PrP refolding phase [GuHCl] M [GuHCl] M 130 Chapter 5.Role of the Disulfide Bond in Stabilizing the Folding Intermediate 5.1. Introduction. Several studies argue for the importance of the sole disulfide bond of PrP (Figure 5.1.A) in prion diseases. Previous reports indicate that the disulfide-free PrP Sc is destabilized, and exhibits a PrP -like conformation(251, 272). However, mutant PrPs in a reduced form, or with mutations that inhibit formation of a disulfide bond, are unable to be converted into PrP Sc (247-249). The reshuffling of this disulfide bond from Sc intramolecular to intermolecular forms has been proposed as a step in the PrPC-to-PrP conversion (239, 250-253), although a later study (240) suggests that disulfide bond reshuffling is not required for PrP Sc propagation. A very recent report has shown that the disulfide bond restricts the motion of the putative transmembrane domain (residues 111135) of Syrian hamster PrP, thus modulating the initiation of misfolding by regulating membrane binding(254). Despite intensive studies, the mechanisms underlying the influence of this disulfide bond on prion diseases remain elusive. Since a disulfide bond imposes conformational constraints on its neighboring residues and stabilizes its local structure(149), it may stabilize the formation of the PrP folding intermediate observed in the Chapter Four. If so, and if the intermediate is indeed the PrP Sc precursor, then the intramolecular disulfide bond may regulate the conversion to PrPC and PrP Sc through the same folding nucleus. To examine whether the interactions in the vicinity of the disulfide bond suffice to form the folding intermediate of PrP, I used a 131 peptide model flanking the disulfide bond to measure the local folding using circular dichroism (CD) and hydrogen/deuterium exchange (HDX). Folding in the vicinity of a disulfide bond has been observed in several other model peptides using this method(151- 154). Figure 5.1. Positions of the disulfide bond and the selected model peptides. (A) Ribbon representation of the structure of ovPrP (PDB entry: 1UW3). The disulfide bond is shown in yellow. P1 and P2 are shown in blue and red, respectively. (B) Sequence alignment of homologous PrP. The symbols α and β correspond to α-helix and β-sheet structure in PrPs, respectively. (A) Helix 1 Helix 2 P1 P2 Disulfide bond Helix 3 132 Figure 5.1. (cont’d) (B) βA Sheep Human Cat Mink Bovine Goat Deer Elk YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDI K I MERVVEQ YSNQNNFVHDCVNITI KQHTVTTTTKGENFTETDVKMMERVVEQ YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDMK IMERVVEQ YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDMKI MERVVEQ YSNQNNFVHDCVNITVKEHTVTTTTKGENFTETDI KMMERVVEQ YSNQNNFVHDCVNITVKQHTVTTTTKGENFTETDI K I MERVVEQ YNNQNTFVHDCVNITVKQHTVTTTTKGENFTETDI KMMERVVEQ YNNQNTFVHDCVNITVKQHTVTTTTKGENFTETDI KMMERVVEQ α1 βB ********************* Sheep MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDR Human MLGSAMSRPI IHFGSDYEDRYYRENMHRYPNQVYYRPMDE Cat MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ Mink MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYKPVDQ Bovine MLGSAMSRPLIHFGSDYEDRYYRENMHRYPNQVYYRPVDQ Goat MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ Deer MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ Elk MLGSAMSRPLIHFGNDYEDRYYRENMYRYPNQVYYRPVDQ α3 α2 ********************************************** ************************* P2 P1 α3 ************************ Sheep Human Cat Mink Bovine Goat Deer Elk MCITQYQRESQAYYQRGASVI LFSSPPVILLISFLIFLIVG MCITQYERESQAYYQRGSSMVLFSSPPVILLISFLIFLIVG MCVTQYQKESEAYYQRRASAI LFSSPPVILLISFLIFLIVG MCVTQYQRESEAYYQRGASAI LFSPPPVILLISLLILLIVG MCITQYQRESQAYYQRGASVI LFSSPPVILLISFLIFLIVG MCITQYQRESQAYYQRGASVI LFSPPPVILLISFLIFLIVG MCITQYQRESEAYYQRGASVI LFSSPPVILLISFLIFLIVG MCITQYQRESEAYYQRGASVI LFSSPPVILLISFLIFLIVG P2 133 5.2. Materials and methods. 5.2.1. Materials. The tris-(2-carboxyethyl) phosphine hydrochloride (TCEP) solutions were freshly prepared before use. NaH2PO4 was purchased from Mallinckrodt Baker (Phillipsburg, NJ). Guanidine hydrochloride (GuHCl) and sodium citrate were obtained from Invitrogen (Carlsbad, CA) and Spectrum Quality Products (Gardena, CA). Acetonitrile was obtained from EMD Chemicals (Gibbstown, NJ). Immobilized pepsin was obtained from Pierce (Rockford, IL). C18 ZipTip was purchased from Millipore (Bedford, MA). All other chemicals were obtained from Sigma-Aldrich. 5.2.2. Construction of peptide models. Multiple sequence alignment and structural alignment were performed to identify the key conserved residues stabilizing this local structure. The homologous sequences of ovine PrP were identified through several rounds of NCBI PsiBLAST (http://www.ncbi.nih.gov/BLAST/Blast.cgi), and the mature homologous sequences were aligned using ESPript server (http://espript.ibcp.fr/ESPript/cgi-bin/ESPript.cgi). Based on the alignment result, two peptide sequences involved in this disulfide bond were selected: (P1) YSNQNNFVHDCVNITVKQH (residues 169-187), and (P2) MERVVEQMCITQYQRE (residues 206-221) (Figure 5.1). 5.2.3. Preparation of peptide models. P1 and P2 were synthesized by SynPep Corporation. Purification was carried out with 5% - 100% acetonitrile gradient in the presence of 0.1% trifluoro-acetic acid in 90 minutes on Vydac C4 and C18 columns coupled with reversephase high-performance liquid chromatography (RP-HPLC) system (Waters) detected at 134 205 nm. Matrix-assisted laser-desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS) measurements were performed to identify the two model peptides (Figure 5.2). To prepare P1P1, P2P2, and P1P2 peptide pairs, roughly equal amount of purified monomers were mixed in a solution containing 5 M GuHCl, 0.2 M Tris-HCl, and 10 mM NaH2PO4 (pH 8), then air-oxidized overnight at room temperature. Separation and identification of these dimeric peptides were carried out by RP-HPLC purification and MALDI-TOF-MS measurement as mentioned above. The combination of RP-HPLC and MALDI-TOF-MS confirms that the P1P1, P2P2, and P1P2 peptides are indeed disulfide-bonded, since these methods cause non-covalent complexes to dissociate. 135 Figure 5.2. Identification of P1 and P2. (A) and (C): Chromatograms of P1 and P2, respectively. (B) and (D) MALDI mass spectra of P1 and P2, respectively. (A) Elute at 44% CAN Absorbance (AU) P1 Retention time (min) (B) Intensity P1 2032.05 kD m/z 136 Figure 5.2. (cont’d) (C) Elute at 45.6% ACN Absorbance (AU) P2 Retention time (min) (D) Intensity P2 2085.68 kD m/z 5.2.4. Hydrogen/deuterium exchange. (1) Technical background of hydrogen/deuterium exchange. Hydrogen/deuterium exchange (HDX) has been applied to probe hydrogen bonding and solvent accessibility in several proteins and peptides. In an unfolded protein or polypeptide, hydrogen exchange 137 is a relatively fast process. However, exchange of amide protons in a native form is often much slower due to these hydrogen being protected from the solvent by secondary and tertiary structure(134, 273). The rate of HDX reaction depends strongly on the temperature and the pH of the reacting system(274). The reaction is catalyzed by both acidic and basic conditions (Figure 5.3.A), with the reaction rate reaching its minimum at roughly pH 2.5(274) (Figure 5.3.B). By utilizing these two characteristics of HDX reactions, exchange can be slowed down significantly for MS measurements by lowering the pH to 2.5 and the temperature to 0oC. 138 Figure 5.3. Mechanism of HDX reaction and its reaction rates plotted as a function of pH for polyalanine. (A) HDX reactions at neutral pH involve base-catalyzed proton abstraction and acid-catalyzed deuterium transfer from solvent. (B) Figure was adopted from Smith, D.L. et al. (1997) J. Mass Spectrom. 32, 135-146. (A) A-H + OD- A- + D3O+ OD- A- + H-OD D2O + A-D log k (sec-1) (B) pH 139 (2) Experimental procedure. Samples were equilibrated in 20 mM phosphate buffer/H2O (pH 7.0) for one hour. 1 µL of equilibrated peptide solution (~100 pmol/µL) was added to 19 µL of 20 mM phosphate buffer either in D2O (pD 6.8) or in H2O (pH 7.0) at 4 oC. After 15 seconds, 19 µL of 0.1 M citrate buffer containing 0.5 M tris-(2-carboxyethyl) phosphine hydrochloride (pH 2.5) was added to quench the exchange reaction and reduce the disulfide bonds. Reduction of the disulfide bond makes it feasible to compare the exchange in a dimeric peptide to its monomeric peptide components. The sample was desalted using a C18 ZipTip (Millipore), and eluted with a matrix solution, followed by being spotted on a pre-chilled MALDI plate. The target plate was immediately dried in a desiccator for 1-2 minutes, then transferred to the MALDI mass spectrometer under vacuum. 5.2.5. Circular dichroism (CD) spectroscopy. Far-UV CD spectroscopy was carried out to detect the secondary structure of these five model peptides (P1, P2, P1P1, P2P2, and P1P2) in a buffer containing 10 mM NaH2PO4 and 10 mM NaF (pH 7.5). The CD spectra were acquired on a Jasco J-810 CD spectropolarimeter using a 1-mm path-length cell from 190 nm to 250 nm with a scanning speed of 1 nm/min at 4 oC. 5.3. Results. CD and HDX measurements were carried out to examine two hypotheses. First, if the local interactions around the disulfide bond suffice to form a PrP folding intermediate, the isolated P1P2 model peptide should adopt the same secondary structure as its 140 counterpart in a native protein, which is α-helical. Second, the association of any two peptides may increase the secondary-structure content of both peptides, since intramolecular hydrogen bonds within a peptide pair will have less competition from intermolecular hydrogen bonds between its monomeric peptide component and water(151). Therefore, the structure induced in the homodimeric P1P1 and P2P2 peptide pairs was also measured, in addition to the heterodimeric P1P2 pair. A control experiment was carried out on their monomeric components (P1 and P2) to determine whether the observations result from the interactions stabilizing by the disulfide bond. As shown in Figure 5.4, CD spectra of these model peptides (P1P1, P2P2, and P1P2) exhibit a single minima at near 200 nm, as expected for random-coil structure(275). A similar situation was observed for their monomeric peptide components (P1 and P2). This observation indicates that interactions within P1P2 peptide pair do not suffice to stabilize a folding intermediate, and that association of the monomeric peptides (P1P1, P2P2, and P1P2) does not provoke significant secondary-structure formation. Consistent with CD measurements, the covalent linkage of two model peptides does not protect them from HDX reactions, suggesting that stable hydrogen bonding does not form within the peptide pair (Figure 5.5). 141 Figure 5.4. CD spectra of the five peptide models. (A) and (B): Monomeric P1 and P2, respectively. (C)-(E): Disulfide-bonded P1P1, P2P2, and P1P2, respectively. Measurements were made in buffer containing 10 mM NaH2PO4 and 10 mM NaF at pH 7.5, ]MRW mdeg dmol-1cm2 4oC. 5 5 (B) (A) 0 -5 -5 -10 -10 [ θ 0 P1 -15 200 220 240 P2 -15 200 0 220 240 wavelength (nm) 0 (C) -4 -4 -8 (D) -8 [ θ ]MRW mdeg dmol-1cm2 wavelength (nm) -12 P1P1 200 220 P2P2 -12 240 200 wavelength (nm) 220 240 wavelength (nm) 142 [ ]MRW mdeg dmol-1cm2 Figure 5.4. (cont’d) 5 (E) 0 -5 θ -10 P1P2 -15 200 220 240 wavelength (nm) 143 Figure 5.5. HDX reactions for the model peptides. Mass shifts calculated for the monomeric P2 and dimeric P2P2 are 6.2 and 6.45, respectively. 2089.24 Relative Intensity (%) (A) P2 in H2O (B) Relative Intensity (%) 2095.44 P2 in D2O 1800.2 2600.4 3400.6 144 4200.8 Figure 5.5. (cont’d) Relative Intensity (%) (C) P2P2 in H2O (D) Relative Intensity (%) 2090.20 2096.65 P2P2 in D2O 145 5.4. Discussion. Hosszu et al. observed that a hyper-stable region involving helices 2 and 3 is retained around the disulfide bond in the C-terminal huPrP upon unfolding(276), suggesting its importance in initiating PrP folding. Presumably, the interactions stabilizing this structured core are essential to form a folding intermediate. Thus, a peptide pair was designed to assess the contribution of the disulfide bond and its local interactions to the formation of this folding nucleus. The data presented here indicate that interactions stabilized by the disulfide bond do not suffice to form this local structure. Although the peptide corresponding to helix 3 (residue 202-230) exhibits high intrinsic αhelical propensity, the peptide corresponding to helix 2 (residue 175-197) shows a strong propensity to adopt random-coil (N-terminus) and β-sheet (C-terminus) structure, instead of the α-helical structure exhibited by its counterpart in a full-length folded PrP. Consequently, non-local interactions between helix 2 and the rest of the protein are required to stabilize its helical conformation. Since the disulfide bond brings residues distant in the primary structure into proximity and strengthens its local interactions, it may play a determining role in helical formation in this region (i.e. helices 2 and 3) (Figure 5.1), particularly for helix 2. Such stabilizing effect of the disulfide bond on PrP has been reported previously(251, 272). Nevertheless, the present data indicate that interactions from other part of the protein, besides those stabilized by the disulfide bond, are required for forming helices 2 and 3. 146 Even if association of P1 and P2 does not suffice to form helical structure, an increased secondary-structure content might be expected in any dimeric peptide pair, as compared to its monomeric components. However, I found that the covalent linkage of these model peptides (P1P1, P2P2, and P1P2) does not provoke secondary-structure formation, stable hydrogen bonding, and burial of solvent-accessible surface area. Polymorphisms of ovine PrPs (ovPrPs) have been associated with differing susceptibility to classical scrapie, particularly at positions 136 (A/V), 154 (H/R), and 171 (Q/R). This genetic modulation has been shown to affect the structural stability of the folding intermediate in Specific Aim Two (see Chapter Four). Since these residues are distant from the disulfide bond, long-range interactions may contribute to the formation of this intermediate, if the folding nucleus is formed around the disulfide bond, as suggested by Hosszu et al.(276). This is in good agreement with the present data. 5.5. Future directions. The present data show that the local structure around the disulfide bond in PrP does not suffice for the formation of the folding intermediate; in other words, interactions involving other parts of PrP are required for folding nucleus formation. To sort out the contribution of interactions to intermediate formation, cleavable GST-tagged constructs were engineered to produce P1 and P2 in E. coli. Based on these constructs, peptides of different lengths flanking the disulfide bond can be easily produced. These constructs also allow the studies on the synthesized peptides from SynPeP Corporation to be repeated and confirmed. 147 5.5.1. Preparation of GST-tagged peptides. PGEX-4T-1 (Amersham Biosciences) was digested with BamHI and XhoI. Oligomers corresponding to the DNA sequences encoding P1 and P2 were designed and cloned into two digested PGEX-4T-1 vectors separately (Figure 5.6): P1: 5’-GAT CCT ATA GCA ACC AGA ACA ACT TTG TGC ATG ATT GCG TGA ACA TTA CCG TGA AAC AAC ATT GAT AAC-3’ and 5'- TCG AGT TAT CAA TGT TGT TTC ACG GTA ATG TTC ACG CAA TCA TGC ACA AAG TTG TTC TGG TTG CTA TAG -3' P2: 5'- GAT CCA TGG AAC GCG TGG TGG AAC AGA TGT GCA TTA CCC AGT ATC AGC GCG AAT GAT AAC -3' and 5'- TCG AGT TAT CAT TCG CGC TGA TAC TGG GTA ATG CAC ATC TGT TCC ACC ACG CGT TCC ATG -3' DNA sequencing was utilized to confirm the validity of these two constructs. 148 EcoR1 BamH1 pTac GST VP16C Xho1 TAND2 PGEX-4T-1 AMP BamH1 and Xho1 digestion DNA sequence encoding P1 or P2 is inserted into this site G Cloning site TCGAG C Figure 5.6. DNA encoding P1 and P2 were cloned into PGEX-4T-1. These plasmids were transformed into competent E. coli BL21 (DE3) cells. To express GST-tagged peptides, E. coli cells containing the desired plasmid were grown in LB medium containing 100 µg/ml ampicillin with shaking at 250 rpm at 37 oC. Based on induction studies, once the optical density reaches 0.6 – 1.0, expression is induced by adding isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM. Purification was carried out using glutathione resin, followed by ion-exchange and gelfiltration chromatography. These two GST-tagged peptides have been identified with SDS-PAGE and Western blotting (Figure 5.7). 149 GST-tagged peptides Figure 5.7. SDS-PAGE and Western blotting show the purified GST-P1 and GSTP2. Lane 1-3, SDS-PAGE (12%) analysis. Lane 4-5, Western blotting analysis using anti-GST as the primary antibody. Lanes 1: Invitrogen protein ladder. Lanes 2 and 4: GST-P1. Lanes 3 and 5: GST-P2. 5.5.2. Future work. Peptides flanking the disulfide bond with various lengths will be produced based on the engineered constructs described here. HDX and CD measurements will be carried out as described previously on disulfide-bridged peptide pairs and their monomeric components to assess whether the interactions within a peptide pair are able to form the folding nucleus. 150 Conclusions and Future Directions I observed a native-like intermediate in ovPrP refolding and unfolding. This intermediate is denoted as Iα because, being similar to the native species N, its secondary structure is largely α-helical. I found that the population and structure of Iα in the variants correlate with their differing propensities towards classical scrapie. Variants with higher classical scrapie propensity are characterized by higher populations and more stable tertiary structure in Iα. Thus, I argue that this partially unfolded, native-like intermediate Iα is a plausible PrP Sc precursor. Consistent with this model, an α-helix-rich species has been observed during the PrPC-to-PrP Sc conversion(222-225, 261). In agreement with the theoretical refolding model (5, 255), the present data suggest a Sc model to explain PrP propagation in classical scrapie: Conformational folding of PrPC: N ↔ Iα ↔ U Sc Sc Sc Assembly of PrP oligomers: (PrP )n + Iα → (PrP )n+1 where polymorphisms determine disease susceptibilities by modulating the population and structure of Iα. In the absence of denaturant (i.e. under folding conditions), the relative population of Iα in the susceptible variant (ALRQ) is about 10 times higher than that in the resistant variant (ALRR). The higher population of Iα in the susceptible variants gives them more opportunities for the PrPC-to-PrP 151 Sc conversion. Iα exhibits differing degree of surface exposure in the four variants: the ALRQ variant shows ~77% of surface-burial in its intermediate relative to its native state, in contrast with ~42% for the ALRR variant, indicating that the intermediate of the ALRQ variant is more folded and structurally closer to its native state, as compared to the ALRR variant. The correlation between the scrapie susceptibility of a variant and the structure of its intermediate suggests the conformation of Iα is crucial to PrP misfolding. According to my model, a poorly populated and/or weakly structured intermediate has difficulty in folding onto a preexisting PrP Sc seed. Therefore, variants with such intermediates are more resistant to scrapie. By contrast, this model predicts that a well-populated and/or mostly folded Iα is more likely to convert to PrP PrP Sc Sc and incorporate into a preexisting oligomer; therefore, variants with such Iα are more susceptible to scrapie. If indeed PrP Sc preexisting PrP is formed by absorption of a native-like intermediate, such as Iα, onto an Sc seed, then it is biophysically more plausible that the formed PrP Sc conformation should likewise have native-like structure. This picture seems to be consistent with the β-helical and spiral models(48, 50) of the PrP Sc conformation, where most of the helical structure is conserved during the PrPC-to-PrP Sc conversion. Such structural conservation is also in agreement with an antibody study, in which α-helices 2 and 3 in PrPC remain largely conserved in PrP Sc (49). Since the residues (136, 154, and 171) involved in the classical scrapie are distant from each other in the primary structure, I conjectured that genetic modulation exerts its effects 152 on this intermediate through long-range interactions. Consistent with this idea, a peptide model indicated that long-range interactions within PrPs are required for intermediate formation. Future directions. Initiation of protein folding depends on the diffusion-driven contact formation between different regions of an unfolded polypeptide chain. This process sets an upper speed limit on a folding reaction, and is determined by the solvent conditions and the protein sequence(148). Since formation of an intermediate is the first step for ovPrP folding, and long-range interactions likely contribute to the differing stability of the Iα in these allelic variants, the formation of long-range interactions in the initial stage likely differs among these allelic variants. In particular, these allelic variants should exhibit differing rates of contact formation in the unfolded state. To test this hypothesis, the rates of contact formation in the unfolded state can be measured using the tryptophan/cystine (Trp/Cys) triplet quenching method(277). In this method, Trp is excited to its triplet state by pulse laser, and its gradual quenching is monitored. Quenching occurs when the excited Trp encounters the disulfide bond (cystine) in a diffusion-limited process (FigureC1). By measuring the quenching rate, the dynamics of the unfolded protein can be characterized. The cystine has a much higher efficiency of quenching the Trp triplet than other elements of the protein; this, and the long lifetime of the Trp triplet excited state in the absence of quencher, allow the rate of Trp-Cys contact formation to be measured(278-280). 153 (A) Quencher (B) (C) (D) Phosphorescence Q/R(171) Q/R(171) R/H(154) R/H(154) Singlet Trp A/V(136) * A/V(136) * hv Triplet Trp (excited) Figure C1. Schematic illustration for quenching reaction. (A) Trp is in the singlet ground state. (B) Trp is excited to its triplet state. (C) The excited Trp contacts the quencher. (D) Trp goes back to the singlet state from the triplet state and releases phosphorescence. Preliminary data has been taken in the laboratory of Dr. Lisa Lapidus of the MSU Physics department. The loop length between the sole Trp and the sole cystine is 80 residues in the wild-type ovPrP (Figure C2). [‘Loop length’ is defined as the distance along the backbone (counted in residues) between Trp and the quencher.] The initial conditions were set to pH 5 at five different temperatures ranging from 0oC to 40oC. Two decay 154 phases with time constants of ~1 and ~15 µs were observed for wild-type ALRQ variant (Figure C3). Similar experiments will be carried out on the other three variants. (A) (His)6 tag MGHHHHHHGQ HHHHHH 102 GGSHSQWNKP 136 GGLGGYMLGS AMSRPLIHFG 171 182 YRPVDQYSNQ NNFVHDCVNI 217 IKIMERVVEQ MCITQYQRES SKPKTNMKHV AGAAAAGAVV 154 NDYEDRYYRE NMYRYPNQVY TVKQHTVTTT TKGENFTETD QAYYQRGA (B) C(182)-C(217) Loop length (80 residues) Figure C2. The positions of quenchers, Trp, and residues involved in sheep polymorphisms. (A) Primary structure of wild-type ALRQ variant. (B) Schematic representation showing the relative positions of Trp , the disulfide bond, and residues involved in sheep polymorphisms. 155 0.03 absorbance (A) (B) 0.02 0.02 0.01 0.01 τ1 = 1.4 µs τ2 = 16 µs 0 0 1e-07 1e-06 1e-05 1e-04 τ1 = 1 µs τ2 = 11 µs 1e-07 1e-06 1e-05 ln Time (sec) ln Time (sec) absorbance (C) (D) 0.02 0.01 τ1 = 0.96 µs τ2 = 11 µs 0 1e-07 1e-06 1e-04 τ1 = 0.09 µs τ2 = 10 µs 1e-05 1e-04 1e-07 ln Time (sec) 1e-06 1e-05 1e-04 ln Time (sec) 0.0175 absorbance (E) 0.0125 0.0075 0.0025 τ1 = 0.89 µs τ2 = 10 µs 1e-07 1e-06 1e-05 1e-04 ln Time (sec) Figure C3. Quenching reactions of the Trp/Cys pair. Experiments were carried out in 50 mM sodium acetate (pH 5) at (A) 0oC, (B) 10oC, (C) 20oC, (D) 30oC, and (E) 40oC. Time constants for triplet decay are indicated in the plots. 156 If more than one quencher is involved in the observed quenching reactions, interpretation of the data could become complicated. Therefore, additional control experiments were carried out to determine whether another quencher, such as the (His)6 tag upstream of Trp (Figure C2), contributes to the observed quenching reactions. Although the quenching rate of a free His is about 10 times slower than that of a free cystine at pH 5, the (His)6 tag may compete with cystine in the quenching reaction due to the shorter loop length of the Trp/(His)6 pair (9 residues) and the higher quencher concentration, as compared to the Trp/Cys pair. Since the quenching rate of a free His becomes about 100 times slower in its deprotonated form(280), a pH value higher than its pKa (~6) should be able to slow down the quenching reaction of Trp/(His)6 significantly. Thus, experiments mentioned above were repeated at pH 7, 8, and 9. A pH dependence was not observed in the range of pH 5 to 9, indicating that quenching reactions of Trp/(His)6 are negligible. 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