ABSTRACT Title of Dissertation: STRUCTURAL STUDIES OF NEP1/EMG1 RNA METHYLTRANSFERASE AND ATYPICAL RIO3 KINASE Seth Thomas, Doctor of Philosophy, 2011 Dissertation Directed By: Assistant Professor Nicole LaRonde-LeBlanc Department of Chemistry and Biochemistry Nucleolar Essential Protein 1 (Nep1) is required for small subunit (SSU) ribosomal RNA (rRNA) maturation and is mutated in Bowen?Conradi Syndrome. Although yeast (Saccharomyces cerevisiae) Nep1 interacts with a consensus sequence found in three regions of the SSU rRNA, the molecular details of the interaction are unknown. Nep1 is a SPOUT RNA methyltransferase, and can catalyze methylation at the N1 position of pseudouridine. Nep1 is also involved in assembly of Rps19, an SSU ribosomal protein, into the SSU. Mutations in Nep1 that result in decreased methyl donor binding do not result in lethality, suggesting that enzymatic activity may not be required for function, and RNA binding may play a more important role. To study these interactions, the crystal structures of the ScNep1 dimer and its complexes with RNA were determined. The results demonstrate that Nep1 recognizes its RNA site via base-specific interactions and stabilizes a stem-loop in the bound RNA. Furthermore, the observed RNA structure contradicts the structures of the Nep1-binding sites within mature rRNA, suggesting that the Nep1 changes rRNA structure upon binding. Finally, a uridine base is bound in the active site of Nep1, positioned for a methyltransfer at the C5 position, supporting Nep1's role as an N1- specific pseudouridine methyltransferase. In addition to the work completed with the Nep1 project, structural characterization of Rio3 Kinase is reported, as well as collaborative work in the structure determination of Ubiquitin and Ubiquitin complexes and Iodotyrosine Deiodinase. STRUCTURAL STUDIES OF NEP1/EMG1 RNA METHYLTRANSFERASE AND ATYPICAL RIO3 KINASE By Seth Thomas Dissertation submitted to the Faculty of the Graduate School of the University of Maryland, College Park, in partial fulfillment of the requirements for the degree of Doctor of Philosophy 2011 Advisory Committee: Professor Jason Kahn, Chair Professor Nicole LaRonde-LeBlanc Professor George Lorimer Professor David Fushman Professor John Moult, Dean?s Representative ? Copyright by Seth Thomas 2011 ii Acknowledgements I would first like to thank my advisor, Dr. Nicole LaRonde-LeBlanc for her support, guidance, advice, and patience with me during the last five years. I can easily remember as if it was yesterday, walking into the lab and seeing empty, black bench tops. Now, five years later, I am unable to express enough appreciation for her time and persistence, without which, this dissertation would not be possible. I would also like to thank all past and present members of the LaRonde Lab, especially Irene Kiburu, Chris Keller, and Dr. Aga Szyk for their daily conversations, help and support. Thank you also to all the members of my review committee. Dr. Kahn for being the chair of my committee and providing leadership. Dr. Lorimer for always having your door open and providing support and direction to the Center for Biomolecular Structure and Organization. Extended thanks to Dr. Fushman for not only being a member of my review committee but also providing me the opportunity to work and teach with members of his laboratory. Dr. Moult for dedicating his time as the Dean?s representative on the review committee. Thanks to the ARCS Foundation for providing support that allowed me to focus on research. Thanks to Dr. Patrick McTamney, Jennifer Adler and Dr. Steven Rokita for our successful collaboration with IYD and I wish everyone in the Rokita Lab success in the future. Working on IYD was an experience I will not forget for the rest of my life. I am a better all around scientist having worked with the Rokita Lab. iii I would like to extend my heartfelt gratitude to Mrs. Paulette Frazier. She is an extraordinary person that put up with me for five years. Without Paulette, I would have been completely lost at the University of Maryland. Simply put, she is amazing. Thank you to all the members of my family: my mom and dad, Dean, Belinda, and Nikki for their continual support and frequent motivational encouragement. Finally, I would like to thank my future wife Misty. From long, long distance to just long distance to our upcoming wedding, you have provided the support, motivation, and love I have needed. I love you and cannot wait to be your husband! iv Table of Contents Acknowledgements....................................................................................................... ii List of Tables ............................................................................................................. viii List of Figures .............................................................................................................. ix Abbreviations............................................................................................................... xi Chapter 1: Introduction ................................................................................................. 1 Chapter 2: Purification and Crystallography of AfNep1 and ScNep1........................ 16 Chapter 3: Description of AfNep1 and ScNep1 in the Free State .............................. 32 v Chapter 4: Nep1 in Complex with RNA..................................................................... 41 Chapter 5: Nep1 Discussion and Conclusions ............................................................ 59 Chapter 6: Purification of Rio3................................................................................... 71 Chapter 7: Optimization of RioK3.............................................................................. 81 vi Chapter 8: My Role in Ubiquitin .............................................................................. 100 Chapter 9: Crystal Structure of Iodotyrosine Deiodinase, a Novel Flavoprotein Responsible for Iodide Salvage in Thyroid Glands .................................................. 113 vii Bibliography ............................................................................................................. 137 viii List of Tables Table 2.1: RNA Oligonucleotides................................................................................25 Table 2.2: A. fulgidus Nep1 Semet Scalepack Logfile................................................26 Table 2.3: A. fulgidus Nep1 Native Scalepack Logfile...............................................27 Table 2.4: S. cerevisiae Nep1 Scalepack Logfile........................................................28 Table 2.5: S. cerevisiae Nep1 with One Molecule of RNA Scalepack Logfile...........29 Table 2.6: S. cerevisiae Nep1 with Two Molecules of RNA Scalepack Logfile.........30 Table 2.7: Nep1/Emg1 Data Collection and Refinement Statistics.............................31 Table 6.1: RioK3 Expressed Constructs......................................................................77 Table 7.1: RioK3 Short Scalepack Logfile..................................................................95 Table 7.2: Matthews Coefficient Logfile.....................................................................96 Table 8.1: Ubiquitin I13S Scalepack Logfile............................................................106 Table 8.2: Ubiquitin Data Collection and Refinement Statistics...............................108 Table 8.3: E2-25K Di-Ubiquitin Scalepack Logfile..................................................110 Table 9.1: ShelX Results from HKL2MAP Program Suite.......................................125 Table 9.2: Data Collection and Refinement Statistics...............................................126 Table 9.3: IYD-Holo Scalepack Logfile....................................................................127 Table 9.4: IYD-MIT Scalepack Logfile....................................................................128 Table 9.5: IYD-DIT Scalepack Logfile.....................................................................129 ix List of Figures Figure 1.1: 35S Pre-rRNA Processing ........................................................................ 11 Figure 1.2: 18S RNA Sequence and Structure............................................................ 12 Figure 1.3: Nep1 Catalyzed Reaction ......................................................................... 14 Figure 1.4: Eukaryotic Rio Family Cladogram and Domain Arrangement................ 15 Figure 2.1: AfNep1 and ScNep1 Crystals................................................................... 24 Figure 3.1: Nep1 Sequence Alignment....................................................................... 36 Figure 3.2: Overall Structure of ScNep1 and Dimer Interface ................................... 37 Figure 3.3: The AfNep1 Dimer................................................................................... 38 Figure 3.4: Conservation Maps on the Surface of ScNep1......................................... 39 Figure 3.5: Conservation Maps on the Surface of AfNep1 ........................................ 40 Figure 4.1: Overall Structure of the Nep1/RNA Complexes...................................... 51 Figure 4.2: Electrostatic Surface of Nep1/1RNA Complex........................................ 52 Figure 4.3: Nep1-RNA Interactions............................................................................ 53 Figure 4.4: Schematic of Interactions Between Nep1 and RNA ................................ 54 Figure 4.5: Conformation Changes Upon RNA Binding............................................ 55 Figure 4.6: Interactions with SAH.............................................................................. 56 Figure 4.7: Alignment of SAM Binding Pockets ....................................................... 57 Figure 4.8: Nep1 and RNA Binding Assays............................................................... 58 Figure 5.1: Proposed Nep1 N1-pseudouridine Methyltransferase Mechanism.......... 66 Figure 5.2: Possible Chaperone Role for Nep1 in S19 Loading................................. 67 Figure 5.3: RNA Structures in Nep1 Binding Sites.................................................... 68 Figure 5.4: AfNep1 and ScNep1 Aligned with Sequence Conservation .................... 69 Figure 5.5: Proposed Chaperone Role of Nep1 in Ribosome Biogenesis .................. 70 Figure 6.1: RioK3-Short Purification Gels ................................................................. 76 Figure 6.2: RioK3-Short Crystals ............................................................................... 78 Figure 6.3: Diffraction of RioK3-Short ...................................................................... 79 x Figure 6.4: MBP-RioK3-1-193 Purification ............................................................... 80 Figure 7.1: ?X? Method .............................................................................................. 93 Figure 7.2: Indexing of Riok3-Short........................................................................... 94 Figure 7.3: RioK3-Short MW Determination and LC MS/MS .................................. 97 Figure 7.4: RioK3-S199A-Short Crystals................................................................... 98 Figure 7.5: Limited Proteolysis of Riok3 Full-length................................................. 99 Figure 8.1: Ubiquitin I13S Overall View ................................................................. 107 Figure 8.2: Possible Crystals Containing E2-25K and Di-Ubiquitin Complex ....... 109 Figure 8.3: E2-25k and Di-Ubiquitin Complex ........................................................ 111 Figure 9.1: IYD structure.......................................................................................... 130 Figure 9.2: Structural Overlay of IYD and BluB...................................................... 132 Figure 9.3: IYD Sequence Conservation .................................................................. 133 Figure 9.4: IYD Active Site Electrostatics and MIT/DIT Alignment ...................... 134 Figure 9.5: Mapping Human Mutations onto the Structure of IYD ......................... 135 Supplementary IYD Figure S1 and Figure S2...........................................................136 xi Abbreviations ? Angstrom Af Archaeoglobis fulgidus Ala Alanine APS Advanced Photon Source Arg Arginine Asn Asparagine Asp Aspartic Acid ATP Adenosine-5?-triphosphate BIS-TRIS Bis(2-hydroxyethyl)amino-tris(hydroxymethyl)methane BME ?-mercaptoethanol CCD Charged Coupled Device CCP4 Collaborative Computational Project No. 4 COOT Crystallographic Object Oriented Toolkit Cys Cysteine Da Dalton ?C Degrees Celsius DIT Di-iodotyrosine DNA Deoxyribonucleic Acid EDTA Ethylenediaminetetraacetic Acid EM Electron Microscopy ePK Eukaryotic Protein Kinase FMN Flavin Mononucleotide Gln Glutamine Glu Glutamic Acid H Hours HEPES 4-(2-hydroxyethyl)-1-pipeazineethanesulfonic Acid HPLC High-Pressure Liquid Chromatography IPTG Isopropyl-?-D-thiogalactopyranoside IYD Iodotyrosine Deiodinase K Kelvin LB Luria Broth LC MS/MS Liquid Chromatography Mass Spectrometry/Mass Spectrometry Leu Leucine M Molar MES 2-(N-morpholino)ethanesulfonic Acid Mg Milligram MIT Mono-iodotyrosine mL Milliliter mM Millimolar MW Molecular Weight NEP1 Nucleolar Essential Protein 1 NECAT Northeastern Collaborative Access Team NMR Nuclear Magnetic Resonance xii PCR Polymerase Chain Reaction PEG Polyethylene Glycol PHENIX Python-based Hierarchical Environment for Integrated Xtallography RMSD Root Mean Square Deviation RNA Ribonucleic Acid RPM Revolutions Per Minute rRNA Ribosomal Ribonucleic Acid S Svedburg Unit SAD Single Wavelength Anomalous Diffraction SAH S-adenosylhomocysteine SAM S-adenosyl Methionine Sc Saccharomyces cerevisiae SDS-PAGE Sodium Dodecyl Sulface Polyacrylamide Gel Electrophoresis Se-Met Selenium Methionine Ser Serine SER Surface Entropy Reduction SPOUT SpoU and TrmD Family Name SSU Small Ribosomal Subunit TBE Tris/Borate/EDTA TEV Tobacco Etch Virus Thr Threonine Tris Tris(hydroxymethyl)aminomethane UV Ultraviolet Val Valine wHTH Winged Helix-Turn-Helix ?L Microliter ?M Micromolar 1 Chapter 1: Introduction 1.1: Overview Biochemistry is known as the chemistry of life. It is the study of structure- function relationships to determine cellular processes. Primary DNA sequences are transcribed into RNA, and then later translated into amino acids by the ribosome. The twenty possible natural amino acids are connected linearly by a peptide bond, forming what is known as a polypeptide chain. These polypeptide chains then take on a secondary structure, specifically determined by the sequence of amino acids in the chain, leading to the formation of alpha helices and beta sheets. The final proteins products are globular arrangements of secondary structure (also known as tertiary structure) stabilized into a low energy state. Once folded correctly into proper three- dimensional globular structures, proteins are tasked to carry out varied cellular processes essential to life of all organisms. At the core of protein biochemistry is the synthesis of the polypeptide chain. A ribonucleoprotein complex called the ribosome performs this process via a mechanism known as translation; this process translates the genetic code into amino acids leading to the synthesis of a fully functional protein. The eukaryotic ribosome is made up of ribosomal RNA (rRNA) and ribosomal proteins assembled into two mature subunits, one larger than the other. The large subunit (60S) and small subunit (40S) come together independently during translation to form the complete 80S ribosome. It takes a highly ordered, well-regulated scheme for the cell to manufacture rRNA and process it into its mature fragments. In eukaryotes, the initial 2 35S pre-rRNA (part of the 90S pre-ribosome) is chopped into smaller pieces through a series of cleavages (1-3). The final mature rRNA products are the 5.8S, 25S and the 18S (Fig. 1.1). There are estimated to be over 170 proteins that are involved in the maturation of the eukaryotic ribosome (3). These proteins carry out multiple different functions toward the final goal of producing a fully functional mature 80S ribosome, including direction of cleavage steps, rRNA modifications (such as methylation and pseudouridylation), transport, and chaperone function (3-9). The structural characterization of known ribosomal processing factors with unknown functions can provide a molecular analysis into their role in ribosome biogenesis. Currently, the only two ways to acquire molecular resolution better than 4.0 ? of protein structure is by x-ray crystallography or nuclear magnetic resonance (NMR). In the late 1950?s, myoglobin was the first protein for which a crystal structure was solved, credited to John Kendrew with a Nobel Prize in 1962 (10). Including the contribution from NMR, which saw its first protein structure in 1984 by Kurt W?thrich, there are greater than 65,675 protein structures deposited to the protein database (11, 12). Amazingly, just five short years ago, there were fewer than 38,000 submitted structures (12). X-ray crystallography methods have advanced so quickly the past half-decade that it is an arduous task to keep up with new technology. The widespread availability/affordability of high-throughput machinery, in house diffraction systems, and improved/new crystallographic software (Phenix/COOT/CCP4) are major factors for this recent success. Even with all the recent success, x-ray crystallography historically and currently has the same rate- limiting step, and that is the growth of suitable crystals that yield high-resolution 3 diffraction patterns. This manuscript will describe multiple projects, both successful and challenging, and will provide a glimpse into the exciting trials of x-ray crystallography. 1.2: Background: Nep1 Nucleolar essential protein 1 (Nep1) is a highly conserved protein required for ribosome biogenesis and found in organisms from archaea to humans. Nep1 was first identified in fission yeast (Schizosaccharomyces pombe) as a high-copy number suppressor of a mating deficiency caused by a mutation in the effector domain of Ras1 and thus named Mra1 (multicopy suppressor of Ras 1) (13). In yeast, Nep1 has also been designated as Emg1, for essential for mitotic growth 1 (14). Emg1 is required for 18S SSU rRNA maturation (14, 15). To release the yeast 18S SSU rRNA from the pre-rRNA, cleavage must occur at four sites, designated sites A0, A1, A2 and D (2, 3). Cleavage at site A0 and A1 truncates the 5?-end of the rRNA transcript, while cleavage at A2 results in separation of a 20S pre-rRNA species from the 5.8S and 25S. Final cleavage at site D produces the mature 18S SSU rRNA (Fig. 1.1). Depletion of Emg1 results in loss of cleavage at site A2, which results in an accumulation of a 21S SSU rRNA species (15). This is a result of cleavage at site A3, which normally happens after site A2 cleavage in the maturation of the 5?-end of the 5.8S rRNA species (2). 1.3: Introduction to the SPOUT Family Recently in 2008, work published by Taylor et al. and Leulliot et al., provided the first looks at Nep1/Emg1 with a dimeric structure from archaebacterium 4 Methanocaldococcus jannaschii and a monomeric structure from Saccharomyces cerevisiae (16, 17). Structural analysis of archaeal Nep1, of our own then unpublished work and the work by Taylor et al, revealed structural homology to the SPOUT family of RNA methyltransferases. The name SPOUT is derived from the family?s original methyltransferase members, SpoU and TrmD (18). The family is now much larger, but they all still have shared structural characteristics (16, 18, 19). The members of the family each contain a SPOUT methyltransferase domain and may contain additional inserted domains required for selectivity of RNA substrates. Members of the SPOUT family all contain a signature methyltransferase knotted fold, where S-adenosylmethionine (SAM) is the source for the methyl group used in the methyl transfer. Thus far, SPOUT methyltransferases have been implicated in the methylation of ribose 2?-OH (2?-O-ribose), guanine N1 (m1G), uridine C3 (m3U) and N3 of pseudouridine (m3?) (19, 20). Within the family, Nep1 proteins are most closely related to the TrmD and RsmE subfamilies, which catalyze m1G and m3U modifications, respectively (19). All SPOUT methyltransferases characterized thus far are dimeric, as is Nep1. 1.4: Nep1 is a Pseudouridine RNA Methyltransferase RNA-binding screens have indicated that Nep1 binds to the consensus sequence C/UUCAAC (21). This sequence is found in three regions of 18S rRNA in yeast including the stem loop of helix 31, bases 1189-1193, and part of the stem of helix 42, bases 1565?1570 (Fig. 1.2). In the RNA three-hybrid screen that identified this consensus sequence, all the binding sites were located in the unpaired region of a stem loop structure predicted by RNA secondary structure predictions (21). 5 Recent work has identified Nep1 as a pseudouridine N1-methyltransferase (22) (Fig. 1.3). Wurm et al. showed that both human and M. jannaschii Nep1 are capable of catalyzing methylation on pseudouridine-containing RNA sequences, especially C/U?CAAC. This sequence is the Nep1 site found in the stem loop of helix 31 of yeast 18S rRNA mentioned above (22). This pseuoduridine, located at position 1189, is hypermodified in yeast, with a 3-amino-3-carboxypropyl group on the N3 position and a methyl group on the N1 position (m1acp3?) (23). Nep1 is the first identified example of an N1-specific pseudouridine methyltransferase (22). 1.5: Nep1 Interactions with Rps19 and SnR57 Nep1 plays an unclear role in the loading of Rps19 into the SSU. Rps19 is a small ribosomal subunit (SSU) ribosomal protein found only in archaeal and eukaryotic ribosomes and is also known as a protein mutated in roughly 25% of cases of Diamond?Blackfan Anemia, a rare congenital disease characterized by defects in red blood cell development and predisposition to cancers (24, 25). Rps19 is an essential protein, and its depletion also results in the loss of cleavage at site A2 (16, 25-28). In yeast, studies performed by Buchhaupt et al., overexpression of Rps19 was shown to partially suppress the phenotype observed with loss of Nep1 function. Surprisingly, this suppression is enhanced by deleting snr57, the C/D box snoRNA that guides 2?-O-ribose methylation at 18S G1572 (21, 29). In fact, deletion of Nep1 is not lethal in an snR57 deletion mutant (21). Taken together, it appears that Nep1 aids in Rps19 loading and is essential in the presence of snr57. This function is either through direct interaction with Rps19, or alteration of the rRNA structure (21). Since the defects in rRNA processing observed in Nep1 and Rps19 depletion are identical, 6 it is likely that some of the ribosome biogenesis defect caused by loss of Nep1 is a consequence of improper Rps19 assembly. As mentioned above and shown in Figure 1.2, Nep1 consensus binding sequence C/UUCAAC is found in helix 31 (1188-1193 and helix 42 (1565-1570). Both these binding sites are near helix 41, where Rps19 was shown to bind using cryo electron microscopy (cryo-EM) reconstructions (21, 30). In addition, one of the sites, between bases 1565 and 1570, overlaps with the predicted binding site of snR57 (bases 1569?1583). 1.6: Nep1 Objectives The initial focus of my research was to determine the first structure of Nep1 and use structural analysis to understand its role in ribosome biogenesis. When, as mentioned, Taylor et al. reported the first Nep1 structure in 2008 (16) my research took on a new direction. Utilizing the Nep1 structure, we asked the following questions: 1. Is it possible to crystallize and solve the structure of Nep1 in complex with cognate target RNA, 2. If so, what drives specificity for the target RNA, and what is a plausible mechanism for RNA methylation. 3. What is the functional role of Nep1 in ribosome biogenesis? To address these questions, in chapter 2 I will describe the purification and initial crystallization and structure solution of Archaeaglobis fulgidus Nep1 (AfNep1) as well as the structure of Saccharomyces cerevisiae Nep1/Emg1 (ScNep1) solved via 7 seleno-methionine substitution and molecular replacement, respectively. Chapter 2 will also describe the methods utilized, including RNA oligonucleotide design, to acquire two x-ray structures of a complex formed between ScNep1 and a 14 base cognate RNA oligonucleotide. Chapters 3 and 4 will introduce all four structures determined from the experiments detailed in chapter 2 and provide structural analysis highlighting the active site, dimer interface and interactions with bound RNA. Conclusions to the Nep1 project will be presented in chapter 5. 1.7: Rio3 Background Kinases are involved in a wide range of cellular processes including and not limited to cell cycle regulation, DNA replication, transcription, and ribosome biogenesis (31, 32). Atypical protein kinases are defined as members of the kinase superfamily that possess phosphoryl transfer activity without significant sequence homology to canonical eukaryotic protein kinases (ePK). The RIO kinase family is classified within this group of atypical kinases because of the very low sequence homology to ePKs and the ability to phosphorylate serine residues (33, 34). Structural studies of Rio1 and Rio2 show the RIO domain is structurally similar to ePK?s kinase domain (35-37). The founding member of the RIO family (Rio1) has been shown along with Rio2 to be essential in ribosome biogenesis, more specifically the maturation of the 20S to the fully mature 18S ribosome (38-41). There are over 500 serine/threonine/tyrosine kinases in the human genome (42). Of those kinases, there are 13 families of atypical kinases, which represent 40 of the 500+ kinases (42). The functional subdomains of the 250-300 amino acid kinase domain are structurally conserved among the canonical eukaryotic protein 8 kinases (43). These subdomains include: nucleotide binding loop, known as the P- loop; hinge region, which interacts with the adenine of adenosine triphosphate; ?catalytic loop? containing catalytic residues asparagine and aspartate; activation loop or APE; metal binding loop; and peptide binding sub-domains on the C- terminus (39, 43-45). Structural studies of archaeal Rio1 and Rio2 have shown the absence of the activation loop sub-domain and C-terminal peptide binding sub- domains (35, 37). All metazoa carry the gene for the third member of the RIO kinase family. Rio3 is conserved in all multi-cellular eukaryotes. However, Rio3 is not present in yeast and archaea, unlike Rio1 and Rio2. Even though Rio1 and Rio2 share conservation through archaea and yeast, Rio3 and Rio1 are more closely related. With an overall sequence alignment (ClustalW) identity of 45% for the RIO kinase domain alone, Rio3 human (RioK3) and Rio1 human (RioK1) are highly homologous and come from the same phylogeny (Fig. 1.4A) (46, 47). Rio2 human (RioK2) and RioK3 are less related, with an overall alignment identity for the RIO domain identity of just 19%. Each member has a unique N-terminal and C-terminal domain/sub-domain. Rio2?s conserved N-terminal winged helix turn helix (wHTH) domain is known via archaeal Rio2 structures. However, eukaryotic Rio1 has an extended N-terminal domain of unknown structure not present in archaea. Rio3 contains a longer unknown domain (in comparison to Rio1) on the N-terminus, consisting of ~200 amino acids (48). Figure 1.4B shows the domain arrangement for the three members of the RIO family. 9 Recently, RioK3 has been identified as an oncogene in pancreatic ductal adenocarcinoma and was shown to significantly alter cytoskeletal architecture when overexpressed (49). The characteristic phenotype is the formation of lamellipodia, an appendage like protrusion commonly seen in migrating and invasive cells (50). Overexpression of RioK3 was also shown to promote this invasive cell phenotype by the activation of a small GTPase known as Rac (49). It is still uncertain as to how RioK3 activates Rac and whether RioK3 requires kinase function, either through autophosphorylation or through phosphorylation of a currently unknown substrate. The significant uncertainties of RioK3 combined with its characterization as an oncogene make RioK3 a prime target for further structural and biochemical studies. 1.8: Rio3 Studies A major portion of my tenure at the University of Maryland has been working on the purification and crystallization of Rio3. There have been successes in purification protocols that will be detailed in chapter 6. For every success in the Rio3 project, there has been an equal roadblock at protein stability, initial crystal screening, as well as in the optimization of diffraction from acquired crystals. Chapter 7 will provide details to the exhaustive approach to improve crystal quality and diffraction. Despite my inability to produce suitable crystals prior to the end of my time here, the work described will lay the foundation for future success in the structural and biochemical characterization of Rio3. 10 1.9: Additional Studies In addition to the previously described work, I have participated in a variety of collaborative projects. Chapter 8 will describe work performed with Dr. David Fushman, a co-member of the Center for Biomolecular Structure and Organization, which focused on Ubiquitin and Ubiquitin in complex with E2-25K. Finally, chapter 9 contains the published work I completed in collaboration with then senior graduate student Dr. Patrick McTamney and his advisor, Dr Steven Rokita, at the University of Maryland. The collaborations were exciting and provided experiences that guided me to life lessons in protein crystallography and collaboration formation. 11 Figure 1.1: 35S Pre-rRNA Processing Thin arrows indicate specific cleavage sites. Cleavage at A0 and A1 occur first off the 5?-end. Next, cleavage at site A2 results in the separation of the 20S pre-rRNA, which later becomes the final mature 18S rRNA. Depletion of Nep1 (loss of cleavage at site A2) results in accumulation of the 21S pre-rRNA portion depicted in green, caused by the continuation of normal cleavage at site A3. 12 Figure 1.2: 18S RNA Sequence and Structure Based on the secondary structure prediction for S. cerevisiae 18S rRNA from the Comparative RNA Web Site (www.rna.ccbb.utexas.edu). Selected regions of secondary structure are colored orange, cyan and green for helix 31, 41 and 42 respectively. Red color indicates Nep1 consensus sites. The snR57-binding site is 13 highlighted in yellow. Purple denotes parts of rRNA within 4?? of S19 (shown as blue cartoon). Asterisk indicates the site of interaction between helix 41 and 42. Small circles around bases represent bases with post-translational methylation modification (51). 14 Figure 1.3: Nep1 Catalyzed Reaction Nep1 catalyzes the transfer of the methyl group (circled in red) from S-adenosyl methionine (SAM) to the N1 position of pseudouridine (?) resulting in N1-methyl psuedouridine and left over S-adenosylhomocysteine (SAH). 15 Figure 1.4: Eukaryotic Rio Family Cladogram and Domain Arrangement A) Cladogram showing evolutionary relationships (47). PKA (c-AMP dependent protein kinase A) represents canonical eukaryotic kinase. B) Rio family members include a conserved atypical Rio kinase domain, with Rio1 and Rio3 being more similar (light grey used for Rio1 and Rio3 to show similarity vs dark grey for Rio2). Each family member has a unique N-terminal domain (Rio1 and Rio3 both unknown folds; Rio3 predicted winged helix turn helix (wHtH) motif and unique C-terminal residues. 16 Chapter 2: Purification and Crystallography of AfNep1 and ScNep1 2.1: Purification Protocol for AfNep1 and ScNep1 Archaeoglobis fulgidus was originally discovered in 1987 deep in an Italian thermal vent (52, 53). This hyperthermophilic organism was later discovered to optimally thrive at 76?C (54). In order to survive, A. fulgidus has evolved to prevent the denaturing of proteins at high temperatures. The elevated thermal stability of A. fulgidus proteins allows the design of a purification protocol for their expression in E. coli that utilizes wide differences in thermal denaturation properties. The full-length Nep1 gene was PCR amplified from A. fulgidus and S. cerevisiae genomic DNA (ATCC) and subcloned into a plasmid vector (pDest527) containing an N-terminal 6X histidine tag followed by a Tobacco Etch Virus (TEV) protease cleavage site (plasmids obtained from Protein Expression Laboratory, SAIC, Frederick, MD) (55). These constructs were transformed into E. coli Rosetta?-DE3- pLysS cells (Novagen). Expression cultures were grown in LB media at 37 ?C and induced with 1 mM isopropyl-beta-D-thiogalactopyranoside (IPTG) at OD600 of 0.6 for 4 hours for AfNep1. For ScNep1, cultures were grown at 37?C until OD600 of 0.6, then cooled to 20?C and induced with 1 mM IPTG for overnight expression. After expression, the cells were harvested by centrifugation. For AfNep1, cells were resuspended in 50 mL of 50 mM Tris pH 8.0, 300 mM NaCl, 0.2 % ?- mercaptoethanol, 0.2 X Bugbuster? (Novagen) and 0.1 mg/ml DNase1 (Roche) per liter of expression culture and stirred at room temperature. After 30 minutes, the lysate was transferred to centrifuge tubes and placed in a 75 ?C water bath for 15 minutes to denature the E. coli proteins and then immediately centrifuged for 20 17 minutes at 18000 rpm (rotor: Beckman 45 Ti) in an ultracentrifuge to remove insoluble/denatured material. For ScNep1, cells were resuspended in 50 mL of 25 mM Tris pH 8.0, 200 mM NaCl, 0.2% ?-mercaptoethanol, 0.2 X Bugbuster? and 0.1 mg/ml DNase1 (Roche) per liter of expression culture and stirred at 4?C. After 45 minutes, the lysate was transferred to centrifuge tubes and centrifuged for 20 minutes at 18000 rpm (45 Ti). The supernatants were passed through a 0.22 ?M filter and loaded onto a 5 ml HisTrap HP column (GE Healthcare) equilibrated in buffer containing 50 mM Tris pH 8.0, 300 mM NaCl and 0.2 % ?-mercaptoethanol for Nep1 and 25 mM Tris pH 8.0, 200 mM NaCl and 0.2% ?-mercaptoethanol for ScNep1. The bound AfNep1/ScNep1 was washed with 100 mM imidazole in the respective equilibration buffer and then eluted with a gradient from 100 mM to 1 M imidazole in equilibration buffer in 20 column volumes. The fractions containing AfNep1/ScNep1 were pooled and dialysed overnight into 25 mM Tris pH 8.0, 300 mM NaCl, 1 mM EDTA and 0.2 % ?-mercaptoethanol for AfNep1 and 25 mM Tris pH 8.0, 200 mM NaCl, 1mM EDTA and 0.2 % ?-mercaptoethanol for ScNep1. In the case of AfNep1, attempts made to remove the tags by TEV protease cleavage were unsuccessful. The tag was cleaved in ScNep1. The resulting protein contained a single extra glycine residue at the N-terminus. The cleaved ScNep1 protein was passed over a 5 ml HisTrap HP column (GE Healthcare) a second time to remove uncleaved protein and TEV protease (55). The proteins were concentrated to 5 - 10 mg/ml and further purified to >99% by size exclusion chromatography using a Superdex 75? column (GE Healthcare) equilibrated in 20 mM Tris pH 8.0, 300 mM NaCl, 1 mM EDTA and 0.2% ?-mercaptoethanol for Nep1, and a Superdex 200? column (GE Healthcare) 18 equilibrated in 20 mM Tris pH 8.0, 200 mM NaCl, 1 mM EDTA and 0.2% ?- mercaptoethanol for ScNep1. The protein was concentrated again to 8 mg/ml for AfNep1 and 20 mg/ml for ScNep1. This preparation was stored at 4?C until crystallization screening. Se-Met AfNep1 was expressed in minimal media with all amino acids supplemented except for methionine, which was replaced by seleno- methionine (Se-Met). Reducing agent (0.2% ?-mercaptoethanol) was added to Se- Met protein immediately prior to crystallization screening. 2.2: Crystallography 2.2A: RNA Oligonucleotide Design Table 2.1 shows the RNA oligonucleotides of varying lengths and sequences purchased HPLC-purified from IDT. At the time of oligonucleotide design, the only information available was the identification of the ScNep1 consensus binding site, C/UUCAAC, and RNA secondary structure prediction of the target sequence (21, 51). It was not known that pseudouridine was required for methylation by ScNep1. The initial design utilized different lengths of the identified sequence (oligo #1, #3 and #4) and extending the consensus sequence with Watson-Crick G-C base pairs to stabilize the predicted stem-loop secondary structure (oligo #2). ScNep1 was mixed at a 1:1 molar ratio with the RNA oligonucleotides resuspended in 10 mM Tris pH 8.0. The mixtures were analyzed by gel electrophoresis on a 6% TBE-polyacrylamide gel and visualized by UV after ethidium bromide staining to determine stoichiometric complex formation. The 14- base RNA oligo #2 containing the sequence 5?-GGGCUUCAACGCCC-3? was the 19 best at producing stoichiometric complexes. Oligo #3 containing 5?- CUCUUCAACGAG-3? also showed good complex formation, however no crystals were obtained with RNA present in the active site. 2.2B: Crystallization Conditions Initial crystallization conditions were obtained through utilization of several sparse matrix screens (Qiagen, Emerald Biosystems Inc., Nextal, Hampton Research) with the sitting drop vapor diffusion method. Crystals of the AfNep1 protein were obtained from drops containing 3-4 M sodium chloride, 100 mM HEPES buffer pH 7.0 ? 8.0 placed over 1 mL reservoirs. Crystals of free ScNep1 were obtained from several conditions, including 50 mM BIS-TRIS, pH 6.5, 50 mM ammonium sulfate, 30% pentaerythritol ethoxylate (15/4 EO/OH) and 20% glycerol, used for data collection. The ScNep1/1RNA complex crystallized in 100 mM HEPES, pH7.0 and 10% PEG (polyethylene glycol) 6000 and crystals of the ScNep1/2RNA complex were obtained in 50 mM MES, pH 6.0, 15 mM magnesium sulfate and 7% PEG 4000. Diffraction quality crystals were obtained after 4-6 days at 20 ?C. (Fig. 2.1) 2.2C: Data Collection Crystals were flash frozen in mother liquor supplemented with 20% glycerol for all crystals. Cryogenic conditions are used to protect crystals from radiation damage during data collection as well as to prevent icing during cryo-preservation. Diffraction data for the native and Se-Met were collected at 100 K at the NECAT beamline at the Advanced Photon Source (APS), Argonne, Illinois. Data from the Se- Met crystals were collected at one wavelength (peak). All data were integrated with 20 DENZO and merged with Scalepack within HKL2000 (56). Tables 2.2-2.6 show condensed Scalepack output log files for all four structures, including the Se-Met log file from data used for phasing (Table 2.2). Split into shells of resolution, the log files are used as a tool to understand the quality of the data after being integrated and merged. Specifically, the log files presented in Tables 2.2-2.6 show redundancy and completeness of the shells, and also provide data quality statistics. Understanding these output files is crucial in determining not only inborn problems in the diffraction quality resulting from poor crystals, but also the quality of the choices made during the integration and merging process. 2.2D: Structure Determination X-rays behave as both a particle and a wave. The ability to ?transform? diffraction from the x-ray beam after hitting a protein crystal into an electron density map via the electron density equation (Eq. 1) can only be accomplished when three variables are known. Equation 1: Electron Density (?) Equation. Red represents the structure factor amplitude. Green represents the indices, hkl, of the unit cell (position). Blue represents the phase term of the electromagnetic wave. V is the unit cell volume. 21 The three terms shown in color in Equation 1 represent the location of the diffraction spots (known as the hkl?s or indices colored in green) in reciprocal space, intensity of the diffraction (known as amplitude colored in red), and finally the phase (?) of the wave colored in blue. A two-dimensional detector can only acquire measurements for position and amplitude. The inability for the detector to simultaneously measure the phase of the wave is known as the ?phase problem?. In order to acquire a three dimensional structure from a two dimensional detector a method for acquiring phases must be addressed. The availability of adjustable wavelengths at synchrotrons has provided a great method for acquiring phases through single wavelength anomalous diffraction (SAD) (57, 58). Phasing via SAD methods requires the presence of anomalously diffracting atoms in the structure. This can sometimes be accomplished using the native sulfur atoms (as you will see in chapter 9). However, this is generally more challenging and requires very high resolution (2.0 ? or less) and clean diffraction patterns, including a robust crystal that withstands radiation damage (59). Heavy atoms can also be introduced artificially, either through replacement during expression or by heavy atom soaking (60, 61). The most common method of heavy atom incorporation is achieved with replacement of methionine residues with selenomethionine (Se-Met). This replacement should not cause any changes to the crystal form (space group and unit cell), and therefore should be ?isomorphic? to the native crystal. The Patterson function is used to take advantage of the heavy atom?s absorption edge and anomalous diffraction associated with collecting data at that particular wavelength and determines the location of the introduced heavy atoms. Once the locations are determined, phases for the heavy 22 atom substructure can be determined and the first electron density map can be viewed. From this point, there are many options for model building. Another option for phasing is called molecular replacement and is now the most common method of phasing (~75%) (12). Molecular replacement is more of a brute force method that rotates and translates a known homologous model (>20% identity) looking for a similar match to the unknown model. Modern search programs, such as Phaser and MOLREP, perform this search by independent rotational and translational searches. First, a proper rotational orientation for the search model is determined in the asymmetric unit by aligning the Patterson maps for the search model with the experimental data. After the correct rotational orientation is determined, the search model undergoes translational movements to better align the Patterson maps. If the resolution is high enough and with today?s impressive processing power in computers, in a relatively short time, initial phases can be determined by this rotation and translation search method. Once the phases are refined, an electron density map can be viewed. The AfNep1 structure was solved using single wavelength anomalous diffraction (SAD) data with the Se-Met protein crystals, collected to 2.0 ?. The AutoSharp program suite was used to obtain the phases using the SAD method, apply solvent flattening and density modification to the initial electron density map, and perform automatic model building with wARP (62, 63). The complete model was finalized by rebuilding in COOT and refinement with REFMAC5 (64, 65). The model was then used to phase the native data set using MOLREP as part of CCP4i program suite (66-69). The ScNep1 structures were solved by molecular replacement in 23 MOLREP or Phaser using PDB ID 2V3J (17, 70). All models were subjected to several rounds of building in COOT and refinement using REFMAC5 or PHENIX (64, 65, 71, 72). Translation libration screw-motion (TLS) groups were determined using the TLSMD server (73). Refinement statistics are provided in Table 2.7. The coordinates and structure factors have been deposited in the Protein Data Bank (accession codes 3O7B for AfNep1, 3OII, 3OIJ and 3OIN for ScNep1 free dimer, 1RNA complex and 2RNA complex respectively). 24 Figure 2.1: AfNep1 and ScNep1 Crystals A) AfNep1 crystals containing Se-Met, (B) ScNep1, (C) ScNep1 with one molecule of RNA and (D) ScNep1 with two molecules of RNA. 25 Table 2.1: RNA Oligonucleotides Green represents consensus yeast consensus sequence (21). Red coloring indicates mutations from native yeast 18S RNA. ? Oligo 1 - 14 base native yeast 5?-GCU CUU CAA CGA GG-3? ? Oligo 2 - mutated yeast *crystallized with Emg1* 5?-GGG CUU CAA CGC CC-3? ? Oligo 3 ? 12 base native yeast 5?-CUC UUC AAC GAG-3? ? Oligo 4 ? 8 base native yeast 5?-CUU CAA CG-3? 26 27 28 29 30 31 32 Chapter 3: Description of AfNep1 and ScNep1 in the Free State 3.1: Overview Structural analysis of AfNep1 and ScNep1 supported previously reported structures that Nep1 does indeed belong to the SPOUT family of methyltransferases and contained all of the characteristic family traits, including the knotted methyltransferase fold (16, 17). This chapter will present a structural overview of Nep1, highlighting Nep1?s relationship to the SPOUT family of methyltransferases. Secondly, all SPOUT family proteins have been shown to be dimers (19), as Nep1 is not an exception, a description of the dimer interface combined with comparisons between AfNep1 and ScNep1 will be presented in detail. Research on the characterization of the Nep1 dimer interface was progressing very quickly and was near write up and submission, however, the competition was fierce and the direction of the project had to adapt (Chapter 4) after other laboratories released initial structural descriptions and also supported the presence of an obligate dimer (16, 17). 3.2: Basic Structural Description The overall structure of the Nep1 protein, shown in Figure 3.1, consists of a central ?-sheet surrounded by ?-helices on either side, characteristic of a Rossmann fold (16, 17). The central ?-sheet is comprised of six ?-strands, five of which are parallel to each other and one anti-parallel (?3), with an additional two loosely ordered strands completing the sheet (?4, ?5). The order in which sequential ?- strands form the ?-sheet is ?5-?4-?3-?2-?1-?7-?6-?8. The helices within the 33 Rossmann domain are located between ?1 and ?2 (?A, ?Ab, ?d?), ?3 and ?4 (?c, Hc) and after ?8 (?E). Two additional helices, Hb? and ?a? are present in a small Nep1 specific 40 amino-acid insertion between ?1 and ?2 (residues 48?88). This insertion and the orientation of ?a? is characteristic of Nep1 SPOUT methyltransferases, and also exhibits the most difference between the archaeal and yeast structures. This domain is moderately conserved based on sequence comparison with eukaryotic Nep1 sequences (Fig. 3.1). Electron density was observed for all the residues except 1?27 for both monomers, 56?64 for monomer A and 54?65 and 153?157 for monomer B, of ScNep1, and for all but the last three for AfNep1. In both models, density was observed in the SAM-binding pocket into which SAH was modeled. 3.3: Dimer Formation Although the structure of Nep1 from yeast has been previously reported, the deposited structure (PDB ID 2V3J) was monomeric (17). In the crystal structure of ScNep1 bound to SAH presented here, the molecule is dimeric. In both archaea and yeast, a large interacting surface is observed between Nep1 molecules that buries surface area in excess of 845 ?2 for AfNep1 and around 1475 ?2 for ScNep1. The dimer interface is formed primarily between residues on helices ?A and ?E and the N-terminal end of ?c'. The interactions form in a 2-fold symmetric interface that can be seen in Figure 3.2A and Figure 3.2B (Fig. 3.3 for AfNep1). The residues at the dimer interface of ScNep1 are highly conserved among eukaryotes, but less so among archaeal organisms (Figs. 3.4 and 3.5). The ScNep1 interface consists of a mixture of electrostatic interactions and hydrogen bonds on the edges and a few 34 hydrophobic interactions at the center, and several water molecules were observed in the charged regions of the interface. This shows a less tight dimer interface than that observed for the AfNep1 interface, which contains a stabilizing disulfide bond, commonly used to stabilize protein structure in thermophiles (Fig. 3.3) (74). Interestingly, in ScNep1, the side chain of the C-terminal Leu 252 forms hydrophobic packing interactions at the interface and the carboxyl group at the end of the chain forms a hydrogen bond with the side chain of Ser 228 and Asn 229 and the backbone amide nitrogen of Asn 229 of the other monomer (Fig. 3.2B and 3.2C). Therefore, the C-terminus participates in dimer formation. In the previously reported structure of ScNep1, the construct used contained a C-terminal histidine tag that may have disrupted this interaction and weakened the dimer to some extent, resulting in monomers in the crystals (17). 3.4: Chapter Summary The conserved dimer interface between the monomers of both AfNep1 and ScNep1 is seen throughout all members of the SPOUT family (19). While we were working on exploring this dimer interface, other research laboratories published work confirming Nep1?s conserved dimer formation. The focus of research was no longer to determine a novel structure, but to determine Nep1?s involvement with target 18S RNA. Focus shifted to crystallization of Nep1 in complex with RNA. 35 36 Figure 3.1: Nep1 Sequence Alignment Sequence alignment from of Nep1 from S. cerevisiae (NEP1_YEAST), Homo sapiens (NEP1_HUMAN), Drosophila melanogaster (NEP1_DROME) and A. fulgidus (NEP1_ARCFU). Yellow, red and blue dots indicate residues involved in dimerization, RNA binding and SAM binding respectively. Secondary structure for yeast (top) and A. fulgidus (bottom) shown as open bars for ?-helices and arrows for ?-strands and numbered according to Leulliot et al. for yeast, and Taylor et al. for A. fulgidus (16, 17). 37 Figure 3.2: Overall Structure of ScNep1 and Dimer Interface A) Overall view of the ScNep1 dimer. RNA binding and catalytic regions of the molecules are colored red and dark blue respectively. The molecules of SAH are shown in sticks. Secondary structure elements are labeled according to Leulliot et al. (17). B) Front view of the dimerization interface of ScNep1. Residues are labeled with chain names and residue numbers. C) Top view of the interface showing the interaction of the C-terminus in the interface. Residues that are identical in eukaryotic sequences are labeled in red text. All 3D structure figures were generated in PyMOL (www.pymol.org) (75). 38 Figure 3.3: The AfNep1 Dimer A) The symmetric AfNep1 dimer generated by symmetry operation of the asymmetric unit. Bound SAH molecules (not added to crystallization conditions) shown in yellow sticks. B) Close-up of dimerization interface. Cysteine involved in dissulfide bond labelled with an asterisk. C) Top view of the dimerization interface. 39 Figure 3.4: Conservation Maps on the Surface of ScNep1 Residues are colored based on conservation in alignments with human and fruit fly Nep1 sequences using ClustalW (red = identical; green = highly conserved; blue = weakly conserved). The ?front? view in A. is rotated 90? in either direction around the x-axis to produce the orientations in B. In C, ?inside? refers to the dimerization interface, and ?outside? is produced by a 180? rotation around the y-axis. 40 Figure 3.5: Conservation Maps on the Surface of AfNep1 A-C) Residues are colored based on conservation in alignments with eukaryotic Nep1 sequences using ClustalW (red = identical; green = highly conserved; blue = weakly conserved) (46). 41 Chapter 4: Nep1 in Complex with RNA 4.1: Overview Our structures of ScNep1 are the first examples of a SPOUT methyltransferase bound to its target RNA. Even more interesting is Nep1 is the only member of the SPOUT family known to date that methylates pseudouridine (22). Presented in this chapter is a base-by-base description of the interactions between ScNep1 and the 14 base RNA fragment. Conformational changes between free ScNep1 versus ScNep1 bound to RNA and comparisons between ScNep1 with one molecule of RNA versus ScNep1 with two molecules of RNA are also presented in dedicated sections within this chapter. 4.2: Interaction of ScNep1 with RNA Using the binding sequences for ScNep1 obtained through RNA three-hybrid screening by Buchhaupt et al. (21), a series of oligonucleotides were designed and tested for binding to ScNep1 in stoichiometric complexes. It was determined that an ideal complex was obtained when ScNep1 and a 14 base RNA fragment with the sequence G1G2G3C4U5U6C7A8A9C10G11C12C13C14 were mixed in a 2:2 (monomer:RNA) ratio. Crystals were obtained for Nep1 bound to this 14 base RNA fragment in two crystal forms (Fig. 2.1). In one crystal form, a dimer of ScNep1 is bound to one RNA molecule (Fig. 2.1A). In the second crystal form, the dimer binds two separate RNA molecules (Fig. 2.1B). In both cases, the RNA forms a stem loop structure in which eight bases forming canonical base pairs in the stem and six bases 42 in the loop (UUCAAC) have base specific contacts with the protein. Electron density was not observed for residues 1-27 and 56-64 of both protein monomers in the RNA complexes, and residues 76-82 of monomer A of the two-RNA complex. There are no significant differences between the RNA molecules bound in the two complexes. In the comparison of the sequences of RNA found to bind Nep1 by RNA three-hybrid screens, all the sequences contained the consensus sequence C/UUCAAC, and the most tightly binding were those sequences that contained the consensus sequence within the loop of a predicted stem-loop (21). These six bases form the basis for specific recognition by the protein. In the crystal containing the one RNA per dimer complex, the empty RNA-binding site is involved in four hydrogen bonds with the RNA backbone from a symmetry-related complex. In both complexes, the RNA binds on highly positively charged surfaces of the dimer as predicted in previous reports (16, 17). However, the protein contains significant positively charged surface that is not contacted by RNA in these structures (Fig. 4.2). This positively charged surface is conserved in AfNep1 as well. This is consistent with the idea that Nep1 binds to RNA on the pre-ribosome, which makes it likely that it will contact significantly more RNA than is observed with these minimal constructs. The regions that are not contacted by RNA in these structures contain many conserved residues as well, indicating that this surface is important for Nep1 function (Fig. 3.3). ScNep1 provides several key interactions that confer the observed specificity for a UUCAAC sequence (Fig. 4.3, schematic in Fig. 4.4). The first base of the consensus sequence, U5, is contacted through base-specific contacts through bidentate 43 hydrogen bond interactions between the NH1 and NH2 of conserved Arg 136 of monomer A (A:Arg 136 NH1 and NH2) and the C2 carbonyl oxygen (O2) of the uridine base (Fig. 4.3A, 4.4). There are no contacts between the protein and C4 carbonyl oxygen (O4), which explains the observation that a cytosine, which differs from uracil solely by an amine group at C4, can substitute at this position (21). The next uracil in the consensus sequence, U6, is the site of modification by the enzyme. It is flipped out from the loop and bound in a pocket between the two monomers, near the bound SAH molecule, in the enzyme active site (Fig. 4.3A, B). It is recognized by a direct hydrogen bond with A:Arg 132 NE (N? of arginine 132 of monomer A) at the U6 O2 (C2 carbonyl oxygen of U6), and water-mediated hydrogen bond interactions between the U6 O4 and the backbone amide of B: Ser 233 and the U6 O4 and N3 and OE (side chain carbonyl oxygen) of conserved B: Gln 94. Arg 132 is held firmly in position by hydrogen bonding between Arg 132 NH1 and NH2 and conserved A: Asp 101 OD1 and OD2. In addition, hydrogen bonds are observed between the U6 O2 and NH1 and NH2 of B:Arg 88 and between the ribose O4 and the NH1 of B:Arg 88. This would hold U6 firmly in position, and may serve as a means to stabilize negative charge on the base during methyl transfer. Arg 88, in turn, is held in place by interactions between its NH2 and NE and the OD1 and OD2 of conserved A:Asp 90. Asp 90 is the equivalent of the human Asp 96 that is mutated to glycine in Bowen-Conradi syndrome. That mutation would have the effect of destabilizing the interaction with the U6 substrate both through disruption of the interaction with the Arg 88 and hydrophobic contacts with Ile 91 through repositioning. Finally C5 of the uridine base is positioned a mere 3.9 ? away from the sulfur atom of SAH, in perfect 44 position for a methyl transfer to the C5 position of the uracil (Fig. 4.3B). If a pseudouridine were in its place, then N1, which is an equivalent position to C5, would be the target of methylation. In the remaining consensus sequence, C7 is recognized through backbone interactions between the carbonyl oxygen, O2, and the backbone amides on either side of A: Arg 132, and a base stacking interaction with A: Arg 136. A8 is sandwiched between a stacking interaction with conserved A: Arg 129 and van der Waal?s contact with conserved A: Ile 159, and donates a hydrogen bond from N6 to the backbone carbonyl oxygen of A: Thr 127. A8 N6 is also involved in a water- mediated hydrogen bond interaction with the side chain OH of Thr 127 and A8 N1 is in hydrogen bonding distance of the backbone amide of A: Arg 129. A9 is recognized through hydrogen bonds between the backbone carbonyl of A: Arg 129 and the N6, a van der Waal?s interaction between the side chain of A: Val 128 and the C2, and a stacking hydrophobic interaction with conserved A: Leu 140. Although Val 128 is not highly conserved, it is Val, Thr or Cys in other sequences, which would be able to maintain the interaction with A9 C2. Finally C10 is specified by two hydrogen bonds with conserved A: Arg 136, one between the Arg NH2 and the C10 N3, and the other between the Arg NE and C10 O2. There is also an interaction between the C10 ribose O2? and the backbone carbonyl of A: Arg136. Arg 136 contacts C10 via three hydrogen bonds, U5 via two hydrogen bonds, and stacks in between A9 and C7. Consequently, Arg 136 is invariant in all sequences. In a functional analysis by Taylor et al., mutations of Arg 136, as well as Arg 88, Arg 129, and Arg 132 to Ala 45 all result in diminished RNA binding activity as measured by yeast three hybrid assays with consensus sequences (16). Of the eight bases that form the stem in canonical Watson-Crick base pairs, only one, G11, is contacted at the base by a protein residue. B: Glu 74 accepts a hydrogen bond from G11 N2, and contacts the G11 ribose O2? via a water-mediated interaction (Fig. 4.5). Other backbone interactions between the protein and the RNA include contacts between A: Gln 143 and C10 O2? and phosphate between C10 and A9, A: Arg 129 and C11 O2P, A: Arg 211 and the phosphate between U5 and U6. Of these, Arg 129 and Gln 143 are highly conserved, while Arg 211 is only weakly conserved (Fig. 3.1A). Based on this data, we can conclude that very little specificity is determined in the stem portion of the RNA. 4.3: Catalytic Residues in the Active Site The methylation reaction that Nep1 catalyzes would require the deprotonation of N1 of the pseudouridine so that it could attack the methyl group from SAM. Stabilization of a negative charge on the uridine ring would result in a decrease of the pKa, which was measured at 8.97 for free pseudouridine (76). The likely candidate for this stabilization would be invariant Arg 88, for which both primary amine nitrogens are within hydrogen bonding distance of O2, the C2 carbonyl oxygen (3.3 and 2.8 ?); (Fig. 4.3B), and which is stabilized by bidentate hydrogen bond interactions with invariant Asp 90, which is mutated in Bowen-Conradi Syndrome (77). Mutation of Arg 88 to Asp resulted in a significant loss of Nep1 RNA binding activity (17). Arg 88 and Asp 90 are located at the beginning of ?A, and are surrounded by Pro 89 and Ile 91, which are also invariant. Ile 91 is in van der Waals contact distance of the 46 uridine ring in the active site. This region is labeled as the ?Catalytic Loop? in Fig. 3.1A and 3.1B. Arg 132 may also provide further stabilization, since it is also within hydrogen bonding distance (3.0 ?) of O2. These interactions would increase deprotonation at the N1 through localization of the negative charge to O2 where it would be stabilized by interaction or proton donation by Arg 88. This would in turn enable N1 to accept the methyl group from SAM. 4.4: Free Nep1 vs Nep1 Bound to RNA When a monomer of free Nep1 is compared to monomers from the RNA Nep1 complex, an RMSD of 0.76 ? is observed for 415 out of 426 residues. Most of the interaction between the RNA and the protein are mediated by the helix ?c? and adjacent loops on the monomer that does not provide the SAM to the active site. However, the most significant conformational changes are seen in the small Nep1 specific inserted domain on the SAM binding side of the dimer interface (Fig. 4.5A, B). This domain, consisting of a 3-10 helix (Hb?), small ?-helix (?a?), and a ?- hairpin, undergoes a shift in positioning towards the dimer interface when RNA is bound. In addition, there is alteration of overall secondary structure, as the structure changes to accommodate interactions with the RNA (Fig 4.5A, B). The domain, which is packed against ?4, houses Arg 88, which may be important for catalysis, and Glu 74 that interacts with G11. The buried surface area in the RNA bound structure (1338 ?2) is slightly decreased, due to an opening of the interface to accommodate the RNA, which buries an additional 721 ?2 on the monomer containing the methyl donor and 351 ?2 on the other monomer. 47 4.5: Two RNA vs One RNA Both the 1:1 and 1:2 (ScNep1 dimer:RNA) complexes for the complexes between Nep1 and RNA were obtained using the same preparation of Nep1 to RNA in a 2:2 (ScNep1 monomer:RNA) ratio. The isolation of these two complexes in the crystal structures suggests that both complexes exist in solution, which implies differences in the binding affinities between the two sites. Comparisons of the structures of these complexes reveal differences in the structures of the RNA binding sites as a function of occupancy with RNA (Fig. 4.5C, D). The RMSD for the alignment of the protein molecules in the complexes averages 0.85 ? for 421 residues of the protein chains, with the largest difference being between chain A of the 1:1 complex and chain B of the 2:1 complex. This number is larger than the RMSD for the comparison between the Nep1/1RNA complex and the free enzyme, indicating a larger conformational difference between the Nep1/1RNA and Nep1/2RNA complexes than between the free enzyme and the Nep1/1RNA complex. Upon closer inspection, we observed that the positioning of one monomer relative to the other is slightly different between the two complexes. In the Nep1/2RNA complex, the two monomers are slightly closer together near the RNA binding site, indicating that RNA binding pulls the two monomers closer together. As a result, the SAH molecule is pulled closer to the RNA. The SAH molecule is contacted mostly by backbone interactions, but one residue, Asp 214, hydrogen bonds the ribosyl 3?OH via one of its side-chain carbonyl oxygens (Fig. 4.6). In the Nep1/2RNA complex, for one monomer site the Asp 214 side-chain is rotated away from the SAM binding site and no longer contacts the bound SAM (Fig. 4.7A, B). This results in a slightly altered 48 positioning of the methionyl moiety in the active site, with the sulfur atom positioned farther away from the uridine base (Fig. 4.7A, B). In addition, the SAM binding pocket on the side of the dimer not bound to RNA in the Nep1/1RNA complex is not visibly occupied by SAH. Based on these observations, and the presence of additional conserved positive charged surface (Fig. 4.2), it is possible that both RNA binding sites would be occupied on rRNA, perhaps with varied binding affinity or specificity, and additional binding surface would be used to interact on the surface of the rRNA. 4.6: Binding Assays with Nep1 and RNA Many attempts were made to determine the binding affinity of Nep1 to the target sequence. Electrophoretic mobility gel shift assay (EMSA) and anisotropy are described. For gel shift, a 12 base RNA oligonucleotide (5'-CUCUUCAACGAG-3') was radioactively 5' end-labelled with T4 polynucleotide kinase and [?-32P]ATP using the KinaseMaxTM kit from Ambion. The labelling reaction was performed with 25 pmol of RNA and 33 pmol of radioactive ATP, 1 ?L of T4 polynucleotide kinase, 2 ?L of the provided 10 X kinase buffer, 13 ?L RNAse free water and incubated at 37 ?C for one hour. After the incubation period, 1 mM EDTA was added and the reaction was halted via heating at 95 ?C for two minutes. Labelled RNA was added to a reaction mixture of ScNep1 in varying concentrations from 1.2 x 10-4 M to 2.4 x 10-7 M in buffer containing 3 % glycerol, 75 mM NaCl, 2 mM MgCl2, 50 mM Tris pH 7.5, and 0.1 % BME and allowed to incubate at 25 ?C for one hour. After incubation, the reaction was stopped via heat denaturation and ran on a 6 % TBE polyacrylamide gel. All gels were run in triplicates, gel dried, exposed to a phosphor screen and scanned with a StormTM Imager. Quantification of bands was performed with 49 ImagequantTM. Figure 4.8A presents an example gel with ScNep1 concentrations showing weak binding, which conflicts with reports from M. jannaschii Nep1 fluorescence experiments performed by Wurm et al. (22). Upon performing the same experiment with a control RNA oligonucleotide with sequence 5'- UGUGUGUGUGUG-3' and obtaining nearly identical results, it appeared Nep1 had the ability to non-specifically bind short RNA oligonucleotides. Fluorescence anisotropy measurements were made on the Fluorolog-3 Jobin Yvon Horiba fluorescent spectrophotometer set at 4 nm bandwidth and 1 second integration times. 5?-Fluorescein (Fl) labeled RNA (5'-Fl-CUCUUCAACGAG-3') and 3'-Fl labeled control RNA (5'--UGUGUGUGUGUG-Fl-3') was used for the binding assay. 10 nM labeled RNA was added to 1 mL of solution containing ScNep1, 75 mM NaCl, 50 mM Tris pH 7.5, 2 mM MgCl2, and 0.2 % BME. A titration curve was obtained by removing 200 ?L of solution and replacing 200 ?L with dilution buffer containing 75 mM NaCl, 50 mM Tris pH 7.5, 2 mM MgCl2, and 0.2 % BME, and 10 nM labeled RNA. The dilution buffer contained 10 nM labeled RNA to keep oligonucleotide concentration consistent throughout the entire titration curve. The reaction was done at temperature controlled 25 ?C with constant stirring. Measurements were taken 8 minutes after every dilution to allow temperature and concentration equilibration with excitation at 495 nm and emission spectra averaged from 518-522 nm. The results of anisotropy reflected those seen in gel shift. Nep1 bound both the control RNA and target RNA with nearly identical affinity. Figure 4.8B shows an example of raw anisotropy data from control RNA. Anisotropy data could not be fit to an acceptable model. Both experiments described here suggest 50 Nep1 non-specifically binds to short RNA oligonuclotides. While, this is not ideal for measuring specific affinities of the target sequence, it does provide support that Nep1's additional positive surface can bind additional RNA (Fig. 4.2), which is expected given Nep1's environment around the 18S rRNA. 4.7: Chapter Summary The detailed structural description of the active site and all interactions associated with the binding of the target RNA presented in the two bound structures results in an understanding of Nep1?s ability to methylate pseudouridine. Through structural analysis, it is possible to present mechanistic insights including the role of a mutation causing Bowen-Conradi disease. 51 Figure 4.1: Overall Structure of the Nep1/RNA Complexes Structures of the A) one RNA and B) two RNA complexes with SAH (pink) and RNA (magenta and yellow) shown in sticks. 52 Figure 4.2: Electrostatic Surface of Nep1/1RNA Complex Electrostatic surface of the Nep1 complex showing front, top and bottom views. The surface is colored red to blue, negative to positive charge (calculated using vacuum electrostatics in PyMOL). 53 Figure 4.3: Nep1-RNA Interactions A) Stereo figures of RNA bound in the cleft between the two monomers. Residues are colored according to monomer from which they originate (green-monomer A; cyan- monomer B). Dash lines represent connections between atoms within hydrogen bonding distance. RNA (magenta) truncated to show only the bases that contact the protein for simplicity. B) Stereo view of a close-up of the uridine base interactions with 2Fo?Fc map contoured at 1?. Protein residues are colored as in (A). U6 is shown in magenta. Distances between atoms are indicated by red text. 54 Figure 4.4: Schematic of Interactions Between Nep1 and RNA Shown here is a diagram of the interactions between the crystallized RNA stem- loop and Nep1 residues (colored text; blue = monomer B; green= monomer A). Red arrows indicate possible hydrogen bonds and black lines terminated by black dots indicate hydrophobic contact. Blue dots indicate water molecules. 55 Figure 4.5: Conformation Changes Upon RNA Binding A) Front and B) Bottom views of the Nep1/1 RNA complex (green-monomer A, cyan-monomer B) superposed on the free dimer (magenta). C) Front view of the alignment of the 1 RNA and 2 RNA (yellow) complexes. D) Bottom view of both RNA complexes and the free dimer. Monomer B, shown here on the right, was used to align the monomers. Asterisk indicates the Nep1 specific domain where significant structural change occurs, and ?c' indicates the RNA-binding helix that shifts relative to the aligned monomer. 56 Figure 4.6: Interactions with SAH Shown here are the interactions holding SAH (yellow) in the active site with 2Fo?Fc map contoured at 1?. Protein residues are colored as in cyan and U6 is shown in magenta. Distances between atoms are indicated by black text. 57 Figure 4.7: Alignment of SAM Binding Pockets A) Alignment of the SAM-binding pockets of Nep1/1RNA complex monomer B (cyan), Nep1/2RNA monomer B (green), Nep1/2RNA monomer A (magenta) and free Nep1 dimer (orange) showing side chains of residues that show the most conformational differences between the complexes. Spheres indicate bound waters, and are labeled ?W?. B) Stereo view of the alignment of SAM binding sites in the Nep1/1 RNA and 2 RNA complexes. Nep1/1RNA monomer B and Nep1/2RNA monomers A and B are colored as in (A). 58 Figure 4.8: Nep1 and RNA Binding Assays A) StormTM scan image from phosphor screen of ScNep1 and target RNA (5'- CUCUUCAACGAG-3') electrophoretic mobility gel shift assay. B) Anisotropy data taken from Fluorolog-3 Jobin Yvon Horiba fluorescent spectrophotometer of ScNep1 binding to control RNA (5'-UGUGUGUGUGUG-Fl-3'). 59 Chapter 5: Nep1 Discussion and Conclusions 5.1: Overview The four deposited structures provided a leap forward in understanding the role of Nep1 in ribosome biogenesis, specifically contributing to a molecular view of the interaction with target RNA. This work supports previous research suggesting Nep1 as a pseudouridine methyltransferase along with a second function that is essential for cell survival (21, 22). This chapter provides discussion and conclusions based on the information gained from the analysis of AfNep1, ScNep1 free, and ScNep1 bound to one and two molecules of target RNA. 5.2: Nep1 Project Discussion and Conclusions Prior to this work, the structures of the ScNep1 in monomeric form, and the archaeal M. jannaschii Nep1 dimer, allowed for the classification of Nep1 as a SPOUT methyltransferase, characterization of the Nep1 specific structural elements and analysis of the SAM binding pocket, and, in combination with mutagenesis and NMR mapping on the M. jannaschii Nep1, allowed for a prediction of the RNA binding site (16, 17, 22, 78). This report provides a view of dimeric ScNep1 in apo form and in complex with RNA, allowing a first look at a SPOUT methyltransferase bound to RNA, a high resolution analysis of the interactions of the only known N1- specific pseudouridine methyltransferase with a substrate analog, and a picture of the RNA structure that Nep1 stabilizes upon binding. These data confirm that the RNA binds to the cleft between the two monomers, which agrees with previously reported NMR mapping data and mutational analysis that reported mutation of the M. 60 jannaschii equivalents of R88, R129, R132 and R136 significantly reduced RNA binding activity (16, 22). In Nep1, the binding site accommodates a stem-loop structure, with the majority of binding observed between the RNA loop and the protein. Both the RNA three-hybrid screening of ScNep1 and the specificity determination on M. jannaschii Nep1 indicate that Nep1 binds more tightly to a predicted stem-loop structure, suggesting that this is the preferred substrate for Nep1 (21). However, for M. jannaschii Nep1, sequences that do not contain a stem can also bind (21, 22). We have no information about the structure of the free RNA, and no similar loops containing this sequence were found in the structure databases, but the stem-loop is predicted to form at 37 ?C by RNAfold (79), although we are not sure if U6 would be flipped out in the absence of protein. The RNA seen in our structures fails to contact much of the positively charged surface of the protein, leaving open the possibility that the actual binding site is larger than the selected consensus sequence. It has been proposed that Nep1 is the first example of an N1-specific pseudouridine methyltransferase and that it catalyzes the only such modification known in 18S rRNA, the N1-methylation on the hypermodified m1acp3? at position 1191 in yeast (22). The structural evidence, seen in the specific positioning of the C5 of uridine in the enzyme active site, supports this. The uridine is held in place by several contacts that impart this specificity, and the interaction with Arg 88 suggests that this residue plays a role in catalysis, by promoting deprotonation of the ring (Fig. 5.1). Asp 90, which is equivalent to Asp 86 that is mutated in Bowen-Conradi Syndrome, also plays a role by holding Arg 88 in place in the structure. There are no candidates for direct hydrogen binding near the C5 of the uridine in our structure that 61 may allow specific selection of pseudouridine, but the absence of an activating nucleophile in the vicinity of C6 in these structures rules out the possibility that this protein functions as an m5U methyltransferase, as would be suggested if uridine was the substrate (80, 81). In addition, m5U modification has not been observed in yeast 18S rRNA. However, this means that uridine is indistinguishable from pseudouridine in the Nep1 active site, but it will only catalyze methyl transfer on pseudouridine. This is supported by fluorescence binding data for M. jannaschii Nep1 reported by Wurm et al. that shows no significant differences in RNA binding Kd when pseudouridine is replaced with uridine in high affinity RNA binding sites (22). Although it is likely that the N1-methyltransferase activity is important for Nep1 function, as supported by the severe defect caused by mutations at Asp 90, it has been reported that mutations in the SAM binding site of Nep1 that reduces its ligand binding activity by 100 fold results in no disruption of Nep1 activity (17). Therefore it is plausible that binding of Nep1 to the rRNA in the absence of methyl transfer activity is sufficient to carry out at least some of its role in ribosome biogenesis. In addition, overexpression of yeast ribosomal protein Rps19 compensates for deficiency in Nep1 function (21). An explanation put forth for these observations was that Nep1 might play a role that is independent of Nep1 methyltransferase activity (17, 21). A possible role may be to act as a chaperone of RNA folding during a specific assembly step. The observation of a two RNA complex in which both binding sites are occupied suggests that it is plausible that rRNA occupies both sites when Nep 1 binds in vivo. Based on structural analysis, the two binding sites are non-equivalent, highlighted by differences in binding to SAH 62 (Fig. 4.7A, B). This idea that both sites would be occupied in the ribosome is further supported by the presence of extensive positively charged surface not contacted by the stem-loops in the structures. Two of the three possible Nep1 binding sites in yeast 18S rRNA are located within 50 ? of each other in the 3-dimensional structure of the eukaryotic ribosome (Fig. 5.2A) (30) (82). Those sites are the loop terminating helix 31 and a portion of helix 42 (Fig. 1.2 p12). Based on the EM reconstruction of the eukaryotic ribosome (30) and the recent crystal structure of the complete yeast ribosome (82), in order for Nep1 to bind the site at helix 42, significant secondary structure alteration would have to occur (Fig. 5.2B; Fig 1.2 p12; Fig. 5.3). Half of the expected stem loop that is expected to form when Nep1 binds is part of helix 42. Therefore, it is possible that when Nep1 binds, there is a rearrangement of the interactions to produce the expected stem-loop (Fig. 5.3C). This rearrangement would also result in the two Nep1 sites being moved closer together, since the distance between active sites in the Nep1 dimer is ~20 ?, while it is shown to be 50 ? apart in the mature rRNA (30). The loop at the end of helix 42 interacts with the loop of helix 41 (Fig. 5.2B). Helix 41 is the site of most of the contacts between the rRNA and S19 (Fig. 5.2B & Fig 1.2 p12). Therefore, a possible model is that Nep1 interacts with the rRNA at two sites ? the loop of helix 31, where it catalyzes the methylation of ?1191 and the stem of helix 42, where it acts as an RNA chaperone and breaks apart helix 42, rearranging the area and the interactions with helix 41 to promote S19 binding. In this scenario, more of the Nep1 surface would be expected to contact rRNA, as predicted by this structure. This is assuming that rRNA in early maturation stages have structures near the predicted structure of the mature rRNA. However, there is 63 evidence that pre-rRNA structure can differ from mature rRNA structure (83). In the absence of snR57, which is predicted to bind between bases 1568 and 1584, a Nep1 deletion is no longer lethal, indicating that Nep1 is only required when snR57 is present (21). Since snR57 guides the methylation of G1572 (29), a site near helix 42 (Fig. 1.2 p12), this interaction between Nep1 and snR57 may be due to structural changes that occur as a result of methylation. Another possibility, however, is that the interaction between Nep1 and its binding site in that region may promote the dissociation of snR57 from rRNA. This dissociation may be necessary for proper rRNA structure, or even for N7 methylation of G1575 catalyzed by Bud23p (84). In any case, the genetic interaction between the snR57 and Nep1 supports the possibility of Nep1 binding in the region near helix 42. M. jannaschii Nep1 binds to yeast 18S rRNA sequences corresponding to the loop of helix 31 (78), and AfNep1 similarly is capable of binding yeast rRNA sequences corresponding to the Nep1 binding site near helix 42 (data not shown). Residues involved in key interactions with the RNA, for example Arg 88, Arg 136, Arg 129, Leu 140 and Leu 159, are identical in AfNep1. Superposition of the archaeal dimer on the yeast structures showed that the RNA interaction mode is likely to be very highly conserved between archaea and yeast (Fig. 5.4). Although these structures do not provide all the answers on Nep1 function, it does provide the basis for continued studies on the ancient structural mechanisms utilized by Nep1 in rRNA processing. 64 5.3: Hypothesis of Nep1's Role in Ribosome Biogenesis 5.4: Future Directions In order to further understand the role of Nep1 in ribosome biogenesis, questions persist surrounding the proposed chaperone function. The ScNep1/1RNA and ScNep1/2RNA complex structures shows that Nep1 can bind RNA simultaneously on both sides of the dimer, but its still unknown if this does indeed happen simultaneously. Does Nep1 preferentially bind one target before the other or do both sites bind at the same time? Next, Does Nep1 direct a major conformational shift in helix 42 that results in the dissociation of snr57 and the subsequent loading of S19? To look at these questions, I believe work must shift from short RNA oligonucleotides to longer sections of the 18S rRNA that encompasses both Nep1 65 target sites and the location of S19 binding. Utilizing a longer section of the 18S rRNA in different binding assays that includes structural biology from both x-ray crystallography and NMR, RNAse secondary structure foot printing and fluorescence binding assays, could provide needed answers to these important questions. Finally, small angle x-ray scattering (SAXS) is currently an emerging field and may provide an ideal technique in the near future for visualizing the proposed rearrangements of the 18S rRNA. 66 Figure 5.1: Proposed Nep1 N1-pseudouridine Methyltransferase Mechanism Putative mechanism for the transfer of a methyl group to the N1-position of pseudouridine (?) by Nep1. 67 Figure 5.2: Possible Role for Nep1 in S19 Loading A) and B) Three dimensional structure of the head region of eukaryotic 18S rRNA, truncated to remove bases 1212?1439 for simplicity. (A) and (B) were generated from PDB ID 3JYV [cryo-EM model of the 40S subunit of the Thermomyces lanuginosus ribosome (30)]. 68 Figure 5.3: RNA Structures in Nep1 Binding Sites A) Structure of RNA in Nep1 complex with schematic of the secondary structure shown on the right. B) Structure of Nep1 site at helix 31 and C) helix 42 (from the cryo-EM reconstruction of the eukaryotic ribosome (30)). Secondary structure predicted to be induced upon Nep1 binding shown on the right. For all figures, the consensus binding site is colored yellow and the * indicates the uracil inserted in the Nep1 active site. 69 Figure 5.4: AfNep1 and ScNep1 Aligned with Sequence Conservation Stereo view of ScNep1 (cyan) bound to one RNA molecule aligned to AfNep1 (with symmetry mate to form dimer), colored to show conservation of the RNA binding site between archaeal and eukaryotic Nep1 proteins (red = identical; green = highly conserved; blue = weakly conserved). Sequence alignment done using ClustalW (46). 70 Figure 5.5: Proposed Chaperone Role of Nep1 in Ribosome Biogenesis Model of the proposed role of Nep1 in ribosome biogenesis highlighted by Nep1's ability to bind both target sequences simultaneously and chaperone the folding of helix 42, which will allow the loading of S19 to helix 41. 71 Chapter 6: Purification of Rio3 6.1: Overview Rio3 is an atypical kinase with an unknown function and unknown structure. By sequence analysis with other family members, Rio3, known as RioK3 in humans, possesses a conserved RIO domain and an extended N-terminal domain of approximately 200 amino acids (35, 37, 39, 48). The goal of this project was to solve the crystal structure of the RIO domain for RioK3. The full-length construct (519 residues) suffered from insolubility problems when expressed in E. coli. Therefore, the focus of research was spent trying to express, purify, and crystallize a shortened construct containing only the RIO domain of RioK3 (residues 193-519). Presented in this chapter is the best purification protocol for the RIO domain of RioK3 and for the N-terminus of RioK3. Initial crystallization trials from the resulting protein acquired from this purification protocol are also described in this chapter. 6.2: Purification of RioK3 The human shortened RioK3 gene (residues 193-519) with an N-terminal TEV cleavage site was PCR amplified using sequence specific primers (5'- GAGAACCTGTACTTCCAGGGAATGGATTTAAAACTATCAAAC-3' and 5'- GGGGACCACTTTGTACAAGAAAGCTGGGTTATCATTCATCATATAGTAGT GGTGGGTC-3') and recombined into pDONR201 utilizing Invitrogen?s GatewayTM technology. The resulting plasmid was sequenced and confirmed the correct sequence for the shortened RioK3 construct with N-terminal TEV cleavage site. The plasmid was recombined into pDest527 designed with an N-terminal 6X-His tag (plasmids 72 obtained from Protein Expression Laboratory, SAIC, Frederick, MD). RioK3-Short construct was transformed into E. coli Rosetta BL21-DE3-pLysS cells via heat shock. The construct was expressed in 1 L LB broth containing ampicillin (100 ?g/ml) and chloramphenicol (34 ?g/ml) at 37 ?C until the optical density of the cells reached 0.5 (approximately 4 h). At the correct cell density, the cells were transferred to 20 ?C incubator/shaker and allowed to cool before induction. The RioK3-Short construct was induced with 1 mM isopropyl thio-?-D-galactoside (IPTG) and grown overnight (16 h) at 20 ?C, shaking at 250 rpm. Cells were harvested by centrifugation and cell pellet was frozen overnight at -80 ?C. The frozen pellet was resuspended in 50 mL lysis buffer containing 500 mM NaCl, 50 mM Tris pH 8.0, 0.2% ?-mercaptoethanol (BME), 2 mM MgCl2, 1 mM adenosine triphosphate (ATP), 0.1 mg/ml DNase1 (Roche), 10 ?g/ml RNaseA (Roche), and 0.1 X Bugbuster (Novagen) per liter of culture. The resuspended pellet lysed on ice after stirring for 1 h. The lysate was transferred to centrifuge tubes and the cell debris was spun down at 17000 rpm in Beckman preparative (rotor: Beckman 45 Ti) ultracentrifuge for 40 minutes at 4 ?C. The supernatant was collected through a 0.22 ?m filter. The supernatant containing the soluble RioK3-Short construct was loaded onto an equilibrated 5 mL GE Healthcare HisTrap HP column using the BioRad Duoflow FPLC. The column was equilibrated with buffer A containing 500 mM NaCl, 50 mM Tris pH 8.0, 0.2% BME, 2 mM MgCl2, 1 mM ATP. The bound RioK3-Short construct was washed extensively with buffer A (>10 column volumes) followed by another extensive wash including 100 mM imidazole. RioK3-Short was eluted from the column using a linear gradient from 100 mM imidazole to 1 M imidazole over ten column volumes. 73 The fractions containing RioK3-Short were combined and set up in dialysis membrane with buffer A and 2 mg TEV protease. The cleaved protein was loaded back onto the 5 mL HisTrap column. The RioK3-Short protein did not come through in the flowthrough: 20 mM imidazole in buffer A was needed to elute the bound protein. The fractions containing RioK3-Short protein were pooled together and concentrated to a final volume of 3 mL. A Superdex 200 16/60 size exclusion column was equilibrated with buffer A and the Rio3-short protein was passed over the column as a final purification step. The fractions containing purified RioK3-Short protein were combined and concentrated to 15 mg/ml for crystallization. All steps of purification were analyzed for purity by SDS-PAGE (Fig. 6.1). Table 6.1 shows the RioK3 constructs that were expressed for purification and crystallization trials. 6.3: Initial Crystallography of RioK3-Short The purified RioK3-Short construct was concentrated to 15 mg/mL and immediately screened for crystals using the Art Robbins PhoenixTM liquid handler. The sitting drop 96 well 3-well Intelliplate was chosen for screening with conditions known to promote crystallization. These varying conditions are combined into the 96 deep-well format and were purchased from Qiagen (PEGTM Suite, CryosTM Suite), Emerald Biosystems (Wizard 1TM, Wizard 2TM, Wizard 3TM, NatrixTM) and Hampton Research (IndexTM). The 3-well format of the intelliplate allowed for a concentration range for each condition by setting up the sitting drops in 1:2, 1:1 and 2:1 protein:well solution ratios. In total, the initial crystallization screen for RioK3-Short consisted of 1,152 different drops. Of those 1,152, one condition yielded extremely small (~5x30?m) protein crystals shown in Figure 6.2A. The crystals grew in 25% PEG- 74 3350, 0.1 M HEPES pH 7.5, 0.2 M MgCl2. Those crystals were reproducible from protein expressed in minimal media (described in chapter 7) via the hanging drop method (Fig 6.2B) and sent to the NECAT APS beamline at Argonne National labs for testing of diffraction. This work was promising, yielding diffraction to ~5.5 ? shown in Figure 6.3. 6.4: Expression and Purification of the N-Terminus of RioK3 While working on the purification and crystallization of the RioK3 RIO domain, the N-terminus of RioK3 was shown to bind to polyubiquitin (85). N- terminal fragment 16-153 was shown to bind to Lys-48 linked and Lys-63 linked Ubuitin (85). RioK3 does not possess a known Ubiquitin binding domain, which suggests RioK3 could bind Ubiquitin via a novel mechanism. Using Invitrogen GatewayTM cloning, RioK3-1-153 and RioK3-1-193 were cloned with an N-terminal histidine tag followed by a TEV cleavage site with forward sequence (5'-GAGAAC CTGTACTTCCAGGGCATGGATCTGGTAGGAGTGGCATCG-3' and reverse primers 5'-GGGGACCACTTTGTACAAGAAAGCTGGGTTATCACTTTTTAGG AGTGGGAACCGGTTT-3' and 5'-GGGGACCACTTTGTACAAGAAAGCTGGG TTATCAAATTCCATCTCCTACCTGAAACTC-3', respectively. After unsuccessful purification attempts due to insolubility, RioK3-1-193 was recombined into pDest566 with maltose binding protein (MBP) as an N-terminal fusion protein resulting in HIS6-MBP-TEV-(RioK3-1-193). The fusion protein was used to improve solubility. The expression and purification scheme was similar to RioK3-Short with two modifications. First, the lysis and size exclusion buffers contained 200 mM NaCl, 50 mM Tris pH 8.0 and 0.2 % BME. The elution buffer contained 200 mM NaCl, 50 75 mM Tris pH 8.0, 0.2% BME and 1 M imidazole. Second, after TEV cleavage MBP- RioK3-1-193 did not require 20 mM imidazole for elution and came out as expected in the flowthrough. All steps of purification were analyzed by SDS-PAGE and shown in Figure 6.4. The successful purification of the N-terminus of RioK3 provides a great starting point for the next researcher to look at interactions with Ubiquitin. 6.5: Chapter Summary The initial crystallography results showing diffraction from crystals acquired from the purification protocol presented in this chapter was very promising. The crystals were reproducible and they diffracted! This beginning success provided a great starting point for crystal optimization. 76 Figure 6.1: RioK3-Short Purification Gels A) SDS-PAGE of RioK3-short after first pass over GE Healthcare HP affinity column (IN=induced sample, Sol=supernatant after lysis, FT=flowthrough, TEV Trial=20uL elutant incubated with 20 ?L TEV protease (1 mg/mL) for 30 minutes on rocker and ran as lane 8, Tag=histidine tag cleaved during TEV trial). B) RioK3-Short after cleavage passed over affinity column to remove tag and uncleaved protein (B=before cleavage, A=after cleavage). Concentrations indicate amount of imidazole required for elution from column. C) Elution from GE Healthcare Superdex 200 16/60. Expected MW of Riok3-Short is 37,717 Da. 77 Table 6.1: RioK3 Expressed Constructs 78 Figure 6.2: RioK3-Short Crystals A) First Riok3-Short crystals obtained from G12 of the IndexTM screen in the 2:1 protein:screen ratio. B) Reproduced crystals from RioK3-Short grown in minimal media. Diffraction of crystals from (B) screened at the synchrotron resulted in diffraction to ~5.5 ?. 79 Figure 6.3: Diffraction of RioK3-Short Diffraction image of a RioK3-Short crystal grown in minimal media, collected at NECAT APS beamine in Argonne, Illinois. Red circle represents 5.5 ? resolution. 80 Figure 6.4: MBP-RioK3-1-193 Purification 81 Chapter 7: Optimization of RioK3 7.1 Overview The crystals acquired from the RioK3-Short construct and purification protocol presented in chapter 6 provided the first look at diffraction. The 5.5 ? resolution diffraction however, does not provide enough resolution to acquire a solution, either by de novo phasing or by molecular replacement. This chapter will describe the many methods used to optimize RioK3 in order to acquire strong enough diffraction for structure solution. 7.2: General Optimization Methods 7.2A: Precipitant, pH, Temperature, Salt, Protein Concentration/Ratio Optimization The first method of optimization is always setting up conditions that vary the original crystal condition discovered during initial screening, in this case IndexTM G12. For RioK3-Short, this general optimization method was performed methodically varying the precipitant concentration (16-30% PEG-3350), buffer pH (0.1 M HEPES pH 5.5-9.0), and salt concentration (0.1-0.4 M MgCl2) independently of each other in a 24-well hanging drop Qiagen Cryotool (crystal plate). Also, concentration of RioK3-Short ranged from 8-30 mg/ml, as crystal size and quality can vary greatly depending on protein concentration. The temperature of crystal tray incubation was performed at 20?C or 4?C. The goal of the optimization was to acquire bigger, more defined crystals. However, these methods of optimization did not work. There was an 82 excess of precipitation in the hanging drops in nearly every condition. What was causing this precipitation and lack of crystal formation? After trial and error and multiple rounds of purification and optimization trays, the reason for this instability became clearer. RioK3-Short was unstable and precipitated when diluted from its original purification buffer containing high salt (500 mM NaCl), high ATP concentration (1 mM) and 2 mM MgCl2. To counteract this dilution, I chose to develop a method of setting up drops that would create a range of dilution (Fig 7.1A). An ?X? of concentrated protein (4 ?L) was created using a pipette tip. At the center of the formed ?X?, 1 ?L of precipitant was added, this is depicted in Figure 7.1A. If the dilution of original condition were the cause of the instability, then this ?X? method would show heavy precipitation in the middle, crystals in the precipitation interface (also known as the metaphase), and finally soluble protein without crystals towards the edges of the ?X?. As seen in Figure 7.1B, this method confirmed RioK3-Short becomes unstable as the original conditions are diluted. Also, this ?X? method of setting up drops, yielded many clean crystals and provided reasoning to set up unconventional ratios of protein:precipitant that was later optimized to 6:1. Although bigger, cleaner crystals were obtained by general optimization, the diffraction was limited 6.0 ?. 7.2B: Minimal Media/ Minimal Media with Se-Met Before changing the construct and introducing mutagenesis, another form of optimization is to modify the method of protein expression. This can be done through expression in minimal media and minimal media containing selenium methionine (Se-Met). Along with experience seen in the lab, trends in literature and recognizing 83 examples at conferences, there is ample supporting evidence that expression in minimal media can improve crystal quality and diffraction (86, 87). It is still unclear and not yet in literature what is the cause of this improvement, but the current hypothesis is the minimal media slows expression and promotes more precise protein folding resulting in a more homogenous protein for better, tighter crystal growth. The presence of a strong diffracting heavy atom (Se-Met), introduced during expression, may also improve diffraction. RioK3-Short was grown in minimal media and also in minimal media containing Se-Met. Bigger cleaner crystals were obtained in minimal media (Fig 6.2B) and resulted in the 5.5 ? diffraction pattern shown in chapter 6 Figure 6.3. The diffraction pattern was indexed with DENZO, part of the HKL2000 program suite, where the point group and unit cell dimensions were determined (Fig 7.2A, B) (56). The unit cell was enormous with dimensions of a=142.0, b=142.9, c=273.5 in the primitive hexagonal point group later determined to be space group P32 during scaling with Scalepack, which is also packaged into HKL2000 (56). Table 7.1 shows the data logfile from Scalepack from a near complete dataset collected from a RioK3-short crystal (Fig 6.2B) grown in minimal media. Although high symmetry is present in the crystal, the Matthews Coefficient (MC) calculation suggests between 13-23 (most likely 18) molecules in the asymmetric unit (88, 89). The MC program uses the space group and unit cell dimensions to calculate a total volume in the cell. After calculating the total volume, the program then uses the expected molecular weight of the protein and predicts the percentage of solvent in the asymmetric unit, or in other words, the specific volume (Matthews Coefficient) contributed from the inputted protein molecular weight. Table 84 7.2 shows Matthews Coefficient output logfile with the number of monomers possible in the asymmetric unit ambiguously between 13-23 monomers corresponding to a solvent content from ~35%-65%. The most common percentage of solvent found in the PDB database is 47% solvent, which corresponds to 18 or 19 monomers for RioK3-Short (89). The oversized unit cell and low resolution diffraction combined with calculations to predict the large number of monomers in the asymmetric unit suggests RioK3-Short crystals were disordered and not packed very tightly. The goal of optimization from this point was to shrink the unit cell and promote tighter crystal packing. 7.2C: Crystal Manipulation (Annealing, Dehydration, Seeding) Crystallography techniques that involve physical manipulation of already grown crystals include annealing, dehydration and seeding. Annealing of crystals is used to eliminate possible disruptions in crystals lattices caused by freezing (90, 91). Cryogenic conditions are supplemented to crystal conditions to protect crystals during freezing, which is required before shooting with high energy x-rays; however, sometimes even in the presence of a suitable cryogenic condition freezing still causes deformation and disorder in the crystal. Specifically, freezing can cause an expansion of the solvent in the crystal and cause disordered crystal packing resulting in poor diffraction. Annealing is the process of warming the crystal to room temperature after flash freezing. Two types of annealing were performed with Riok3-Short crystals. The first, known as annealing on the loop, consists of blocking the cryostream while the crystal is mounted on the goniometer just long enough to thaw and then flash refreeze by unblocking the cryostream (91). This method destroyed 85 nearly all diffraction; leaving just smeared diffraction, which are common in severely damaged crystals. The second, known as macromolecular crystal annealing, was performed by removing a crystal from the cryostream and placed back into the crystal condition containing 20% ethylene glycol for five minutes (91). After sitting for five minutes, the crystal is then looped again and flash frozen. This method of annealing also did not improve diffraction. Dehydration is performed for a completely different reason than annealing. Dehydration is used to remove water molecules from the crystal in order to shrink the unit cell (91, 92). This method was very appealing for RioK3-Short because of the enormous unit cell observed in previous diffraction data. Dehydration was performed via variations of two general methods (91). One method is replacing water molecules with a step-wise increase of dehydration agents, allowing the crystal to equilibrate at each step. Dehydration agents used for Riok3 were glycerol, ethylene glycol and PEG-3350. Crystals of Riok3-Short were looped and transferred to mother liquor (original crystal condition) plus 5% dehydration agent and allowed to sit for 5 minutes. Once equilibrated, the crystal was looped again and transferred to mother liquor plus 10% dehydration agent. These 5% dehydration agent jumps and incubation steps were continued until dehydration agent reached mother liquor plus 30%. One crystal was allowed to rest overnight in the final 30% for each dehydration agent. This method of dehydration requires very robust crystals that can withstand a large amount of manipulation. RioK3-Short crystals did not appear to break down or become damaged during transfers. The other method of dehydration involves a single step allowing air to dehydrate crystals by evaporation of water molecules in a 86 reservoir of mother liquor containing the appropriate cryo condition and/or increased precipitant. RioK3-Short crystals were exposed to air for thirty minutes or one hour in 100 ?L reservoirs containing mother liquor plus 20% ethylene glycol, mother liquor plus 10% PEG-3350, or mother liquor plus 10% ethylene glycol and 10% PEG-3350. Unfortunately, the diffraction of crystals from either method of dehydration did not improve diffraction. All annealing and dehydration experiments were performed on the University of Maryland?s Bruker home source macromolecular diffraction system. While the synchrotron is ideal to see small changes in diffraction quality, the home source was sufficient to determine if annealing or dehydration significantly improved diffraction, which did not occur. Finally, the last type of crystal modification performed on RioK3-Short was seeding. There were two types of seeding attempted, macroseeding and microseeding (91, 93, 94). The names are very telling, as macroseeding is the process of looping a single large crystal, washing the crystal, and then placing the crystal for further growth in either an already equilibrated crystal condition or unequilibrated condition with fresh protein. Microseeding is pulverizing a small crystal into microscopic crystals, and then using a horse or cat whisker to transfer the microcrystals, or diluting the stock seed mixture, into an already equilibrated crystal condition below the precipitant concentration required for crystallization. The goal of microseeding is to set up a range of conditions with a decreasing number of nucleation points (microseeds) by streaking across multiple drops much like sequentially streaking bacteria onto an agar plate, with the purpose of acquiring a single colony. RioK3- 87 Short was subjected to both types of seeding. Microseeding did not yield any crystals, however macroseeding was successful in growing larger crystals. The larger crystals suffered from the same poor diffraction as the original pre-seeded crystals, with diffraction limited to ~6.0 ? when screened at the NECAT beamline. 7.3: RioK3-S199A-Short Crystallography techniques to improve existing crystal diffraction were unsuccessful. Because RioK3-Short was soluble and a successful purification protocol was already determined, the next logical direction was the optimization of the RioK3-Short construct via site-directed mutagenesis. Pure RioK3-Short protein was sent previously for molecular weight determination via mass spectrometry at the University of Maryland Proteomics Core Facility managed by Dr. Yan Wang. The expected molecular weight of RioK3-Short (193-519) is 37,717.2 Da. The resulting spectra confirmed the expected MW of RioK3-short with a peak centered at 37,717.6 Da (Fig. 7.3A). A second peak centered at 37,794.7 Da suggested the presence of a singly phosphorylated version of RioK3-short because the difference between the two masses is the expected change for a post-translational phosphorylation modification. Excess protein was digested with trypsin and analyzed by tandem LC MS/MS by Dr. Yan Wang. Peptide determination of the trypsin fragments identified 84% of the sequence with only one possible site for phosphorylation (Fig. 7.3B). The phosphorylated residue was serine 199. Stratagene?s Quikchange Lightning Site- Directed Kit was used to mutate serine 199 to alanine to abolish RioK3?s ability to autophosphorylate. RioK3-S199A-Short purified identically to RioK3-Short with no deviations. Again, the S199A mutant was screened and surprisingly the same 88 condition in which RioK3-Short crystallized did not grow S199A crystals. Only one condition of 1152, containing 24% PEG-8000, 0.1 M Tris pH 8.0 and 0.2 M MgCl2, grew crystals (Fig 7.4A). These crystals were optimized in the same manner as RioK3-Short and grew much larger (~50x500 ?m) than anything previously acquired from RioK3-Short (Fig 7.4B). Even though the crystals appeared more ordered than the RioK3-Short crystals, the diffraction did not match that same visual improvement. In fact, diffraction images collected at the synchrotron were poorer and provided data to ~8.0 ?. 7.4: Surface Entropy Mutations Surface entropy reduction (SER) is an accepted practice for improving crystallization chances for hard to crystallize proteins (95, 96). After the failure to improve diffraction past 5.5 ? over all previous forms of optimization, the RioK3 project needed a new crystal form, and SER could provide the change necessary to promote crystallization in a new form. SER is achieved by mutagenesis of surface residues with high conformational flexibility and thus a high amount of surface disorder/entropy. Mutation of these residues, which are typically lysine or glutamate, to alanine reduces this flexibility and reduces the entropy on the surface of the protein. The less disorder on the surface of the protein promotes tighter packing during crystallization, which leads to better diffracting, higher resolution crystals. It was shown that sequential mutations of residues directly next to each other improves the likelihood of crystallization and an online server (SERp) from UCLA can identify good targets for mutagenesis (95, 97). For RioK3, SERp identified two locations, each a series of three residues that are most likely to improve crystallization. 89 Residues 222-224 and 276-278 had the sequences Glu-Lys-Lys (EKK) and Glu-Lys- Glu (EKE), respectively. To make these mutations, Qiagen?s QuikchangeTM Lightning kit was used with RioK3-Short as the template DNA. For both locations, the mutants were first double mutated yielding constructs RioK3-KK223AA and RioK3- KE277AA, and then the third mutation was introduced creating RioK3-EKK222AAA and RioK3-EKE276AAA. For all four mutated constructs, there was no difference in the expression and successful purification protocol from the original RioK3-Short fragment. All four mutants were concentrated with no issue and subjected to crystallization screening via the PhoenixTM. There were no crystals observed for RioK3-KE277AA and RioK3-EKE276AAA. Crystals were observed for RioK3- KK223AA and RioK3-EKK222AAA in the same condition and same crystal form as the original RioK3-Short. The inability to acquire a new crystal form from SER was disappointing and suggested an entirely new direction had to be taken to acquire a new crystal form for RioK3. 7.5: Limited Proteolysis RioK3-Short, RioK3-S199A-Short, and all the surface entropy mutants were unsuccessful. The optimization methods described above exhausted crystallization trials for the 193-519 constructs. Focus shifted to determine a new RioK3 fragment that was both soluble and purifiable. The RioK3 full-length construct was expressed with a histidine tag in E.coli overnight and purified by affinity chromatography. Originally, the RioK3 full-length construct was insoluble. However, lessons learned from RioK3-Short (addition of ATP and MgCl2) yielded small amounts of near pure soluble protein (~1 mg/L culture). The quantity of RioK3 full-length was not suitable 90 for crystallization trials, but fitting for limited proteolysis using thermolysin, trypsin, and elastase. The goal of the assay was to determine a stable construct that could be used for further crystallization trials. The time based incubation assay was designed to identify a gel band that was consistent over more than one time point and seen in more than one protease. For trypsin and thermolysin, 50 ?L of Riok3 full-length concentrated to 1 mg/mL was mixed with 9 ?L of enzyme buffer containing 100 mM Tris pH 8.0, 100 mM NaCl, 5 mM CaCl2 and 1 ?L of protease. For elastase, the enzyme buffer only contained 200 mM Tris pH 8.0 and was set up in the same volumes. The reaction was incubated at both 37 ?C and on the bench at ~25 ?C, with 15 ?L samples taken at 5, 10, 30, and 120 minutes. The proteolysis reaction was stopped through denaturation by the addition of gel loading buffer containing SDS and heated to 95 ?C. SDS-PAGE was used to analyze the samples (Fig 7.5A, B). Lane 3 from the elastase gel in Figure 7.5A shows a slice cut from the gel, highlighted by an arrow. This slice was sent to Dr. Yan Wang for complete in-gel tryptic digest and peptide determination via LC MS/MS. Dr. Wang was able to identify coverage from the gel band for residues 248-452 with 67% sequence coverage over that range. This result suggested a new design for the short constructs with deletions from both the N-terminus and the C-terminus. When combined with additional information from previously reported Rio1 archaeal structure and PSI- PRED secondary structure prediction tool, three new constructs (248-519, 248-474, and 193-474) were cloned with primer sets 5'- GAGAACCTGTACTTCCAGGGAATGTTGGAGACAATCACTGGC-3' and 5'- GGGGACCACTTTGTACAAGAAAGCTGGGTTATCATTCATCATATAGTAGT 91 GGTGGGTC for 248-519, 5'-GAGAACCTGTACTTCCAGGGAATGTTGGAGAC AATCACTGGC-3' and 5'-GGGGACCACTTTGTACAAGAAAGCTGGGTTATCA TGTGATGTTTAAGCCTGAAACAGC-3' for 248-474, and 5'-GAGAACCTGTACT TCCAGGGAATGGATTTAAAACTATCAAAC-3' and 5'-GGGGACCACTTTGTA CAAGAAAGCTGGGTTATCATGTGATGTTTAAGCCTGAAACAGC-3' for 193- 474 utilizing previously described Invitrogen?s GatewayTM cloning (Fig 7.5C) (35, 98, 99). These three constructs were subjected to expression and solubility testing. RioK3-248-474 and RioK3-248-519 were completely insoluble and could be collected in the cell pellet. RioK3-193-474 expressed and was partially soluble for the first step of purification. However, upon elution from the affinity column, the soluble protein immediately precipitated. The buffers for RioK3-193-474 were modified to prevent precipitation by varying the components (200 mM and 500 mM NaCl, ?1 mM ATP, ?10% glycerol). Buffer variations were unable to overcome RioK3-193- 474?s insolubility. 7.6: RioK3 Conclusions My goal of solving the structure of RioK3 was an exhaustive lesson on the difficulties of both crystallography and protein purification. The only way to learn the ?tricks? of crystallography, whether it be surface entropy mutations or an outside the box ?X? method of optimization, is to gain experience through a challenging project. Ideally, one of the methods employed to improve diffraction works. However, this was not the case for RioK3. I confidently leave this project knowing the next person can successfully take lessons learned from my trials and apply that knowledge to acquiring a structure. I leave the RioK3 project with multiple soluble 92 constructs and a roadmap toward success based on the results from past experiments presented in this manuscript. My successes with expression, purification and crystallization overshadow the lack of a ?hero? crystal. The next researcher fortunate enough to take over this project will be one step away from success. Maybe that researcher will continue to eliminate possible paths as I have during my tenure, or maybe that researcher will choose correctly on their first try. In any case, I believe my work will lead to success in the near future. 93 Figure 7.1: ?X? Method A) Cartoon representation of ?X? method. Protein is streaked into the formation of an ?X? with pipette tip and then precipitant is added to the center of the ?X? as shown. B) Image taken with digital camera of crystals growing in the metaphase region created by dilution between precipitant near the center and soluble Riok3- Short on the corners. The box in (A) represents the location of the image taken for (B). 94 Figure 7.2: Indexing of Riok3-Short A) Diffraction image from Riok3-Short crystal grown in minimal media showing the fitment of spots after refining in the primitive hexagonal point group. B) Choices of point groups determined form Riok3-Short diffraction image in (A). The highest symmetry with the lowest score (primitive hexagonal colored green) was chosen and confirmed during scaling, using Scalepack. DENZO, a program within the HKL2000 suite, was used for indexing (56). 95 96 Table 7.2: Matthews Coefficient Logfile Matthews Coefficient logfile showing the number of possible monomers in the asymmetric unit, based on MW 37,717 Da. Highlighted in yellow are the most likely number of monomers. 97 Figure 7.3: RioK3-Short MW Determination and LC MS/MS A) Molecular weight determination of RioK3-Short. The expected MW for RioK3- Short is 37,717 Da. B) Results from peptide identification via tryptic digestion followed by LC MS/MS. Red residues indicate identified residues with a box around serine 199, the only phosphorylated residue identified. Dr. Yan Wang at the University of Maryland Proteomics Center performed all mass spectrometry for RioK3. Data was analyzed using Mascot (www.matrixscience.com). 98 Figure 7.4: RioK3-S199A-Short Crystals A) Digital image of RioK3-S199A-Short crystals grown in the Wizard II screen position E3, containing 24% PEG-8000, 0.1 M Tris pH 8.0, 0.2 M MgCl2. B) The largest crystals grown for RioK3. This image was photographed from an optimization tray containing 20% PEG-8000, 0.1 M Tris pH 8.5, 0.2 M MgCl2. 99 Figure 7.5: Limited Proteolysis of Riok3 Full-length SDS-PAGE of RioK3 full-length subjected to limited proteolysis with elastase, shown in (A), trypsin or thermolysin, both shown in (B). Horizontal arrows point to band of interest and the red diagonal arrow in (A) points to gel slice removed for mass spectrometry. C) Representation of fragments created from limited proteolysis assay (green = 248-519, purple = 248-474, blue = 194-474, black = original Riok3- Short 193-519). MVNSGM cuts off the ?extended? archaeal Rio1 domain, supported by the MS/MS results (35). RELF-NIT is homologous to the last archaeal Rio1 alpha helix I (35). ?????? represent unfolded region predicted from PSI-PRED (99). 100 Chapter 8: My Role in Ubiquitin 8.1 Overview and Thanks I would like to thank Dr. David Fushman at the University of Maryland, a member of the Center for Biomolecular Structure and Organization (CBSO), for the opportunity to learn and teach while working on his Ubiquitin projects. My role in Ubiquitin has been two fold. First and foremost, my responsibility was to provide advice and consultation to members of his laboratory who had an interest in learning the basics of crystallography. Shirley Lee, current graduate student at the University of Michigan was the primary contact for Dr. Fushman. My second purpose for the multiple Ubiquitin projects was to provide expertise and resources not available to Dr. Fushman. This goal was met by providing beamtime during synchrotron trips and access to instrumentation in the Dr. LaRonde-LeBlanc lab, along with guidance on all shared instrumentation. It should be noted that my role in Ubiquitin was strictly crystallographic in nature, meaning I only provided suggestions and guidance in crystal growth/optimization, crystal harvesting, data collection, and structure solution. I was not responsible for project direction or the selection of crystallographic targets. Many of the targets chosen by Dr. Fushman were site-directed mutants of human Ubiquitin, designed to look at confirmation changes introduced by the selected mutations. Three of these mutants provided crystals that diffracted to 2.0 ? or better. In chronological order, Ubiquitin leucine 69 to serine (L69S), isoleucine 13 to serine (I13S), and double mutant lysine 63 to aspartatic acid with glutamatic acid 64 to glycine (KE63DG) were all selected for x-ray structure determination. Near the end 101 of my tenure at the University of Maryland, I also helped with the structure determination of lysine 48-linked di-ubiquitin in complex with E2-25K, the protein known to be responsible for creating chains of ubiquitin through a covalent bond between lysine 48 and glycine 76 (100). The following is a description of results from my time spent with Dr. Fushman and his laboratory members. 8.2: Methods for Structure Determination of Ubiquitin Mutant I13S Initial crystallization was done via sparse matrix screening (Wizard I & II & IIITM, NatrixTM, PEGTM Suite, CryoTM Suite, IndexTM) using Art Robbins PhoenixTM high-throughput robot. The Phoenix utilizes the sitting drop method for screening for crystallization conditions. Ubiquitin I13S mutant crystals grew in 5-7 days. Optimization of crystals screened for diffraction were done via hanging drop method using a 24-well tray purchased from Qiagen. Crystallization conditions for Ubiquitin I13S mutant include 0.3M Ammonium Iodide, 26% PEG 3350, 100mM Hepes pH 6.8, and 20% glycerol. Crystals were screened for diffraction and cryo conditions at the home souce (Microstar H2 generator with Proteum Pt135 CCD detector) at 100 K. Diffraction data sets were collected via Advanced Photon Source (APS) located at Argonne National Laboratory NECAT Beamline 24IID. Ubiquitin crystal datasets were collected at a wavelength set at 1.00931 ?. All data collected was integrated and scaled using HKL2000 (Table 8.1) (56). The human Ubiquitin monomer (PDB: 1ubq) was used as the search model for molecular replacement using Phaser within CCP4, for the structure solution of Ubiquitin I13S mutant (66, 70, 101). Refinement of the model was done primarily using Refmac5 within CCP4 and model building was done using COOT (64-66). The final solution intended for submission was 102 refined with PHENIX (72). I13S crystallized with four molecules in the asymmetric unit in space group P21. Figure 8.1 shows an overall cartoon view of the asymmetric unit. Statistics for I13S data collection and refinement are shown in Table 8.2. 8.3: Ubiquitin Mutants L69S and KE63DG Crystals were obtained for L69S and KE63DG in the same manner as I13S (screening then optimization) and data was collected at both the home source and synchrotron. Unfortunately, both L69S and KE63DG were not solved due to problems with data quality. Specifically, both crystal forms used for data collection suffered from pseudosymmetry-related twinning that could not be resolved. Pseudosymmetry confuses space group selection due to non-crystallographic symmetry being close to crystallographic symmetry. This results in two or more possible spacegroups contained within the crystals. In this case, we were unable to successfully de-twin the data and molecular replacement could not effectively place molecules to provide accurate phases. This resulted in poor electron density and incorrect solutions. The first approach to resolving pseudosymmetry is going back to the original data to determine if there is a possibility that this space group confusion is caused by overlapping crystals, otherwise known as non-merohedral twins (common in plate crystals), thus giving two distinct overlaid diffraction patterns. After taking a closer look at diffraction patterns and scaling statistics, multiple crystals did not appear to be the cause of twinning. The presence of a single lattice in the diffraction pattern and confusion between symmetry suggested pseudo-merohedral twinning. Pseudo- merohedral twinning is the presence of multiple lattices of differing symmetries 103 directly overlaid on top of each other giving the appearance of a single crystal lattice. The next approach in this type of twinning was to re-index the collected data into lower symmetry, in this case primitive monoclinic (P2) instead of primitive orthorhombic (P222). Initially, the most likely space group determined for both L69S and KE63DG was P212121 (orthorhombic), which can be confused with P21 (monoclinic) in a case of pseudosymmetry. Ideally, the elimination of symmetry not actually present should result in a usable dataset. This was not the case for the Ubiquitin mutants evident by the continued failure of molecular replacement. This suggested a more severe partial twinning that would require separation of the twinned data using a partial twin law. I was never successful in finding an appropriate method for solving these mutant structures. In hindsight, work should have focused on the growth of many crystals and required full-time dedication to optimization of these twinned Ubiquitin crystals. 8.4: E2-25K in Complex with Di-ubiquitin Near the end of my time spent with Shirley Lee, she grasped many of my teachings and was fully capable of making decisions and providing assistance to other members of Dr. Fushman?s laboratory. Ms. Lee provided her knowledge to help Dr. Rajesh Singh, postdoctoral reseacher in Dr. Fushman?s lab, prepare E2-25K and lysine 48 linked di-Ubiquitin protein complex and set up screening trays using the PhoenixTM liquid handler. Shortly after setting up the initial screening, Ms. Lee left the University of Maryland and beginning graduate school at the University of Michigan. I picked up where Ms. Lee left off and began work with Dr. Singh. The results from the screen were very encouraging with over 50 possible targets for 104 optimization. There were two distinct crystals forms, clean plates and circular ?blob? crystals (Fig 8.2A, B). I utilized the home diffraction source to collect a data set from both plate crystals and a ?blob? crystal and confirmed the presence of the complex only in the ?blob? form via a low-resolution (3.0 ?) structure solved by molecular replacement (72). This structure in P212121, solved with E2-25K and one monomer of Ubiquitin in the asymmetric unit. Crystallographic symmetry was present between the two monomers of linked Ubiquitin, resulting in the single monomer of Ubiquitin. There was no discernible electron density to model residues 73-76 between the Ubiquitin monomer and its symmetry related mate, and the covalent linkage between lysine 48 and glycine 76. This could have been due to low-resolution diffraction, the crystallographic axis between the two monomers, or flexibility in the short polypeptide connecting the Ubiquitin monomers. The synchrotron was needed to collect higher resolution data. Crystals harvested from Qiagen?s CryosTM Suite containing 10.2% PEG-20000, 0.085 M MES pH 6.5 and 15% glycerol, diffracted to 2.3? at the NECAT beamline (Table 8.3). Molecular replacement was performed with Phaser within PHENIX, using a previously solved crystal structure of E2-25K and ubiquitin complex (PDB: 3K9P) (70, 72, 102). In space group P21, there are two monomers of E2-25K and two monomers of Ubiquitin (Fig. 8.3A). Residues 74-76, including the covalent linkage between lysine 48 and glycine 76, still could not be modeled due to poor electron density insufficient for accurate placement of those residues (Fig 8.3B). This suggests high conformational flexibility for these residues. Statistics for E2-25K and Ubiquitin complex are shown in Table 8.2. 105 8.5: Ubiquitin Conclusions Again, it has been an incredible opportunity to provide assistance to Ms. Lee, Dr. Singh, and Dr. Fushman. Ubiquitin has truly been a lesson in both the successes and pitfalls of crystallography. 106 107 Figure 8.1: Ubiquitin I13S Overall View Cartoon Representation of the four Ubiquitin monomers (colored cyan, green, yellow, and magenta) in the asymmetric unit. Serine 13 is mutated from isoleucine and shown highlighted in red sticks. 108 Table 8.2: Ubiquitin Data Collection and Refinement Statistics 109 Figure 8.2: Possible Crystals Containing E2-25K and Di-Ubiquitin Complex A) Digital image of one of many conditions that grew ?blob? like crystal form. This crystal form was confirmed to contain E2-25K and di-Ubiquitin complex. B) Digital image of plate like crystals that also grew in many conditions (>30). This crystal form was later confirmed at the synchrotron to only contain Ubiquitin and not the E2-25K and di-Ubiquitin complex. 110 111 Figure 8.3: E2-25k and Di-Ubiquitin Complex A) Cartoon representation of the contents of the E2-25K and di-Ubiquitin complex asymmetric unit, which includes two molecules of E2-25K (magenta) and two molecules of Ubiquitin (green). B) Electron density 2Fo?Fc map contoured at 1?, 112 showing the gap in electron density between the last built residue (leucine 73) and the expected covalent bond with lysine 48. 113 Chapter 9: Crystal Structure of Iodotyrosine Deiodinase, a Novel Flavoprotein Responsible for Iodide Salvage in Thyroid Glands 9:1: Overview Chapter 9 includes portions of the work published with Dr. Patrick McTamney, Jennifer Adler, and Dr. Steven Rokita. As first author on the article, and author of this dissertation manuscript, it is my commitment to discuss my responsibilities for the project and those responsibilities for which I was not the primary scientist. This chapter will include modified text from the article including crystallographic methods (crystal screening, data collection, structure determination and refinement). Included, and not present in the published article, are crystallographic tables (Tables 9.1, 9.2, 9.3-9.5) representing the original data used to solve the protein structures. Results and Discussion has been limited here, the full article can be found through the Journal of Biological Chemistry (103). Dr. Patrick McTamney was responsible for all expression and purification of iodotyrosine deiodinase (IYD) protein. His work to achieve a soluble construct was exemplary and not described in this dissertation manuscript. The full experimental procedure for expression/purification work can be found through the University of Maryland library system in Dr. Patrick McTamney?s recently published dissertation (2009, http://hdl.handle.net/1903/9848). Jennifer Adler contributed to growing crystals and provided an excellent figure depicting the alignment of secondary structure between IYD and family members. This alignment figure is not included with from the following and can be 114 found online in the official published copy from the Journal of Biological Chemistry (104). As senior author and the principle investigator for this project, Dr. Steven Rokita provided expert discussion and background knowledge for the IYD project. Dr. Rokita wrote all active site mechanistic results and discussion. Also Dr. Rokita provided an in-depth introduction, which was essential in providing the basis for structural analysis and guidance throughout the article. Finally, I have to acknowledge my advisor Dr. Nicole LaRonde-LeBlanc. She drove the crystallization and structure determination part of this project to success and provided all the necessary resources, both in terms of expertise and instrumentation, to complete this work. 9.2: Statement of Authorization This research was originally published in Journal of Biological Chemistry. Seth R. Thomas, Patrick M. McTamney, Jennifer M. Adler, Nicole LaRonde- LeBlanc, Steven E. Rokita. Title Crystal Structure of Iodotyrosine Deiodinase, a Novel Flavoprotein Responsible for Iodide Salvage in Thyroid Glands. Journal of Biological Chemistry. 2009; 284(29):19659-67. ? the American Society for Biochemistry and Molecular Biology. 9.3: Introduction Iodotyrosine deiodinase (IYD) salvages iodide from mono-iodotyrosine (MIT) and di-iodotyrosine (DIT) (105, 106). MIT and DIT are by-products from the proteolysis of thyroglobulin, a large protein (>300kD) responsible for the 115 biosynthesis of thyroxine (105). IYD is responsible for salvaging the iodide from MIT and DIT before excretion via a dehalogenation reaction for the purpose of recycling the iodide back into thyroglobulin, and subsequently the biosynthesis of thyroxine in the thyroid. If iodide is not recycled properly by IYD, then the acquisition of iodide from outside sources (diet) becomes the only pathway to gather iodide into the thyroid (107, 108). If for any reason the quantity of available iodide is insufficient for the biosynthesis of the essential thyroid hormones, then the unfortunate individual may suffer from hypothyroidism. Weight gain, fatigue, depression, weakness, constipation, and many other symptoms characterize hypothyroidism, resulting in a very serious condition (109). Currently, the most successful treatment for hypothyroidism is an oral supplement to replace insufficient levels of thyroxine (109). However, the treatment does not help everyone and more fundamental research needs to be performed to better understand the homeostasis of iodide in the human body. Presented in this chapter are three structures of IYD, one with IYD and cofactor flavin mononucleodite (FMN) and two structures solved in the presence of both cofactor and their substrate MIT or DIT, at 2.0 ?, 2.45 ?, and 2.6 ?, respectively. These structures provide an understanding for IYD mutations found in patients suffering from hypothyroidism. 9.4: Experimental Procedures 9.4A: Gene Construction of a Soluble and Affinity-tagged IYD Gene construction of the soluble fragment used for crystallography is attributed to the hard work of Dr. Patrick Mctamney and Dr. Steven Rokita. The full 116 description of cloning can be accessed in the experimental procedures section of Thomas et al. (103) and in Dr. Patrick McTamney?s dissertation (2009, http://hdl.handle.net/1902/9848). Briefly, the successful construct was truncated by 33 amino acids on the N-terminus to remove the membrane anchor domain and included an uncleavable His6 tag on the C-terminus for purification. This truncation resulted in a soluble construct that met the purity and quantity requirements needed for crystallography. 9.4B: Expression and Purification of IYD Dr. Patrick McTamney and Dr. Steven Rokita were responsible for the expression and purification of IYD. One of the major factors in determination of the feasibility of IYD structure determination was their ability to provide ample amount of pure protein for crystal screening and optimization. The full description of the expression of IYD in Sf9 cells and purification by affinity chromatography can be accessed in the experimental procedures section of Thomas et al. (103) and within Dr. Patrick McTamney?s dissertation (2009, http://hdl.handle.ne/1902/9848). 9.4C: Crystallization Initial crystallization was explored with sparse matrix screening (WizardTM I, II, and III (Emerald Biosciences); PEGSuiteTM and CryoSuiteTM (Qiagen); and NatrixTM and IndexTM (Hampton Research)) using an Art Robbins PhoenixTM high throughput liquid handler. Ultimately IYD was crystallized at 20 ?C in three different crystal forms by the hanging drop vapor diffusion method using two parts IYD (16 mg/ml; 10 mM potassium phosphate, pH 7.4) to one part reservoir solution. IYD 117 crystals were obtained using a reservoir solution containing 20% (w/v) polyethylene glycol 3000, and 0.1 M sodium acetate, pH 4.5. IYD?MIT co-crystals were obtained by supplementing the enzyme solution with 2 mM MIT prior to addition of the reservoir solution containing 0.2 M magnesium chloride, 20% (w/v) polyethylene glycol 3350, 10 mM potassium phosphate, pH 7.4. IYD?DIT co-crystals were obtained by supplementing the enzyme solution with 2 mM DIT prior to addition of the reservoir solution containing 0.2 M ammonium acetate, 45% 2-methyl-2,4- pentanediol, 0.1 M BisTris, pH 5.5. In each case, yellow crystals appeared within 24 h. Crystals grew as connected clusters that required dehydration and micro-dissection to separate out single crystals. 9.4D: Data Collection Initial crystals were screened for diffraction and cryo conditions using a Bruker Microstar H2 generator with Proteum Pt135 CCD detector at 100 K. Diffraction data sets were then collected at the Northeastern Collaborative Access Team Beamline 24-ID, Advanced Photon Source, Argonne National Laboratories. Diffraction data for IYD was collected at a wavelength of 1.653 ? for single anomalous diffraction phasing using sulfur atoms. In order to obtain the required redundancy and signal to noise ratio for the sulfur anomalous signal, data sets were collected using varying parameters including exposure time to minimize radiation damage. Diffraction data of co-crystals containing MIT and DIT alternatively were collected at 0.9795 ?. All collected data were integrated and scaled using HKL2000 (56). 118 9.4E: Sulfur Phasing and Refinement All nine sulfur positions within IYD were found using SHELX within HKL2MAP program suite (Table 9.1) (110-112). Heavy atom positions were refined, and initial phases were determined using the AutoSHARP program set (63). Initial model building was performed by WARP within AUTOSHARP (63). Because of the complexity of the monomeric fold, nearly half of the residues were not built and required manual building. Several rounds of model building and refinement were needed for IYD using Refmac5 with TLS refinement and COOT for molecular visualization and rebuilding (64-66). A monomer of IYD was used as a model for molecular replacement to solve the structure of the co-crystal containing MIT. One dimer of this co-crystal was used in turn as the model for molecular replacement to solve the co-crystal containing DIT. Molecular replacement was performed by PHASER within the CCP4 program suite (66, 70). These structures were also rebuilt and refined by iterations of Refmac5 and COOT (64). IYD?DIT was initially solved in space group P212121 and suffered from pseudosymmetry, evident after multiple rounds of refinement with high R-factors. Re-indexing IYD?DIT diffraction data into primitive monoclinic and subsequent determination of the structure in P21 removed the overestimated symmetry and allowed for proper refinement. Refinement statistics are provided in Table 9.2. Tables 9.3-9.5 show data collection log files for all three structures matching those submitted with Table 9.2. 119 9.5: Results 9.5A: Overall Structure of the Soluble Domain of IYD In the absence of substrate, IYD crystallized with one monomer per asymmetric unit. However, analysis of the crystal packing interactions revealed an extensive interface with a symmetry-related molecule (2490 ?2 per monomer) indicating that the enzyme forms a domain-swapped dimer as observed in related proteins within the same structural superfamily (113-116). The IYD?MIT complex crystallized with two monomers per asymmetric unit, and the IYD?DIT complex crystallized with eight monomers per asymmetric unit. All three structures were solved from distinct crystal forms (Table 9.2). Electron density for the first 34 amino acids of the truncated enzyme was not observed for any of the three crystals. These residues in the native protein likely provide a flexible linker between the soluble domain and the N-terminal anchor embedded in the membrane. As mentioned, IYD contains the characteristic ?-? fold that is common to all proteins of the NADH oxidase/flavin reductase superfamily (Fig. 9.1, Supplementary Fig. S1). The closest structural neighbor to IYD as determined by DaliLite (117) is BluB (Protein Data Bank code 2ISK) (116) with an (RMSD) of 3.1 ? for 198 structurally equivalent residues of 219 possible residues. Despite the structural similarities, their sequence identity is quite low (19%). The net catalytic turnover of BluB appears to be very different from IYD, and BluB has been recently identified as the source of the lower ligand (5,6-dimethylbenzimidazole) of vitamin B12 by sacrificing its bound FMN cofactor to oxidation (116). Still IYD and BluB form similar dimer interfaces at their core with criss-crossing helices and similar domain 120 swaps with their extended N- and C-terminal fingers (Supplementary Fig. S2). Upon binding substrate, MIT/DIT for IYD and molecular oxygen for BluB, the lid forms holding in the substrate near FMN. The differences between the IYD and BluB active site lid positioning may determine the specificity and explain the difference in function (Fig. 9.2). Blub active site lid caves in much closer to the FMN molecule, which would not be enough room for MIT or DIT to fit in the active site. 9.5B: Interface Structure of the Homodimer For IYD, N- and C-terminal extensions of each polypeptide wrap around the other. These extensions span distances of greater than 37 ? and comprise a minimum of 26 amino acids near the N-terminus and 18 amino acids at the C-terminus. Two equivalent active sites are located within the dimer interface, and each active site is comprised of residues from both subunits. Accordingly subunit association is essential for FMN binding and catalytic activity. Because the IYD structure was solved as a monomer per asymmetric unit, its dimer was generated by crystallographic symmetry, and consequently no differences between the two monomers were observed. Differences observed between the monomeric units of IYD?MIT and IYD?DIT were minimal as evident from their root RMSD values of 0.196 ? and 0.254 ?, respectively. 9.5C: Substrate-induced Conformational Changes in the Active Site Two unstructured regions were identified within substrate-free IYD by the lack of electron density corresponding to residues 156?177 and 195?208. Even in the substrate-bound co-crystals, the regions containing residues 195?208 exhibit 121 relatively weak electron density compared with other regions of the molecule. Only portions of this region, some with higher than average B-factors, could be built for IYD bound to substrate. In contrast, residues 156?177 gain detectable structure based on the additional electron density observed for the co-crystals IYD?MIT and IYD?DIT. Substrate binding appears to induce the extension of one helix (?C) and the formation of another short helix (?D) in this region to cover the active site and protect substrate and flavin from solvent (Fig. 9.1B, D). This active site lid is stabilized in part by numerous interactions with the bound substrate that is in turn anchored by aromatic stacking with the isoalloxazine ring of FMN as well as by polar contacts with FMN and Ala-126 (Fig. 9.1E). Eight residues of the lid assemble within 4 ? of the substrate, and three of these form polar contacts (Glu-153, Tyr-157, and Lys-178). An analysis of sequence conservation onto the surface of IYD indicates a high level of conservation on one face of the molecule, but not the other. I have designated those as front and back faces for analysis. Figure 9.3A shows the conserved face (front) of the protein along with solvent accessible FMN. As expected, there is significant residue conservation near the active site, and this conservation extends across this face. It is assumed that IYD requires interaction with a currently unknown reductase that binds and reduces the FMN. Therefore, the remaining conservation on the front face may be required for interaction with the reductase. The reductase would presumably bind to IYD and FMN when ligand is not present. Figure 9.3B displays the conserved surface when MIT is bound and the helical flap encloses the active site. The disappearance of the majority of the conserved surface supports the idea that the 122 conserved surface would only be required and open for binding when the substrate is absent. Electrostatic comparisons between IYD?Holo and ligand bound IYD?MIT show distinct structural changes, both in shape and electrostatic charge density on the front face of the protein in response to substrate binding (Fig. 9.1C, D). It is clear that upon binding ligand the protein becomes more structured around the active site and the face of the protein becomes more defined with hydrophobic residues near the center and postively charged residues around the edges. This analysis also highlights that there is ample solvent accessibility to the FMN in the IYD?Holo to allow for substrate binding and interaction with a reductase. Both MIT and DIT are stabilized in the active site by electrostatic interactions, expected by the conserved positively charged arginine residues near the phosphate tail of FMN and the negatively charged area surrounding the amino group of the substrate (Fig. 9.4A). The active site lid forms locking in the substrate near the FMN cofactor and provides just enough space for the specific substrate to stack on top of the FMN molecule. There is a slight conformational change of three nearby residues (Leu-169, Thr-174 and Leu-172) to allow enough room for DIT to fit into the active site (Fig. 9.4B). 9.6: Structural Basis for Deficiency of IYD in Humans Moreno et al. discovered four mutations within the IYD gene from patients suffering from hypothyroidism (106). All mutations discovered are conserved in the mouse IYD gene, along with over 90% sequence identity between mouse and human homologs. This sequence conservation between human IYD and mouse IYD allows 123 for the acceptable explanation of the phenotypes expressed by the human mutations using the mouse IYD crystal structures. Mutations discovered at Arg101 to Trp (mouse: Arg97) and Phe105 and Ile106 to Leu (mouse: Phe101-Ile102) were observed in patients, and shown to completely abolish deiodinase activity (106). Arg97 is located at the rear of the active site, forming hydrogen bond to the phosphate group on the FMN cofactor (Fig. 9.5). Mutation of Arg97 to tryptophan would leave no room for FMN to bind correctly, resulting in the abolished activity. Phe101-Ile102 are also located near the active site (Fig. 9.5). Neighboring residue Arg100 extends and interacts with FMN intimately, with interactions to the 2?-ribityl hydroxyl group, N-1 and O-2. Mutations of Phe101-Ile102 to leucines would change the local structure of the loop and eliminate Arg100 interaction with FMN (Fig. 9.5). The fourth mutation, Ile116 (mouse: Ile112) to Thr was found in patients but still exhibited some deiodinase activity. Ile116 is not located near the active site but is buried within the protein in a hydrophobic core, clear from any interactions of the active site (Fig 9.5). Introducing threonine into the hydrophobic environment could create instability in the protein and could explain the decreased activity, albeit less severe. 9.7: IYD Acknowledgements This work is based upon research conducted in part at the Northeastern Collaborative Access Team beamlines of the Advanced Photon Source supported by Award RR-15301 from the National Center for Research Resources at the National Institutes of Health. Use of the Advanced Photon Source is supported by the United States Department of Energy, Office of Basic Energy Sciences, under Contract DE- 124 AC02-06CH11357. We are grateful to the excellent staff at the Northeastern Collaborative Access Team for assistance with data collection and analysis. 125 Table 9.1: ShelX Results from HKL2MAP Program Suite. The nine sulfur atoms are identified as shown above the line in the ShelXE results logfile, with the strength of anomalous signal present shown in the ShelXC results logfile (111, 112). 126 Table 9.2: Data Collection and Refinement Statistics Statistics for last shell are shown in parenthesis. 127 128 129 130 Figure 9.1: IYD structure A) An overall view of the native homodimer of IYD crystallized in the absence of substrate. Each monomer is distinguished by color. Disordered regions consisting of residues 156?177 and 195?208 connect to the structure as indicated by ? and ?, respectively. B) Native homodimer of IYD crystallized in the presence of its substrate, MIT. Only the structure induced upon substrate binding is highlighted in 131 the colors of the monomers shown in (A). The surface properties of IYD (C) and its complex with monoiodotyrosine (D) were calculated using vacuum electrostatics in PyMOL (75). Blue indicates positive charge, and red indicates negative charge. E) Ionic interactions and hydrogen bonding stabilize the FMN?monoiodotyrosine complex formed by IYD. F) The interaction between FMN and MIT in the active site of IYD. An Fo ? Fc electron density map calculated after refinement in the absence of FMN and MIT is shown contoured at 3?. 132 Figure 9.2: Structural Overlay of IYD and BluB Structural differences between IYD?MIT (gray) and BluB (blue) are highlighted using an overlaid stereoimage. Structural changes near the active site are depicted with the schematic representation. The full-view overlay can be seen as a stereoimage in the supplemental information. 133 Figure 9.3: IYD Sequence Conservation A) and B) shows front views of sequence conservation modeled onto the surface of IYD?Holo and IYD?MIT, respectively. Red=sequence identity. Green=strongly similar. Blue=weakly similar. Gray=no similarity White=FMN surface. C) and D) are rotated 180? around the x-axis in respect to (A) and (B). ClustalW used for alignment from the following: M. musculus (mouse), X. laevis (frog), D. rerio (zebra fish), N. vectinsus (sea anemone), S. purpuratus (sea urchin), D. melengaster (fruit fly), and C. elegans (worm). 134 Figure 9.4: IYD Active Site Electrostatics and MIT/DIT Alignment A) The surface characteristics of the active site of IYD for the IYD?MIT co-crystal crystal calculated using vacuum electrostatics in PyMOL (47). Blue indicates positive charge, and red indicates negative charge. B) Alignment of active site structures of IYD bound with MIT (orange) and DIT (cyan) illustrates the minor conformational change required to accommodate the larger substrate. 135 Figure 9.5: Mapping Human Mutations onto the Structure of IYD Native residues of IYD (M. musculus) highlighted in red correspond to sites associated with human mutations identified clinically to cause hypothyroidism (106). 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R., Walsh, C. T., and Walker, G. C. (2007) Blub cannibalizes flavin to form the lower ligand of vitamin B12. Nature 446, 449-453. 117. Holm, L., and Park, J. (2000) Dalilite workbench for protein structure comparison. Bioinformatics 16, 566-567. Seth R. Thomas 3400 Pennsylvania St. Hyattsville, MD 20783 937-623-3653 sethomas101@gmail.com EDUCATION Ph.D in Biochemistry. 2006-Current University of Maryland College Park. Department of Chemistry and Biochemistry Doctoral Graduate Student, Dr. Nicole LaRonde-LeBlanc B.S. in Biochemistry. 2006 The Ohio State University Bachelors of Science Biochemistry Minor: General Business Administration University of Maryland College Park, 2006-Current Graduate Research Assistant/Teaching Assistant ? Research Experience: My research focus at the University of Maryland has been on structural studies of ribosome processing factors. During my tenure, I have worked on an atypical Rio kinase (Riok3) and Emg1, an N1-pseudouridine RNA methyltransferase. We have shown primary sequence and secondary structure specificity through a crystal structure of Emg1 bound in complex as an obligate dimer with target RNA, containing a base-flipped uridine (substrate analog of pseudouridine to promote crystallization) and a hairpin loop spanning across the dimer interface. I have also collaborated on multiple projects including a breakthrough on a novel structure of iodotyrosine deiodinase (IYD), which salvages iodide from mono- and diiodotyrosine via a dehalogenation mechanism. Without IYD?s salvage pathway in thyroid glands, iodide deficiency can lead to hypothyroidism. The combination of diverse projects has given me the experience needed to pursue a post-graduate research position using what I?ve learned in project design, cloning, protein purification, crystallization, data collection (both home source and synchrotron), and structure solution/refinement. ? Organic laboratory teaching assistant ? General biochemistry teaching assistant JamesCare for Life, Arther G. James Cancer Hospital and Richard J. Solove Research Institute Columbus, OH. 2005-2008 Student Administrative/Research Assistant ? Keep team members informed of current cancer research throughout the world. ? Accelerate responses to research/grant opportunities. ? Benchmark and develop patient support events. The Ohio State University Columbus, OH. 2003-2004 Resident Advisor ? Led 48 students throughout their first year experience ? Facilitated floor meetings and planned/executed floor programs ? Actively Involved in Biological Sciences Living Learning Program TECHNICAL EXPERIENCE ? Structural Biology ? Extensive training in X-ray crystallography including growing/optimizing/freezing crystals, collecting data using the synchrotron and in- house sources (Bruker), data processing (HKL2000, Proteum) and structure refinement (Phenix, CCP4, COOT), experience using both molecular replacement and single wavelength anomalous dispersion (SAD) for phase determination (PHASER, SHELX, Solve/Resolve, ARP/wARP) using both sulfur and selenium anomalous diffraction. ? Biochemistry ? Radioactive labeling/purification of RNA for footprinting/sequencing/nitrocellulose/DEAE binding assays, fluorescence anisotropy using labeled RNA, thermal shift assay using real-time PCR, analytical ultracentrifugation (sedimentation), mass spectrometry, gel-shift assays, Protein/RNA complex crystallization. ? Protein Purification ?Affinity chromatography, gel filtration, ion exchange, widespread experience troubleshooting/protocol optimization of purification protocols. ? Molecular Biology ? Recombinant cloning, PCR, DNA and RNA purification, sequencing, transformations, protein expression, site-directed mutagenesis, mammalian cell culture ? Lab Management ?Maintaining and repairing Bruker x-ray diffraction system, maintain and repair ALL laboratory equipment, including Phoenix high-throughput liquid handler and chromatography systems, laboratory inventory and ordering. PUBLICATIONS 1. Thomas, S. R., McTamney, P. M., Adler, J. M., LaRonde-LeBlanc, N., and Rokita, S. E. (2009) Crystal Structure of Iodotyrosine Deiodinase, a Novel Flavoprotein Responsible for Iodide Salvage in Thyroid Glands. Journal of Biological Chemistry. 284. 19659-19667. 2. Thomas, S. R., Keller, C., Szyk, A. Cannon, J. LaRonde-LeBanc, N. (2010) Structural Insight into the Functional Mechanism of Nep1/Emg1 N1-specific Pseudouridine Methyltransferase in Ribosome Biogenesis. Nucleic Acids Research. Advance online publication. PROTEIN DATA BANK SUBMISSIONS 1. 2009 - 3GB5 M. musculus IYD bound to MN 2. 2009 - 3GFD M. musculus IYD bound to FMN and MIT 3. 2009 - 3GH8 M. musculus IYD bound to FMN and DIT 4. 2010 - 3O7B A. fulgidus Nep1 bound to S-adenosylhomocysteine (SAH) 5. 2010 - 3OII S. cerevisiae Emg1 bound to SAH 6. 2010 - 3OIN S. cerevisiae Emg1 bound to SAH and 1 molecule of RNA 7. 2010 - 3OIJ S. cerevisiae Emg1 bound to SAH and 2 molecules of RNA _______________________________________________________________ AWARDS/FELLOWSHIPS ? Achievement Awards for College Scientists (A.R.C.S.) Foundation. 2009-2010 and 2010-2011 Metro Washington fellowship recipient. ? Jacob K. Goldhaber International Travel Award. American Crystallographic Association 2009 Toronto Conference. Poster presentation. ?Crystal Structure of Iodotyrosine Deiodinase, a Novel Flavoprotein Responsible for Iodide Salvage in Thyroid Glands.? ? ?2005 Making a Difference Award.? Presented to JamesCare for Life team for Outstanding Customer Service. October 2005. RESEARCH PRESENTATIONS ? July 2009. American Crystallographic Association International Conference. Toronto, Ontario, Canada. Poster presentation. ? May 2009. 39th Macromolecular Mid-Atlantic Crystallography Conference. University of Maryland College Park. Poster presentation ? April 2009. A.R.C.S. Seminar and Presidential Luncheon. University of Maryland College Park. Oral presentation. ? November 2008. Bioscience Day. University of Maryland. Poster presentation. ? June 2007. 37th Macromolecular Mid-Atlantic Crystallography Conference. University of Virginia. Poster presentation. REFERENCES Dr. Nicole LaRonde-LeBlanc (Assistant Professor) University of Maryland, Department of Chemistry and Biochemisty Biomolecular Sciences Bldg #296 College Park, MD 20742 301-405-0462 nlaronde@umd.edu Dr George H. Lorimer (Distinquished University Professor) Director of Center for Biomolecular Structure and Organization University of Maryland, Department of Chemistry and Biochemistry Biomolecular Sciences Bldg #296 College Park, MD 20742 301-405-1828 glorimer@umd.edu Dr. Dorothy Beckett (Professor) University of Maryland, Department of Chemistry and Biochemistry Biochemistry Building #091 College Park, MD 20742 301-405-1812 dbeckett@umd.edu