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Cloning and characterization of a novel ferritin from the marine diatom Pseudo-nitzschia multiseries Moccia, Lauren Paul 2008

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CLONING AND CHARACTERIZATION OF A NOVEL FERRITIN FROM THE MARINE DIATOM PSEUDO-NITZSCHIA MULTISERIES by Lauren Paul Moccia B.Sc., University of British Columbia, 2005 A THESIS SUBMITTED TO THE FACULTY OF GRADUATE STUDIES IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Oceanography) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) June 2008 © Lauren Paul Moccia, 2008 ii Abstract Diatoms play a fundamental role in marine food webs, and significantly contribute to global primary production and carbon sequestration into the deep ocean. In many offshore areas of the open ocean, iron (Fe) input is low, and its availability often limits phytoplankton biomass. Recently, gene sequences encoding ferritin, a nearly ubiquitous iron storage and detoxifying protein, have been identified in pennate diatoms such as Pseudo-nitzschia, but not in other Stramenopiles (which include centric diatoms, brown algae and some protist plant parasites) or Cryptophyte relatives. Members of this genus readily bloom upon addition of iron to Fe-limited waters, and are known to produce the neurotoxin domoic acid. Until now, the reason for the success of pennate diatoms in the open ocean was uncertain; however, expressing ferritin would allow pennate species to store Fe after a transient input, using it to dominate Fe stimulated algal blooms. Here, the ferritin gene was cloned from the coastal pennate diatom Pseudo- nitzschia multiseries, overexpressed in Escherichia coli, and purified using liquid chromatography. The ferritin protein sequence appears to encode a non-heme, ferritin- like di-iron carboxylate protein, while gel filtration chromatography and SDS-PAGE indicate that this ferritin is part of the 24 subunit maxi-ferritins. Spectroscopically monitoring the addition of Fe(II) to a buffered ferritin solution shows that the P. multiseries protein demonstrates ferroxidase activity, binding iron and storing it as Fe(III) in excess of 600 equivalents per protein shell. In keeping with the typical stoichiometry of the ferroxidase reaction, oxygen (O2) is consumed in a 2 Fe:O2 ratio while hydrogen peroxide is produced concurrently. iii Diatoms evolved from secondary endosymbiosis involving eukaryotic red algae; however, a broad phylogenetic comparison suggests that P. multiseries ferritin was likely acquired via lateral gene transfer from cyanobacteria – not from its ancestral endosymbionts. Until recently, no other ferritins have been identified in diatoms, and the protein characterized here is unique in that it seems to be derived from a prokaryotic organism yet it occurs in a marine eukaryote. These findings have direct implications for the success of pennate diatoms in both Fe rich coastal waters and upon Fe addition in the open ocean. iv Table of Contents Abstract........................................................................................................................... ii Table of Contents ........................................................................................................... iv List of Tables ................................................................................................................. vi List of Figures...............................................................................................................vii List of Symbols, Abbreviations and Nomenclature .......................................................viii Acknowledgements ......................................................................................................... x CHAPTER ONE: LITERATURE REVIEW AND THESIS OBJECTIVES..................... 1 1.1 General Introduction.............................................................................................. 1 1.2 The Role of Iron in Marine Primary Production..................................................... 3 1.3 Phytoplankton Iron Physiology.............................................................................. 6 1.4 Impact of Iron Availability on Phytoplankton Community Structure...................... 8 1.5 Classification and Molecular Properties of Ferritin Proteins................................. 12 1.5.1 Eukaryotic Ferritins ..................................................................................... 14 1.5.2 Bacterial Ferritins ........................................................................................ 17 1.5.3 Dps Ferritins................................................................................................ 20 1.6 Thesis Objectives ................................................................................................ 21 CHAPTER TWO: MATERIALS AND METHODS ..................................................... 22 2.1 PCR Based Cloning of Pseudo-nitzschia Ferritin ................................................. 22 2.2 Colony Selection and Sequence Verification ....................................................... 28 2.3 Protein Sequence and Phylogenetic Comparison.................................................. 30 2.4 Purification of Recombinant Ferritin.................................................................... 31 2.5 Determination of Protein Size and Purity by Gel Filtration and SDS-PAGE ........ 34 2.6 Protein Determination.......................................................................................... 35 2.7 Spectroscopic Assay of Iron Binding by Apoferritin ............................................ 36 2.8 Determination of Ferritin Iron Content ................................................................ 37 2.9 Oxygen and Hydrogen Peroxide Measurements................................................... 39 CHAPTER THREE: RESULTS.................................................................................... 41 3.1 General Properties of Recombinant Ferritin ......................................................... 41 3.2 Phylogenetic Analysis ......................................................................................... 42 3.3 Protein Purification.............................................................................................. 49 3.4 Spectroscopic Analysis of Ferritin ....................................................................... 51 3.5 Spectroscopic Determination of Ferroxidase Activity .......................................... 52 3.6 Ferritin Iron Content and Binding Capacity ......................................................... 55 3.7 Dissolved Oxygen and Hydrogen Peroxide Measurement .................................... 56 CHAPTER FOUR: DISCUSSION ................................................................................ 58 4.1 Behaviour of Recombinant Ferritin in Solution.................................................... 58 4.2 Spectroscopic Analysis........................................................................................ 59 4.3 Fe Binding Capacity ............................................................................................ 62 4.4 Ferroxidase Fe/O2 and Fe/H2O2 Stoichiometry ..................................................... 65 4.5 Classification of P. multiseries Ferritin ................................................................ 66 4.6 Evolution of Ferritin in the Marine Environment ................................................. 67 vCHAPTER FIVE: CONCLUSIONS.............................................................................. 71 REFERENCES ............................................................................................................. 73 vi List of Tables Table 3-1 Collection of 29 selected ferritin taxa including closest relatives of P. multiseries used for phylogenetic analysis. ............................................................ 43 Table 3-2 Ferene S® determination of Fe bound by ferritin. .......................................... 55 vii List of Figures Figure 1-1 High nutrient, low chlorophyll regions of the surface ocean (100 ˚E – 320 ˚E). ..........................................................................................................................4 Figure 1-2 Pennate diatom Pseudo-nitzschia sp. forming chains. .....................................8 Figure 1-3 Crystal structures of various ferritins............................................................ 13 Figure 2-1 Pseudo-nitzschia ferritin amino acid sequence and DISOPRED2 predicted secondary structure. ............................................................................................... 23 Figure 2-2 PCR products from the amplification (lane 1) and extension (lane 2) of the putative ferritin gene from Pseudo-nitzschia multiseries. ....................................... 24 Figure 2-3 Diagram of ligase independent PCR based cloning of P. multiseries ferritin. .................................................................................................................. 26 Figure 2-4 PCR products from megaprimer insertion (reaction 3) after 18 (lane 1), 21 (lane 2), and 25 cycles (lane 3). ............................................................................. 28 Figure 2-5 Colony PCR to determine the presence or absence of an inserted sequence. . 30 Figure 2-6 Confirmation of induction and heat shock purification of overexpressed ferritin. .................................................................................................................. 32 Figure 2-7 Source 15Q anion exchange purification of recombinant ferritin .................. 33 Figure 2-8 Two-point calibration of O2 electrode (Zero O2 and atmospheric O2)............ 40 Figure 3-1 Selectively Degapped MAFFT amino acid sequence alignment of 29 selected ferritin amino acid sequences ................................................................... 44 Figure 3-2 Unrooted phylogenetic tree of selected ferritin sequences............................. 47 Figure 3-3 Maximum likelihood rooted phylogenetic tree.............................................. 48 Figure 3-4 Denaturing 12% SDS-PAGE of ~1 µg purified ferritin. ................................ 49 Figure 3-5 Gel filtration chromatograph of apoferritin ................................................... 50 Figure 3-6 UV-visible absorbance spectrum of apoferritin............................................. 51 Figure 3-7 Difference absorbance spectra of ferritin titrated with ferrous iron................ 53 Figure 3-8 Change in absorbance at 295 nm versus iron added per ferritin shell. ........... 54 Figure 3-9 Sample oxygen electrode trace showing O2 consumption following ferrous iron addition to apoferritin ..................................................................................... 57 viii List of Symbols, Abbreviations and Nomenclature Symbol Definition ε Molar extinction/absorptivity coefficient ∆A295 Change in absorbance at 295 nm A280 280 nm absorbance AU Absorbance units AXC Anion exchange chromatography BFR Heme-binding 24 subunit bacterioferritin bp Base pairs cDNA Complimentary deoxyribonucleic acid DA Domoic acid DNA Deoxyribonucleic acid dNTPs Deoxyribonucleotide triphosphates Dps DNA binding protein from starved cells dsDNA Double stranded deoxyribonucleic acid EDTA Ethylene-diamine-tetra-acetic acid EST Expressed sequence tag Euk Heme-binding heteropolymer eukaryotic ferritin Fe’ Free iron Fe-Qhigh Maximum Fe requirement at maximum growth rate (Fe storage) Fe-Qlow Minimum Fe requirement at maximum growth rate Fox Ferroxidase FTN Non-heme 24 subunit homopolymer ferritin GF Gel Filtration (chromatography) HNLC High nutrient, low chlorophyll HPLC High performance liquid chromatography IPTG Isopropyl β-D-1-thiogalactopyranoside kb Kilobase kDa Kilodaltons ix MCO Multi-copper oxidase MES 2-N-morpholino-ethanesulfonic acid MW Molecular Weight NCBI RefSeq National Center for Biotechnology Information Reference Sequence NOAA PMEL National Oceanic and Atmospheric Administration Pacific Marine Environmental Laboratory OD600 Optical density at 600 nm absorbance PCR Polymerase chain reaction pH -log[H+] Pi Inorganic phosphate Redox Reduction/oxidation reactions ROS Reactive oxygen species SA:V Surface area to volume ratio SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis TCA Trichloroacetic acid TCEP Tris 2-carboxyethyl phospine US DoE JGI United States Department of Energy Joint Genome Institute UV-vis Ultraviolet-visible Ve Elution volume xAcknowledgements I am infinitely grateful for the continued support of my research supervisor, Dr. Maria T. Maldonado – her passion for biological oceanography is truly inspiring, and her expertise invaluable. With Maite’s guidance I have enjoyed a period of scientific discovery that has improved my observation skills and taught me to think more systematically and to reason more logically. My time in the lab has also shown me the value of patience, yet another key ingredient for success in anything. I acknowledge Michael E.P. Murphy and Beverley R. Green, committee members extraordinaire, for their ongoing support and guidance – it was always helpful and forthcoming. Your office and lab doors were always open to me, an instrumental factor in the completion of this work. Philippe D. Tortell also played a highly positive role in my development as a scientist - his advice was always sound. The commitment that he continually shows in guiding the graduate program is an asset to all graduate students in the department of Earth and Ocean Sciences. I also extend warm regards to all my fellow lab members, past and present, from the Maldonado lab, the Murphy lab and from the Armbrust lab, from whom so many ideas have come. Their friendship and encouragement has positively influenced my personal and academic development. Finally I graciously appreciate my parents, Dr. Rino and Rita Moccia, for their ongoing support, and for teaching me what it takes to get the job done. This thesis is dedicated to them. 1Chapter One: Literature Review and Thesis Objectives 1.1 General Introduction Phytoplankton are unicellular photosynthetic organisms that form the base of most aquatic food webs. This group accounts for nearly half of global inorganic carbon (C) fixation (Falkowski, 1994), using carbon dioxide (CO2), sunlight, and water to produce organic molecules required in cellular metabolism. A significant portion of the fixed organic C in the surface ocean is exported to the deep ocean where it may be sequestered, away from the surface, for long periods of time (Falkowski et al., 1998). Since the dissolved gases in the surface ocean are in equilibrium with the atmosphere, increased sequestration of organic C, termed the ‘biological pump’ (Volk and Hoffert, 1985), leads to a reduction in atmospheric CO2, a greenhouse gas projected to rise substantially in concentration in the near future due to anthropogenic inputs (IPCC, 2001b). Currently about one third of anthropogenic CO2 is entering the surface ocean, primarily in the Southern Ocean (Orr et al., 2001). Both the biological pump and its chemical/physical counterpart, the ‘solubility pump’ of dissolved inorganic carbon, dictate the extent to which CO2 is removed from the atmosphere (Raven and Falkowski, 1999; Falkowski et al., 2000); but ultimately this removal is suboptimal due to iron (Fe) limitation of primary production in vast areas of the open ocean. Under favourable conditions, when the pressure of nutrient limitation or zooplankton grazing is alleviated, algal blooms have the potential to occur. Fast growing diatoms, in particular, play a key role in the biological pump, having been suggested to dominate marine primary production  (Falkowski et al., 1998). In contrast to most other microalgae, diatoms are contained within a dense silicate frustule, and exhibit more rapid 2sinking rates leading to increased export production (Buesseler, 1998). Thus, algal blooms that are high in diatom biomass are major components of the ocean’s biological C sink. However, processes that interrupt the downward flux of C such as zooplankton grazing (Boyd et al., 2004) or viral lysis (Suttle, 2007) in the euphotic zone, and bacterial remineralization throughout the water column (Azam, 1998) may limit C sequestration into the deep ocean. Surface waters are generally low in the macronutrients nitrogen (N), phosphorus (P) and silicon (Si) due to their utilization by phytoplankton; these macronutrients limit algal growth in major oligotrophic waters of the ocean. However, there are significantly large areas of the ocean where these macronutrients are in high abundance, but where primary production is very low. Following the first shipboard iron (Fe) addition experiments (Martin and Fitzwater, 1988), John Martin (1990) postulated that low Fe concentrations were limiting phytoplankton biomass in what are referred to as “high nutrient, low chlorophyll” or HNLC regions. Since then, a series of in situ Fe fertilization experiments have contributed to a compelling body of work supporting the iron hypothesis (e.g. Martin et al., 1994; Coale et al., 1996; Boyd et al., 2000; Gervaise et al., 2002; Tsuda et al., 2003; and Coale et al., 2004). In these studies, Fe addition boosted primary productivity and algal biomass, increased photosynthetic efficiency, and favoured larger algal cells such as diatoms (Martin et al., 1994; Boyd et al., 2001). At the height of such Fe stimulated algal blooms, two pennate diatom genera, Pseudo-nitzschia and Fragilariopsis, have been repeatedly shown to dominate assemblages (Landry et al., 2000; Gall et al., 2001; Gervais et al., 2002; Coale et al., 2004; Marchetti et al., 2006b). 3The question arises, what promotes the success of the pennate diatoms during sporadic influx of Fe in HNLC areas? Recent findings indicate that a gene encoding ferritin, an Fe storage and detoxification protein, is found in several species of pennate diatoms, potentially providing a selective advantage to these diatoms through improved Fe retention after a transient pulse (Marchetti et al., in prep). Preliminary sequence alignments indicated that ferritin of the coastal diatom Pseudo-nitzschia multiseries is somewhat unique from previously characterized ferritins, a ubiquitous but structurally diverse class of proteins. The main objective of this thesis was to clone the ferritin gene from P. multiseries, and to generate a recombinant protein construct to facilitate its characterization. Ultimately, the goal was to employ the purified protein as a suitable standard to be used in HPLC analysis, in the hopes of confirming the presence and abundance of ferritin in cultures of a coastal diatom, P. multiseries, and an open ocean isolate, Pseudo-nitzschia granii, grown under Fe limited and Fe replete conditions. Results of the latter objective could support pre-existing mRNA expression data, and help address the hypothesis that the presence and upregulation of ferritin expression in these pennate diatoms is responsible for their dominance during Fe enrichment experiments. 1.2 The Role of Iron in Marine Primary Production From in situ Fe enrichment experiments, it is now known that HNLC regions (Fig. 1-1) comprise ~30% of the global surface ocean including the North East Pacific (Boyd et al., 2004), Western subarctic Pacific (Tsuda et al., 2003), the Southern Ocean (Boyd et al., 2000) and parts of the Equatorial Pacific (Martin et al., 1994). In these 4regions, surface macronutrients are abundant and could support a dense community of photosynthetic organisms, but phytoplankton biomass is much lower than expected (Falkowski, 1995). As a result, the lack of primary production leads to the accumulation of N, P, and Si. Early research determined that Fe, although required in trace amounts by photosynthetic organisms, may be too low in surface seawater to support growth (Brand et al., 1983; Martin and Fitzwater, 1988; Morel and Price, 2003). Thus, in examining the HNLC hypothesis, it is important to understand the biogeochemistry of Fe in the ocean that controls its supply and availability to phytoplankton.          Figure 1-1 High nutrient, low chlorophyll regions of the surface ocean (100 ˚E – 320 ˚E). The left chart represents areas of sub maximal chlorophyll a levels, and the right chart indicates areas where surface nitrate has not been completely used. Ovals indicate the North East Pacific, parts of the equatorial Pacific, and the entire Southern Ocean (NOAA PMEL). 5HNLC areas experience sporadic aeolian (windblown dust) inputs from land that are up to three orders of magnitude lower than those in coastal ecosystems (Duce and Tindale, 1991; Wu and Luther, 1996). Other sources of Fe to the surface ocean include river runoff, precipitation, upwelling, eddy diffusivity and anthropogenic inputs; however, it is the Fe-rich dust input that is thought to play the most important role in increased export production during glacial periods (Petit et al., 2000; Bopp et al., 2003). Other models indicate that the upwelling of recycled Fe could account for the majority of open ocean primary production, but suggest that on short temporal scales it is the high reactivity and low residence time of Fe combined with the presence of organic ligands that ultimately control its availability to phytoplankton (Johnson et al., 1997; Archer and Johnson, 2000). In the open ocean, roughly 90% of Fe forms insoluble particulates sequestered in living organisms, adhering to sinking detritus, or forming macromolecular organic colloids (Price and Morel, 1998; Nishioka and Takeda, 2000). Of the dissolved Fe pool, up to 99.9% is complexed by organic ligands (Rue and Bruland, 1995); these Fe complexes are not directly available for phytoplankton growth (Maldonado and Price, 2001). The net effect of these additive factors is subnanomolar concentrations of dissolved inorganic iron (Fe’) (Rue and Bruland, 1995; Wu and Luther, 1995) such that bioavailable Fe in the ocean is a sparse commodity, for which competition among microorganisms likely fuels the development of physiological adaptations to reduce cellular Fe demands, to acquire it more effectively, and/or to store surplus Fe when possible. 61.3 Phytoplankton Iron Physiology Phytoplankton require Fe for nitrogen uptake and utilization, the biosynthesis of chlorophyll, and in oceanic diatoms, it is utilized most heavily in the electron transport proteins of photosynthesis (Raven et al., 1999). To meet cellular Fe demand, oceanic phytoplankton employ several key adaptations that allow them to grow, albeit more slowly, in the Fe starved waters of the open ocean. For some time it has been known that oceanic species demonstrate greatly reduced Fe requirements compared with their coastal counterparts, and are able to sustain higher growth rates under Fe limiting growth conditions (Brand, 1991; Sunda and Huntsman, 1995; Maldonado and Price, 1996). In general, the reduction of cell size under Fe limitation helps to ameliorate nutrient acquisition by increasing the surface area to volume ratio (SA:V) of the cell (Brand et al., 1983), but the high iron use efficiency of oceanic diatoms (Maldonado and Price, 1996) is derived from more specialized methods to cope with Fe deficiency. Plasticity in metalloprotein utilization may help to conserve photosynthetic iron requirements in a number of ways. In the pennate diatom Phaeodactylum tricornutum, for example, the Fe-containing protein ferredoxin may be replaced with the Fe-free variant flavodoxin (containing manganese) during periods of low Fe stress (Entsch, 1983; La Roche et al., 1995). There are also instances where metalloproteins containing alternative transition metals such as copper (Cu), manganese (Mn), and zinc (Zn) can be used in place of their Fe-containing counterparts in various marine phytoplankton including green algal and diatom genera (Raven et al., 1999). A recent example in the small centric oceanic diatom Thalassiosira oceanica describes the substitution of the Fe-containing electron transport protein cytochrome c6 with Cu-containing plastocyanin (Peers and 7Price, 2006). More drastic alterations to the photosynthetic architecture are also possible; under Fe-limitation, T. oceanica also shows markedly reduced relative quantities of Fe- rich photosystem I and cytochrome b6f complexes compared to its coastal counterparts (Strzepek and Harrison, 2004). In the latter case, T. oceanica reduces the demand for Fe without significantly affecting its photosynthetic rate. Diatoms are also well equipped to endure Fe limitation in that they are able to access the vast pool of Fe that is complexed with organic ligands such as siderophores (Maldonado and Price, 1999, 2001), which are produced by marine bacteria to scavenge Fe when it is in low abundance (Granger and Price, 1999). The coccolithophorid Emiliania huxleyi is known to produce organic ligands of its own, although elevated ligand concentrations have been noted upon Fe addition rather than depletion (Rue and Bruland, 1997; Boye and van den Berg, 2000). T. oceanica is able to capitalize on the presence of these organically bound iron by possessing an inducible, high affinity Fe uptake system that allows the reduction and subsequent internalization of Fe, albeit slowly, from organic ligands (Maldonado and Price, 2001). Further evidence for this suggests that the diatom employs a multi-Cu oxidase (MCO) dependent Fe uptake system to obtain organically bound Fe (Maldonado et al., 2006). This is in contrast to the conventional model whereby phytoplankton were thought to be dependent on the small pool of unchelated inorganic iron (Fe’) present in the marine environment (Hudson and Morel, 1990; Sunda and Huntsman, 1995). The ability of diatoms to efficiently derive Fe from the marine environment is likely augmented by the upregulation of the number of Fe uptake transporters at the cell surface (Kustka et al., 2007). 8Despite these adaptations, and their large impact on global carbon sequestration via the biological pump, the small oceanic phytoplankton that characterize low productivity HNLC waters do not dominate bloom assemblages when Fe is added. Instead, in situ Fe fertilization experiments result in dramatic community structure shifts with a bias towards larger diatom cells, in particular pennate diatoms. 1.4 Impact of Iron Availability on Phytoplankton Community Structure When pulses of Fe are introduced to HNLC regions, the community structure changes markedly. Pseudo-nitzschia (class Bacillariophyceae, Fig. 1-2) is one such genus that almost universally responds to the Fe influx, and numerically dominates the resulting phytoplankton blooms, despite a characteristically low abundance in HNLC regimes (Landry et al., 2000; Gall et al., 2001; Gervais et al., 2002; Coale et al., 2004; Marchetti et al. 2006b). At ocean station Papa (50 ˚N, 145 ˚W), a frequently sampled HNLC locale, small pennates are generally present in higher abundance than centric diatoms (Horner and Booth, 1990), despite observations by Ishizaka et al. (1986) that pennate diatoms sink faster than centrics, especially when this effect is magnified by nutrient deprivation (Bienfang et al., 1982). Figure 1-2 Pennate diatom Pseudo-nitzschia sp. forming chains. (Rennie, 2008) 9 Pseudo-nitzschia is widely distributed in the oceans, and includes nearly 30 species (Lundholm et al., 2002; Hasle, 2002). The cells are several µm in width and often exceed 100 µm in length, making them just visible to the naked eye in culture. These pennate diatoms are encased in a weakly silicified frustule, with striated girdle bands, and an extremely eccentric raphe (a thin slit that runs along the cell centreline). They also demonstrate colony formation by overlapping each other in a stepwise fashion at the cell ends (Hasle, 1994) (Fig. 1-2). The majority of research on members of this genus involves coastal species that produce domoic acid during harmful algal blooms. Domoic acid is a neurotoxin implicated in amnesic shellfish poisoning and can cause digestive and respiratory difficulty, permanent shor-term memory loss, confusion, nausea and in severe cases even death in humans (Bates, 2004). Recent studies indicate that coastal strains may secrete domoic acid for use as an extra-cellular Fe and Cu chelator (Rue and Bruland, 2001) such that the uptake of Fe is enhanced (Maldonado et al., 2002). Although this effect has not been confirmed in HNLC isolates, when Fe is limiting, DA is known to improve the uptake of copper, which is used in the high affinity Fe uptake system of diatoms, e.g. MCO (Wells et al., 2005). In eutrophic marine environments such as the coast or areas of upwelling where Fe limitation is less common, large individual and chain forming diatoms dominate the phytoplankton community. This success is mainly attributed to the fact that these fast growing cells are able to store macronutrients in central vacuoles (Stolte and Riegman, 1995), giving them a competitive advantage by depriving other phytoplankton of access to those nutrients (Raven, 1997; Tozzi et al., 2004). Conversely, when surface Fe 10 availability is low, it is clear that small pennate and centric diatoms and picoplankton comprise the majority of the open ocean assemblage (DiTullio et al., 1993). While diatoms in general may exhibit high efficiency acquisition and utilization of subnanomolar Fe supplies, the smallest centric diatoms demonstrate faster nutrient uptake rates per cell volume (Sunda and Huntsman, 1995) and it might stand to reason that they could easily out-compete their larger pennate counterparts with increased nutrient demands. However, the pennate diatoms also have an advantage in that their elongated shape and tendency to form chains increases SA:V and improves the rate of nutrient uptake (Pahlow et al., 1997). In fact, when the uptake rates determined by Sunda and Huntsman (1995) were normalized to SA, the calculated rates were independent of algal cell size. Thus, it is difficult to predict the success of either morphology based on size and shape alone; it is necessary to compare their biochemical differences as well. It is unlikely that improved SA:V or the use of DA as an extracellular metal ligand alone would allow such widespread success of pennate diatoms over their centric counterparts upon relief of Fe limitation. A recent study has shown that the minimum Fe requirement at maximum growth rate (Fe-Qlow) for Fe-limited oceanic Pseudo-nitzschia sp. is comparable to those of the small centric T. oceanica, but that the Fe storage (Fe- Qhigh) of the oceanic pennate species is comparatively higher and more variable when Fe is abundant (Marchetti et al., 2006a). This added plasticity may thus reflect a potential luxury Fe uptake system in members of this genus, whereby Fe is taken up when it is available during enrichment events, then retained in the cell for use long after external Fe levels have been depleted (Marchetti et al., 2006a). 11 Despite its utility within the cell, Fe does not exist freely at high concentrations in the cytoplasm due to its high reactivity, and insolubility in an oxidizing environment at physiological pH. While the electrochemical properties of this transition metal make it ideal for electron transport, free intracellular iron can lead to redox reactions that generate reactive oxygen species (ROS), such as free radicals and peroxides that lead to oxidative stress. Complexes incorporating Fe(III), the favoured oxidation state of iron at neutral pH, are in equilibrium with free Fe(III), an extremely reactive Fe species. Free Fe3+ reacts readily with the superoxide radical (O2•-), thereby being rapidly reduced to Fe(II) (Eqn 1). This species can react with hydrogen peroxide (H2O2) in the Fenton reaction (Eqn 2), generating highly reactive hydroxyl radicals. Fe3+ + O2•-  Fe2+ + O2 [1] Fe2+ + H2O2  Fe3+ + OH- + OH• [2] O2•- + H2O2  O2 + OH- + OH• [3] Equation 3 is the overall Fe catalyzed Haber-Weiss reaction, the products of which lead to the degradation of deoxyribonucleic acid (DNA) and other biomolecules (Haber and Weiss, 1934; Halliwell and Gutteridge, 1984). The Fenton reaction can also be generated with Fe(II) as a starting material (Chiancone, et al., 2004) in the following series of reactions, although in this case the overall reaction (Eqn 3b) includes the net oxidation of Fe(II):    Fe2+ + O2  Fe3+ + O2•- [1b]  2O2•- + 2H+  H2O2 + O2 [1c] Fe2+ + H2O2  Fe3+ + OH- + OH• [2] 2Fe2+ + 2H+ + 2O2•-  2Fe3+ + OH- + OH• [3b] 12 Thus, any organism that stores surplus Fe must do so by sequestering it safely away from ROS, yet in a readily accessible form. In plants, mammals and many microorganisms, proteins of the nearly ubiquitous yet structurally diverse family of ferritins perform intracellular Fe detoxification and storage. All ferritins are involved in Fe homeostasis, and the absence of the protein is potentially lethal in higher organisms (Liu and Theil, 2004). During a recent analysis of Pseudo-nitzschia spp., an expressed sequence tag (EST) encoding a ferritin-like protein was identified, and transcript levels of this gene were found to increase two-fold upon iron addition to Fe limited cultures (Marchetti et al., in prep). In a recent comprehensive survey of algal taxa including 23 diatom species (Marchetti et al., in prep), the pennate diatoms Phaeodactylum tricornutum (US DoE JGI, 2006), Fragilariopsis spp. and Pseudo-nitzschia spp., were found to possess putative ferritin genes, as was the red algal Cyanidioschyzon merolae (Matsuzaki et al., 2004). Interestingly, however, no ferritin gene sequences were found bioinformatically or by PCR-based methods in the genome of Thalassiosira pseudonana (Armbrust et al., 2004), or any other centric diatom examined. These findings suggest that pennate diatoms have followed a unique evolutionary trajectory, and may derive their success during Fe enrichment from the ability to express the iron-storing protein ferritin. 1.5 Classification and Molecular Properties of Ferritin Proteins There are three main groups of ferritin proteins discussed in the literature: eukaryotic ferritins (ie from mammals and plants), bacterial ferritins (including heme- binding and non-heme subgroupings), and Dps (DNA binding protein from starved cells) 13 ferritins. The first two are considered to be maxi-ferritins, with nanocages constructed of 24 subunits, while the smaller Dps class are mini-ferritins, comprised of only 12 subunits. Ferritins are part of a broad group of di-iron carboxylate proteins that bind and oxidize Fe(II) (Levi et al., 1988; Lawson et al., 1989; Nordlund and Eklund, 1995; Treffry et al., 1997), and store many atoms of Fe(III) within a ferrihydrite mineral core with the rough stoichiometry 9Fe2O3•5H2O (Towe, 1981). Maxi-ferritins commonly have a MW of 450 kDa, are roughly spherical, and measure 12 nm in diameter with an 8 nm hollow core when empty (Harrison and Arosio, 1996); mini-ferritins are comparatively smaller, around 240 kDa (Bozzi et al., 1997). Extensive inter and intrasubunit interactions make ferritins heat stable up to ~80˚C (Liu and Theil, 2004). While it is clear that there is great diversity among members of this superfamily of proteins, the differences can be intricate. Representative crystal structures of the three main types of ferritins from the Protein Data Bank (PDB, Berman et al., 2000) are shown below (Fig. 1-3). A         B         C Figure 1-3 Crystal structures of various ferritins (not to scale). Human mitochondrial ferritin (3A, PDB 1R03), Non-heme ferritin (FTN) from Thermotoga maritima (3B, PDB 1VLG), and Dps ferritin from Escherichia coli (3C, PDB 1DPS) (Berman et al., 2000). 14 1.5.1 Eukaryotic Ferritins Mammalian ferritins (Fig. 1-3A), most commonly isolated from human and horse, are composed of two types of subunits, H (MW ≈ 21 kDa) and L (MW ≈ 19.5 kDa) for heavy and light, or heart and liver, respectively (Orino et al., 1997). The subunits are arranged in various proportions of the two, depending on the tissue from which they were isolated, and the fully formed structure of ~450 kDa can bind up to 4500 Fe atoms (Arosio et al. 1978; Harrison and Arosio, 1996; Orino et al., 1997). While 450 kDa is the commonly reported MW of animal ferritins, larger ones have also been described, for example the 530 kDa ferritin of the chiton Aeanthopteura hirtosa (Kim et al., 1986). In vitro, mammalian ferritins are able to bind approximately 12 heme groups, but unlike heme-containing bacterioferritins, these proteins are normally purified in a heme-free form (Kadir et al., 1992). The characteristic ferroxidase (Fox) activity of ferritins has been linked to the H- chain, which is evolutionarily related to the subunit type that is present in plant and bacterial ferritin homopolymers (Andrews et al., 1991), while the L chain is better suited to Fe core nucleation and release (Levi et al., 1988; Levi et al., 1992; Lawson et al., 1989; Baaghil et al., 2003). Within the H and L groupings, sequence identity is well conserved, around 80% on average, but between H and L chains there is closer to 50% amino acid identity. Most of this similarity between the subunits is associated with the residues that form the 8 hydrophilic channels through which Fe(II) is thought to enter the protein cavity in eukaryotic ferritins (Boyd et al., 1985; Hempstead et al., 1997). The mechanism by which mammalian H-chain ferritin binds Fe(II) has been determined (Yang et al., 1998), and is described by the following reaction series (for 15 simplicity the overall charges of the protein complexes are not included, as they have no impact on overall stoichiometry): Diferrous binding (phase 1):       2Fe2+ + P  [Fe(II)2-P] [4] Ferroxidation (phase 2):        [Fe(II)2-P] + O2 + 3H2O  [Fe(III)2O(OH)2-P] + H2O2 + 2H+ [5] Core formation (phase 3):    [Fe(III)2O(OH)2-P] + H2O  2FeO(OH)(core) + P + 2H+ [6] Overall Reaction:    2Fe2+ + O2 + 4H2O  2FeO(OH)(core) + H2O2 + 4H+ [7] The overall reaction (Eqn 7) is the sum of 3 phases: the fast pairwise binding of 2 Fe2+ to a di-nuclear Fox site of the apoprotein (P) (phase 1, Eqn 4), the slower oxidation to Fe3+ by O2 producing H2O2 (phase 2, Eqn 5), and finally iron hydrolysis where Fe3+ is incorporated into the ferrihydrite mineral core (phase 3, Eqn 6). After sufficient core formation has occurred, auto-oxidation of Fe(II) directly onto the mineral core is described by the following reaction:     4Fe2+ + O2 + 6H2O  4FeO(OH)(core) + 8H+ [8] 16 Higher plant ferritins (phytoferritins) are functionally similar to animal ferritins, sharing up to 49% amino acid sequence identity and likely a common ancestry (Andrews et al., 1992; Harrison and Arosio, 1996). Like their mammalian counterparts, pea, maize, and soybean (Glycine sp.) ferritins are also comprised of two different subunits, with the larger of two around 26 - 28 kDa, and an assembled weight of the 24-mer up to 540 and 570 kDa (Ko et al., 1987; Laulhère et al., 1988). Plant ferritin genes are typically located in the nucleus, and include an N-terminal signal and transit peptide that directs the functional gene product to the chloroplast (Ragland et al., 1990). Since non-functional, vestigial remnants of bacterial ferritin nucleotide sequences are noted in some chloroplast genomes, it has been suggested that plant ferritin genes were initially derived through endosymbiosis, but were later replaced by a different gene within the nucleus (Ragland et al., 1990; Briat et al., 1999). A dual benefit of maintaning genes nucleus versus the chlroplast is to protect the genetic material from free radical damage in the oxygen (O2) and Fe rich plastid, and to propagate them where fitness improving recombination is possible (Martin, 2003). 17 1.5.2 Bacterial Ferritins Bacterial ferritins (Fig. 1-3B) are homopolymer ferritins, comprised of 24 identical H type subunits (~18.5 kDa each, 450 kDa for the assembled structure) and contain both oxidation and nucleation sites that lead to the storage of up to 2000 Fe (Andrews, 1998). They can be further subdivided into two groups: bacterioferritins (BFR) that binds heme groups between subunits, and non-heme ferritins (FTN) (Harrison and Arosio, 1996). According to the NCBI RefSeq non-redundant protein database (Pruitt et al., 2007), species that express BFR include Rhodobacter capsulatus, Neisseria gonorrheae, Pseudomonas aeruginosa and Synechocystis sp., while Thermotoga sp., Helicobacter pylori, Campylobacter jejuni, Mycobacterium tuberculosis, and a number of archaeal genera, such as Candidatus methanoregula and Pyrococcus furiosus, belong to the FTN subgroup. Escherichia coli expresses both types of maxi-ferritins, heme BFR and non-heme FTN. Proteins belonging to these two classes are indistinguishable with respect to the rate at which they bind iron, but heme-ferritins accumulate less iron overall, and it is thought that the heme promotes the release of Fe from the protein (Andrews et al., 1995; Baaghil et al., 2003). While the Fox sites tend to be highly conserved, bacterial ferritins share low sequence identity with mammalian ferritins (up to 23% to the H-chain, and 21% to the L chain), and the two types of E. coli ferritin share only 14% identity with each other (Andrews et al., 1991; Andrews et al., 1992). Structurally, however, bacterial ferritins and clonal mammalian ferritins (containing only H-chains) are somewhat similar, in that their α carbons can be superimposed to within 0.2 nm (Arosio and Levi, 2002). 18 So far, only the reaction stoichiometry of heme-containing BFR has been proposed (Yang et al., 2000a; Bou-Abdallah et al., 2002) and is shown in the following series of reactions: Diferrous binding (phase 1):        2Fe2+ + P  [Fe(II)2-P] + 4H+ [9] Ferroxidation (phase 2): Step one,        [Fe(II)2-P] +O2 + H2O  [Fe(III)2O-P] + H2O2 [10] Step two,      [Fe(II)2-P] + H2O2  [Fe(III)2O-P] + H2O [11] Net reaction,        2[Fe(II)2-P] + O2  2[Fe(III)2O-P] [12] Core Formation (phase 3): Step one,    2Fe2+ + O2 + 4 H2O  2FeO(OH)(core) + H2O2 + 4H+ [13] Step two,        2Fe2+ + H2O2 + 2 H2O  2FeO(OH)(core)  + 4H+ [14] Net reaction,     4Fe2+ + O2 + 6H2O  4FeO(OH)(core) + 8H+ [15] The stepwise reaction mechanism involves oxidation at two separate di-nuclear Fox sites, and was proposed by Yang et al. (2000a) to explain the complete conversion of O2 to water that was observed during the addition of Fe(II) to BFR. They suggest that production of hydrogen peroxide during ferroxidation can then be used as an intermediate oxidant during a second proximal diferrous binding event, thereby avoiding the generation of reactive hydrogen peroxide. Thus, during ferroxidation the net 19 stoichiometry in equation 12 appears to involve a 4:1 ratio of Fe(II):O2; in actuality, 2 Fe(II) are being oxidized by one molecule of O2, while another 2 Fe(II) are oxidized by the short-lived H2O2 intermediate. In contrast to E. coli BFR, non-heme EcFTN was found to contain a third iron oxidation site, separate from the di-nuclear Fox site (Hempstead et al., 1994; Treffry et al., 1998). In vitro Fe(II) titration showed that O2 consumption decreased markedly after 48 Fe(II)/EcFTN were added (saturating the 24 ferroxidase sites), and that once sufficient core formation had occurred, further Fe(II) was oxidized and added directly to the core, the Fox sites having been superseded (Treffry et al., 1997; Treffry et al., 1998). This shift in core formation was accompanied by a change in Fe oxidation stoichiometry as well, from 2 Fe(II):O2 (as observed for mammalian ferritins in equation 7) to a 4:1 Fe(II):O2 ratio (as per equation 8, auto-oxidation in mammalian ferritins, and 15, net core formation in BFR) (Treffry et al., 1998; Yang et al., 2000a). These data suggest that although the lack of heme in EcFTN seems to make it unique from both mammalian ferritins and BFR, its ferroxidase and core formation mechanism is more similar to the mammalian type than to BFR, which demonstrates its own unique Fe oxidation stoichiometry. 20 1.5.3 Dps Ferritins Dps ferritins (DNA binding proteins from starved cells, Fig. 1-3C) are named after their isolation from bacterial culture under conditions of oxidative or nutritional stress (Almirón et al., 1992; Bozzi et a., 1997). Species that produce Dps ferritins upon starvation include E. coli, the cyanobacterium Trichodesmium sp., the bacterium Listeria innocua, as well as archaeal and crenarchaeal members (Almirón et al., 1992; Bozzi et al., 1997; Yang et al., 2000b). In addition to the ability to bind up to 500 Fe (Bozzi et al., 1997), Dps ferritins are thought to function in DNA protection, preventing ROS from damaging the genetic material of the cell (Almirón et al., 2002). This class of ferritin is unique in that its members are comprised of only 12 subunits (18 kDa per subunit and around 240 kDa when assembled), which comprise a homo-dodecamer. Although incorporating only one type of subunit as in the bacterial ferritins, their chemical behaviour is quite different (Bozzi et al., 1997). Dps ferritins generally utilize H2O2 more efficiently than O2 as an oxidant, simultaneously removing both Fe(II) and H2O2, (both components of the Fenton reaction (Eqn 2) that lead to damaging free radical formation) (Zhao et al., 2002; Su et al., 2005). The proposed overall equation for Fe hydrolysis in Dps ferritins is shown in equation 16 below (Wiedenheft et al., 2005): 2Fe2+ + H2O2 + 2H2O 2FeO(OH)(core) + 4H+ [16] Ferritin from L. innocua is an exception to the chemistry of other Dps ferritins, as it binds Fe like BFR and mineralizes Fe as in mammalian auto-oxidation, but does not produce any net H2O2 (Wiedenheft et al., 2005). Like BFR, H2O2 produced at one Fox site might be consumed at a second site, and does not enter into net equations 9, 12, or 15. 21 1.6 Thesis Objectives The body of literature summarized here serves as a framework to characterize new ferritins. Of central interest to this thesis is the newly cloned ferritin protein of the pennate diatom Pseudo-nitzschia multiseries (strain CLN-47). By obtaining a stable recombinant protein construct, the protein’s ferritin-like properties (size, Fe binding characteristics, O2 and H2O2 reactivity) could be identified. Alignment of the protein sequence with those of other known ferritins also allowed for a phylogenetic classification of P. multiseries ferritin. The aims of this thesis were to: a) utilize new cDNA sequence data from Pseudo-nitzschia multiseries (CLN-47) to design primers to amplify the putative ferritin gene, using a PCR based, ligase independent methodology to generate a recombinant ferritin expression construct using the pET28a expression system. b) transform Escherichia coli with the pET28a plasmid construct, to induce the cells to over-express the cloned ferritin protein, and purify the recombinant protein in its spontaneously assembled form. c) determine the ferritin-like properties of the purified protein, in order to classify this ferritin using the available literature as a framework. d) use the purified protein as an effective protein standard with which to develop a purification scheme for native ferritin from algal culture, and to develop a polyclonal antibody for use in future studies. 22 Chapter Two: Materials and Methods 2.1 PCR Based Cloning of Pseudo-nitzschia Ferritin After discovering a putative ferritin gene obtained from an EST (expressed sequence tag) library from Pseudo-nitzschia australis (03184-6D), the full cDNA sequence encoding a ferritin-like protein in this species, as well as in P. multiseries (CLN-47), was determined (Marchetti et al., in prep). Using SignalP 3.0 online prediction software, it was confirmed that the first 31 amino acids of the translated protein sequence comprise a signal peptide (Nielsen and Krogh, 1998; Bendtsen et al., 2004), and the next 36 amino acids represent a plastid targeting peptide (Nielsen et al., 1997; Emanuelsson et al., 2000). Prediction of α-helices in the rest of the protein was performed using DISOPRED2 (Ward et al., 2004) (Fig. 2-1). Genomic DNA at an approximate concentration of 8 µg/µL was extracted from cultured cells of P. multiseries with the DNeasy Plant Mini Kit (Qiagen, Valencia, CA, USA), and the gene encoding the ferritin monomer was amplified from a plasmid containing P. multiseries cDNA, using the forward and reverse primer sequences 5’ GA AGG AGA TAT ACC ATG GGC TCT GAA GAA TTG TTA GAT TTG TTC AAC AGG CAG 3’ and 5' CA AGC TTG TCG ACG GAG TTA AGA ACG GAA CAG ACA TGG ACC AAG AAA AC 3', respectively. These hybrid primers allowed the first five full helices of the ferritin gene to be inserted, without its signal peptide or plastid targeting sequences into a pET28a plasmid (Novagen®, Madison, WI, USA). The portion of the gene used to generate the recombinant ferritin construct lies between the black arrows in Figure 2-1. 23 Figure 2-1 Pseudo-nitzschia ferritin amino acid sequence and DISOPRED2 predicted secondary structure. The 6 α-helices are shown as coloured bars and black lines represent coil structure. Conserved residues of the ferroxidase site are highlighted in pink. Black arrows represent the portion of P. multiseries ferritin that was PCR amplified and inserted into the pET28a vector. STOP indicates the terminus of the natural peptide. As a PCR based method was utilized to insert the gene into the plasmid, the first 20 bp of the forward primer and 17 bp of the reverse primer are complementary to pET28a. By redesigning only the portion of these primers that is complimentary to the gene of interest, they may be used to clone virtually any gene into pET28a. In PCR reaction 1, 200 µM dNTPs and 0.4 µM of the forward and reverse primers were combined with 10 ng of genomic DNA, and the 541 bp megaprimer product (containing the ferritin gene with flanking sequences complimentary to the plasmid multiple cloning STOP 24 site) in Figure 2-2 was generated using the rapid, high fidelity DNA polymerase Phusion™ (New England Biolabs, Ipswitch, MA, USA). Amplification was performed using an initial heat denaturation step (98 ˚C for 30 s), 25 cycles of denaturation, annealing and extension (98 ˚C for 10 s, 60 ˚C for 20 s, 72 ˚C for 15 s), and a final elongation step (72 ˚C for 5 min). The PCR product was purified using Wizard® SV centrifugal cleaning devices (Promega Corporation, Madison, WI, USA) Figure 2-2 PCR products from the amplification (lane 1) and extension (lane 2) of the putative ferritin gene from Pseudo-nitzschia multiseries genomic DNA, and 1 kb plus DNA marker (Invitrogen Corporation, Carlsbad, California, USA). 2% agarose gel stained with ethidium bromide. 100 bp 200 bp 300 bp 400 bp 500 bp 650 bp 850 bp 1000 bp 1650 bp 2000 bp 1        2 25 To facilitate annealing to the plasmid in the third and final ligase-free PCR reaction, 60 bp primers complimentary to pET28a and to the ends of the megaprimer were designed, such that the 541 bp megaprimer was further extended by 40 bp at the beginning and 43 bp at the end. This extending PCR, reaction 2, employed the same conditions as in reaction 1, yielding a single 624 bp dsDNA product that was visualized by agarose gel electrophoresis (Fig. 2-2). The forward extension primer sequence for this reaction was 5’ ATA ACA ATT CCC CTC TAG AAA TAA TTT TGT TTA ACT TTA AGA AGG AGA TAT ACC ATG GGC 3’ and the reverse was 5’ GCC GGA TCT CAG TGG TGG TGG TGG TGG TGC TCG AGT GCG GCC GCA AGC TTG TCG ACG GAG 3’. To insert the ferritin sequence, a modified, directional ligase-independent method was employed (Aslanidis et al., 1990; Zhou and Hatahet, 1995; Tillett and Neilan, 1999). The pET28a plasmid was first digested with EcoR1 (New England Biolabs) for one hour at 37 ˚C to linearize the plasmid, and then heated to 65 ˚C for 20 min to inactivate the enzyme. Using 10 µL PCR reactions containing 15 ng of cut plasmid, 60-90 ng of extended megaprimer, 200 µM dNTPs, and Platinum Pfx DNA polymerase (Invitrogen Corporation, Carlsbad, CA, USA), the plasmid with ferritin inserted was recircularized. The program included an initial heat denaturation step (95 ˚C for 45 s) and a varied number of cycles of denaturation, annealing and extension (95 ˚C for 30 s, 68 ˚C for 30 s, 68 ˚C for 7 min, respectively). A conceptual description of the 3 cloning steps is shown in figure 2-3. 26 Figure 2-3 Diagram of ligase independent PCR based cloning of P. multiseries ferritin (not to scale). Amplification (step 1) and extension (step 2) of the putative ferritin gene from P. multiseries genomic DNA (thick black strands) using hybrid primers that allow the insertion of the megaprimer (step 3) into the pET28a plasmid vector (thin black strands). Orange regions of the hybrid primers are complimentary to the pET28a plasmid, and red regions are complimentary to the ends of the ferritin gene. PCR Step Two PCR Step One P. multiseries Genomic DNA Ferritin Megaprimer PCR Step Three pET28a Plasmid With Ferritin Insert 27 Upon visualization with agarose gel electrophoresis, it was determined that the greatest yield was generally achieved after at least 25 cycles (Fig. 2-4). With increasing cycle number, the amount of megaprimer decreases while the recircularized nicked plasmid band intensifies. The largest band in Figure 2-4 represents supercoiled plasmid, and corresponds to an increased number of colony forming units; however, visualization of this band is not required to generate successful transformants. The PCR product from this final reaction was treated with Dpn-1 (New England Biolabs) for 1 hour at 37 ˚C to degrade any remaining un-amplified methylated plasmid DNA, heated to 65 ˚C for 20 min to inactivate the enzyme, and dialyzed on a nitrocellulose filter disc against 10% PEG 6000 for 30 minutes. 1 uL of the remaining sample was transformed by electroporation into E. coli DH5α to retain and propagate the plasmid. Transformants were incubated in 500 µL of Luria Bertani (LB) medium at 37 ˚C for one hour, and 10 µL were plated on LB agar containing 20 µg mL-1 kanamycin and incubated overnight at 37 ˚C. 28 Figure 2-4 PCR products from megaprimer insertion (reaction 3) after 18 (lane 1), 21 (lane 2), and 25 cycles (lane 3). Small band represents the megaprimer that is consumed as it is inserted into the cut plasmid. The ~6.5 kb band represents nicked recircularized plasmid with insert. The large ~12 kb band represents supercoiled plasmid. 1% agarose gel stained with ethidium bromide. 2.2 Colony Selection and Sequence Verification Seven isolated colonies were chosen at random and lysed in 50 µL of 20 mM Tris-HCl (2-Amino-2-hydroxymethyl-propane-1,3-diol hydrochloride) pH 8, 2 mM EDTA, and 1% Triton X-100. One µL of the lysate was added to 9 µL of Pfu polymerase PCR buffer, and 200 µM dNTPs, such that products were amplified with Pfu polymerase 200 bp 300 bp 500 bp 650 bp 850 bp 1000 bp 1650 bp 2000 bp 5000 bp 12000 bp 3000 bp 7000 bp 1     2     3 nicked, recircularized plasmid with insert supercoiled pET28a plasmid with insert extended megaprimer 29 using primers (0.4 µM) complementary to the T7 phage promoter and terminator located on the pET28a plasmid. Colonies with plasmids containing an insert were identified by the presence of a 600 bp band visualized by 1% agarose gel electrophoresis stained with ethidium bromide (Fig. 2-5). Colonies containing plasmids with no insert were predicted to yield bands at 200 bp. Due to the high efficiency of the employed method, all colonies tested were positive for inserts. DNA sequencing performed by Agencourt Bioscience Corporation (Beverly, MA, USA) confirmed that the gene was successfully inserted in- frame into the pET28a vector, and that the predicted translated amino acid sequence was: 1 N-GSEELLDLFNRQVTQEFTASQVYLSASIWFDQNDWEGMAAY 40 41 YMLAESAEEREHGLGFVDFANKRNIPIELQAVPAPVSCAEWSSP 84 85 EDVWQSILELEQANTRSLLNLAEAASTCHDFAVMAFLNPFHLQ 127 128 QVNEEDKIGSILAKVTDENRTPGLLRSLDVVSFLGPCLFRS-C 168 The N-terminal glycine was inserted via the initial megaprimer development to maximize pET28a expression efficiency (Murphy, M. personal communication). The reverse hybrid primer used in the first round of PCR imparted a stop codon sequence such that translation was terminated at the same point dictated by the P. multiseries ferritin gene. Except for the initial glycine, the final 168 amino acid sequence matched the peptide expected from the originally determined gene sequence (Marchetti et al., in prep). 30 Figure 2-5 Colony PCR to determine the presence or absence of an inserted sequence. Distance between T7 promoter and terminator is ~200 bp without insert, and ~600 bp with ferritin sequence. Insertion efficiency with this method was generally high, with only one colony in approximately 20 containing a plasmid with no insert. 2.3 Protein Sequence and Phylogenetic Comparison The confirmed P. multiseries ferritin sequence was compared with known amino acid sequences in the NCBI RefSeq non-redundant protein database (Pruitt et al., 2007) using the BlastP 2.2.18 algorithm (Altschul et al., 1997). Translated nucleotides encoding putative ferritins from the genome sequencing projects for Phaeodactylum tricornutum (US DoE JGI, 2006) and Cyanidioschyzon merolae (Matsuzaki et al., 2004) were conducted using tblastn (Altschul et al., 1997). Multiple sequence alignment was performed using MAFFT v. 6.240 (Katoh et al., 2002, 2005) with Blosum62 weighting matrix, and poorly aligning gapped regions were removed using BioEdit v. (Hall, ~600 bp ~200 bp 31 1999). Maximum likelihood bootstrapped phylogenetic trees (100 replicates) and unrooted trees were generated using PhyML v. 2.4.4 (Guindon et al., 2005) and PHYLIP (Felsenstein, 1993), respectively. 2.4 Purification of Recombinant Ferritin For expression of ferritin monomer, purified plasmids were transformed by electroporation into E. coli BL21 strain DE3 competent cells, and incubated overnight at 37 ˚C in 4 mL 2YT medium with 20 µg/mL kanamycin. 1 L cultures were grown at 37 ˚C in 2 L flasks of 2YT medium until they reached OD600 of at least 0.6, then were induced by adding 0.5 mM IPTG (isopropyl β-D-1-thiogalactopyranoside) and incubated overnight at 25 ˚C. Expression tests with varied induction temperature (25, 30, and 37 ˚C) and IPTG concentrations (0.1, 0.5, 1.0 mM) indicated that these conditions provided the optimal yield of soluble protein. Cultures were harvested by centrifugation for 10 min at 5000 g in 150 mL bottles, resuspended in approximately 30 mL of 20 mM Tris-HCl pH 8 with 1 mM TCEP (Tris 2- carboxyethyl phospine, Sigma) and 5% glycerol, and lysed by passing the sample through an Emulsiflex® Homogenizer (Avestin Inc., Ottawa, ON, Canada) 5 times at 10,000- 15,000 psi. The inclusion of the reducing agent TCEP helped to maintain the stability of ferritin in solution during the purification procedure, preventing non-specific disulfide bonds from forming. To minimize DNA contamination, the sample was also treated with DNase type II (100 units per mL) for 30 min at room temperature. Since ferritins are known for superior thermal stability (Bertrand and Harris, 1979; Stefanini et al., 1996), initial purification was achieved by apportioning the DNase 32 treated sample into mini centrifuge tubes, and heating for a maximum of 5 min at 65 ˚C in a heating block to selectively denature other proteins. Tubes were transferred to an ice bucket for immediate cooling, then centrifuged at 16,000 g for 10 min to pellet denatured proteins. The resulting supernatant was pooled, and spun at 12,000 g for 40 min in a refrigerated centrifuge. 15% SDS-PAGE confirmed that the clarified supernatant contained the majority of ferritin, with minimal losses to the heat shock pellet (Fig. 2-6). Comparison of pre and post-induction samples showed high levels of recombinant ferritin expression, making potential contamination by constitutive E. coli ferritins negligible. Figure 2-6 Confirmation of induction and heat shock purification of overexpressed ferritin. 15% SDS-PAGE of E. coli supernatant after overnight induction (lane 1), before induction (lane 2), heat shock denatured proteins (lane 3), and remaining supernatant following heat shock (lane 4). Protein molecular weight marker (Page Ruler™ unstained) from Fermentas International Inc. (Burlington, ON, Canada).  1      2       3      4 116.0 kDa 66.2 kDa 45.0 kDa 35.0 kDa 25.0 kDa 18.4 kDa 14.4 kDa 20 kDa ferritin monomer 33 The pI of the ferritin protein was estimated to be around 4.1 (Toldo, 1995) thus, half of the clarified supernatant (~15 mL) was injected into a GE ÄKTAexplorer™ FPLC™ system (GE Healthcare Bio-Sciences Corp., Piscataway, NJ, USA) and loaded on a 8 mL GE Source™ 15Q anion exchange column equilibrated with 20 mM Tris-HCl pH 8, 1 mM TCEP, and 5% glycerol at 4 ˚C. The buffer was prepared and titrated to pH 8 at room temperature, but was approximately pH 8.6 at 4 ˚C due to the temperature dependence of Tris. The progress of the run was monitored at 280 nm absorbance (relative absorbance units) and conductivity (mS cm-1) versus elution volume (Fig. 2-7). Figure 2-7 Source 15Q anion exchange purification of recombinant ferritin. UV absorbance at 280 nm (blue trace) and conductivity (pink trace) were monitored during the run. Ferritin peak eluted at 85 mL and 27 mS cm-1 conductivity. 34 Following the elution of the flow through, the NaCl concentration of elution buffer (1 M NaCl in equilibration buffer) was increased to 15% (150 mM NaCl and conductivity of 16 mS cm-1) and held there for 10 mL to displace most other contaminant proteins from the column. Ferritin was then eluted with a concentration gradient of elution buffer from 15% to 45% over 30 minutes or ~10 column volumes. The large ferritin containing peak began to elute at ~290 mM NaCl and 27 mS cm-1 conductivity. Ferritin containing fractions were collected and pooled with an estimated concentration of 4 mg mL-1, about 80 mg total ferritin yield. The pooled sample was then concentrated and further purified using an Amicon Ultra-15 centrifugal device (Millipore, Inc., Billerica, MA, USA) with a nominal molecular weight cut-off of 30 kDa. The final concentration after centrifugation was ~30 mg mL-1 and no protein precipitate was noted. The pinkish brown concentrate was mixed and stored in 1.5 mL centrifuge tubes in aliquots of 200 µL; half were stored at 4 ˚C and half were flash frozen in liquid nitrogen and stored at –80 ˚C. After one week, the samples stored at 4 ˚C had lost their pink tint, and appeared more yellowish brown; this had no apparent effect on stability or function. 2.5 Determination of Protein Size and Purity by Gel Filtration and SDS-PAGE To further assess purity and to determine the size of the native ferritin multimer, a 500 µL aliquot of the protein sample (~2 mg mL-1) was run on a Superdex® 200 HiLoad 16/60 gel filtration column (Amersham Biosciences) calibrated with 5 other native protein standards catalase (240 kDa, bovine), aldolase (160 kDa, rabbit), albumin (67 kDa, bovine), ovalbumin (45 kDa, bovine), and myoglobin (17.8 kDa, equine) (SERVA 35 Electrophoresis, Heidelberg, Germany; distributed by Helixx Technologies, Inc., Toronto, ON, Canada). The void volume of the column, 44.0 mL, was determined by the elution time of blue dextran (2000 kDa). The molecular weight determination of ferritin (as purified) was initially performed with the running buffer 20 mM Tris-HCl pH 8, 200 mM NaCl, 1 mM TCEP and 5% glycerol, similar to the conditions at which ferritin eluted from the anion exchange column. This was later repeated with apoferritin using the buffer in which Fe assays were conducted, 50 mM MES (2-N-morpholino-ethanesulfonic acid), 100 mM NaCl, pH 6.5, with no difference in ferritin elution volume (Ve). Using denaturing 12% acrylamide SDS-PAGE and linear regression, the molecular weight of the ferritin monomer was also determined. 2.6 Protein Determination Protein concentration was determined prior to each assay using a modified Lowry method (DC protein assay, Bio-Rad Laboratories, Hercules, CA, USA) with bovine serum albumin used to generate a standard curve (R value no less than 0.99). 100 µL of several dilutions of the ferritin solution to be measured were mixed with 500 µL of an alkaline copper tartrate solution and 4 mL of a dilute folin reagent, and vortexed immediately. Using a Cary 50 Bio UV-visible spectrophotometer (Varian Inc., Palo Alto, CA, USA), the absorbance at 750 nm was recorded after at least 20 min. The third component of the kit, reagent S (an SDS solution) was not added since there was no detergent present in any of the samples. The modified Lowry method is thought to be superior to the original assay (Lowry et al., 1951; Peterson, 1979), since it can be done in one step at room 36 temperature, and the colour develops quickly and is stable for up to two hours or more (Stoscheck, 1990). The linear range of this assay is superior to the Bradford assay (Bradford, 1976), it is compatible with detergents, and does not precipitate, making it a more reliable choice for most protein concentration determinations. The non-linearity sometimes observed with the Bradford assay is mainly due to the equilibrium that exists between the two forms of the dye used, and this is perturbed in the presence of protein. It is possible to enhance the sensitivity and linearity of the assay by measuring the ratio of the absorbance at 595 to 450 nm (Zor and Selinger, 1995); however, this was not explored in this study. For Fe-unloaded apoferritin, corresponding absorbances at 280 nm were compared with Lowry protein determinations in order to generate a molar extinction coefficient. 2.7 Spectroscopic Assay of Iron Binding by Apoferritin Apoferritin was prepared by dialyzing several mL of purified ferritin in anion exchange elution buffer (~2 mg mL-1) against a solution of 3% sodium dithionite (Na2S2O4, Sigma) in 1 M sodium acetate buffer pH 4.8, for 8 hours to remove endogenous Fe from the protein (Granick and Michaelis, 1942; Macara et al., 1972; Treffry et al., 1992). The sodium dithionite reduces Fe(III) bound to ferritin, and the acetate chelates the Fe(II) released from the protein. Although a small amount of fine white precipitate was noted, this disappeared completely after the sample was dialyzed overnight against 50 mM MES buffer pH 6.5 with 100 mM NaCl, the buffer in which all downstream Fe binding assays were conducted. pH 6.5 was chosen for all assays because 37 this has been shown to be optimal, balancing Fox activity and minimizing Fe precipitation in horse spleen ferritin (Yang and Chasteen, 1999). Following the methods of Yang et al., 2000a and Bou-Abdallah et al., 2002, the presence of a dinuclear ferroxidase site was assayed spectroscopically in a quartz cuvette by aerobically titrating 1 mL of a ~1 µM solution of purified ferritin with 1 or two µL of 6 mM ferrous sulfate heptahydrate solution (FeSO4•7H2O, Sigma, pH 3.5 in water), up to a maximum of 1370 Fe:ferritin (1040 µM added Fe). Following each Fe addition, absorbance spectra (800-200 nm) were recorded after 3 min, and the formation of the ferroxidase products (di-ferric peroxo mineral precursors and the mineral core) was tracked by monitoring the increase in absorbance at 295 nm (Treffry et al., 1992). When the ∆A295 peaks were plotted against Fe added (µM) per µM ferritin, the mean molar absorptivity coefficient (ε) per bound Fe atom was calculated using Beer’s Law (Eqn 20), where b is the pathlength (1 cm) and c is the protein concentration, which was carefully measured prior to each assay.       ∆A295 = εbc [20] 2.8 Determination of Ferritin Iron Content Fe titrated protein samples were dialyzed extensively against 50 mM MES, 100 mM NaCl, pH 6.5 to remove any unbound Fe. Their Fe content was then determined using the Ferene S assay (Haigler and Gibson, 1990), accurate to nmol quantities of Fe in a 3 mL reaction volume. All glassware was soaked overnight in 1% HCl to remove extraneous Fe contamination. The Ferene S reagent was prepared by dissolving 45% sodium acetate, 10 mM ascorbic acid, and 0.75 mM Ferene S® reagent (3-(2-pyridyl)-5,6 38 bis-[2-(5-furyl sulfonic acid)]-1,2,4-triazine-5’,5”-disodium salt) (Sigma-Aldrich Canada, Ltd., Oakville, ON, Canada) in 50 mL deionized water with heat and constant stirring. The reagent was then covered and kept on ice until use. Standards were freshly prepared in 1.5 mL centrifuge tubes by mixing 0, 2, 4, 6, 8, 12, and 16 µL of 2mM FeCl3 with a volume of 50 mM MES buffer pH 6.5 with 100 mM NaCl, equal to that used for the measurement of the protein samples, and then diluted to 780 µL with H2O. This yielded amounts of total Fe equal to 0, 4, 8, 12, 16, 24, and 32 nmol Fe, respectively. Sample volumes were adjusted such that measured Fe content fell within the bounds of the calibration curve. To each standard and protein sample in triplicate, 120 µL of 10 M HCl was added and mixed by inverting periodically for 10 minutes. 100 µL of 80% TCA (trichloroacetic acid, Sigma-Aldrich) was added, and solutions were centrifuged for 10 min at 16,000 g. The resulting supernatant was transferred to glass test tubes where 200 µL of 45% acetic acid and 1.8 mL of Ferene S® reagent were quickly added and vortexed. Measurements were taken using a Cary 50 bio spectrophotometer blanked with water, and the absorbance of all standards and unknowns was measured at 593 nm, a wavelength corresponding to the deep blue colour generated by Fe containing solutions. Fe content of the unknowns was determined by comparison to the standard curve (R value no less than 0.99). Protein concentrations were determined in triplicate using the modified Lowry method described above. 39 2.9 Oxygen and Hydrogen Peroxide Measurements To categorize the chemical characteristics of the recombinant ferritin ferroxidase reaction, an Apollo 4000 Free Radical Analyzer (World Precision Instruments, Sarasota, FL, USA) was used to monitor the consumption of oxygen following Fe(II) addition to the protein solution. Measurement of dissolved O2 was conducted using a Clark type electrode (Clark et al., 1953) consisting of a platinum working electrode with a poise voltage held at –0.7 V and an Ag/Cl reference electrode covered by a gas permeable membrane. The electrode measures the change in current generated by the reaction:       O2 + 2e- + 2H2O  H2O2 + 2OH- [21] The O2 electrode was equilibrated in 2 mL of 50 mM MES buffer pH 6.5 with 100 mM NaCl (5.844 g kg-1 salinity) open to the atmosphere for at least 4 hours with constant stirring, such that the initial level of dissolved oxygen was in equilibrium with the atmosphere. A temperature probe was also inserted into the solution, enabling the calculation of theoretical atmospheric O2 saturation in order to generate an accurate value for dissolved oxygen in this solution. Adding 1 mg of sodium dithionite allowed the current at zero oxygen to be measured; the oxygen electrode was calibrated with these two points (Fig. 2-8). After calibration, re-equilibration of the electrode was performed with a 2 mL solution of ~1 µM ferritin in the same buffer. Upon stabilization of the O2 reading, 4µL of 6mM FeSO4•7H2O in water (pH 3.5) was added (increasing the final concentration of Fe(II) by 12 µM) and the change in [O2] was recorded (similar to Yang et al., 2000a). 40 Following Fe addition and maximum O2 response, the evolution of hydrogen peroxide from the Fox reaction was determined as in Xu and Chasteen, (1991) and Bou-Abdallah et al. (2002) by adding 1 µL of 20 mg mL-1 catalase, recovering oxygen in the ratio 2H2O2:1O2. O2 saturation of the assay solution was corrected for temperature and salinity according to the equation based on Henry’s Law by Garcia and Gordon (1992), and was calculated using O2sol Version 1.1 (Hamme, 2005). The electrode current readings were converted to dissolved oxygen concentration (µM) using the daily calibration equation (Fig. 2-8) and plotted. Figure 2-8 Two-point calibration of O2 electrode (Zero O2 and atmospheric O2). Atmospheric measurement was made after at least 4 hours of stirring with the electrode assembly open to the air. Zero was obtained by adding 1 mg sodium dithionite. The calibration was performed daily and the resultant correction equation was applied to electrode current data. 41 Chapter Three: Results 3.1 General Properties of Recombinant Ferritin Prior to attaining the 168 amino acid protein upon which all assays in this study are based, a different construct was initially created, which included an N-terminal 6x HIS tag and thrombin cleavage site. It was also slightly truncated, terminating after proline 162 (Fig. 2-1). The shortened construct was created to avoid the inclusion of residue 163, a potentially destabilizing 3rd cysteine residue. Unfortunately, SDS-PAGE of various thrombin-digested ferritin treatments indicated that only some of the 24 HIS tags could be cleaved before degradation of the protein occurred. Furthermore, while the HIS- tagged construct demonstrated Fox activity upon titration with Fe(II), the protein’s stability below pH 7.5 was very low, and the total Fe binding capacity of the protein was limited to about 40 Fe per ferritin shell (data not shown). Addition of Fe beyond ~40 Fe:protein resulted in the formation of iron and protein precipitate. To improve the stability and activity of the recombinant ferritin, the carboxyl terminus of the protein was extended in length by 5 amino acids to include the entire fixth helix predicted by DISOPRED2 (Fig. 2-1), and the 6x HIS tag was omitted. Transformation efficiency was high overall (Fig. 2-5), due primarily to the use of the enzyme Dpn-1 to degrade unamplified DNA. Expression of ferritin was also excellent with the extended construct (Fig. 2-6), yielding up to 200 mg ferritin per litre of culture, the majority of which was soluble. Downstream assays of the protein were therefore ameliorated, as it was stable in a variety of buffers at a range of pH. 42 3.2 Phylogenetic Analysis BlastP query of the confirmed P. multiseries ferritin protein sequence against the NCBI RefSeq non-redundant protein database (Pruitt et al., 2007) indicates that it belongs to the ferritin-like protein superfamily, and its conserved domains compare closely with non-heme ferritins (Altschul et al., 1997; Pruitt et al., 2007). Table 3-1 represents a collection of 29 ferritins from various organisms and encompasses the main classes of ferritin examined in this study, and includes the closest known relatives of the P. multiseries protein (E-values < 1e-13). Overall amino acid sequence identity with its most similar homolog is about 36%, with less than 60% amino acid similarity (Fig 3-1). Despite the sequence variability among the diverse classes of ferritin, the most common feature of nearly all sequences is the conservation of the amino acyl residues that comprise the ferroxidase site. Figure 3-1 shows the location of residues that characterize the Fox sites of the protein; other conserved regions are involved in the formation of the channels by which Fe enters the Fox sites, Fe release, and protein folding. From this alignment it seems that the position of the Fox residues is particularly unique in the three Dps ferritins. 43 Table 3-1 Collection of 29 selected ferritin taxa including closest relatives of P. multiseries used for phylogenetic analysis. Organism name (common name) Strain/isolate (if known) Accession number Type of ferritin1 Pseudo-nitzschia multiseries (coastal pennate diatom) CLN-47 FTN Synechococcus sp. (marine cyanobacterium) WH 5701 ZP_01085349 FTN Chloroflexus aggregans (thermophilic phototrophic bacterium) DSM 9485 ZP_01516889 Likely FTN Macrobrachium rosenbergii (freshwater prawn) ABY75225 Euk Candidatus methanoregula boonei (methanogenic peat bog archaeon) 6A8 YP_001403955 Likely FTN Pacifastacus leniusculus (signal crayfish) CAA62186 Euk Prochlorococcus marinus (marine cyanobacterium) MIT 9215 YP_001484100 FTN Pyrococcus furiosus (hyperthermophilic archaeon) DSM 3638 NP_578471 FTN Anaerostipes caccae (bacterium) DSM 14662 ZP_02420883 Likely FTN Thermotoga maritima (hyperthermophilic bacterium) MSB8 NP_228934 FTN Synechococcus sp. (Red Sea cyanobacterium) RS9916 ZP_01471843 FTN Equus caballus (horse H-chain) NP_001093883 Euk (H-chain) Homo sapiens (human H-chain) NP_002023 Euk (H-chain) Salmo salar (salmon H-chain) AAB34575 Euk (H-chain) Rana catesbeiana (bullfrog H-chain) P07729 Euk (H-chain) Helicobacter pylori (bacterium) CAA76033 FTN Phaeodactylum tricornutum (coastal pennate diatom) CCAP1055/1 16343 (JGI ID) Likely FTN Cyanidioschyzon merolae (extremophilic red alga) 10D AP006498 Likely Euk Glycine soja (soybean) ABP68836 Euk Chlamydomonas reinhardtii (green alga) ABW87266 Euk Thermosynechococcus elongatus (thermophilic cyanobacteria) BP-1 NP_682531 Likely FTN Escherichia coli (bacterium) K-12 MG1655 NP_417795 BFR Trichodesmium erythraeum (marine nitrogen-fixing cyanobacterium) IMS101 YP_723752 Dps Pseudomonas aeruginosa (proteobacterium) PAO1 NP_252925 BFR Synechocystis sp. (freshwater cyanobacterium) PCC6803 NP_442825 BFR Uncultured Crenarchaeote (archaeon) 4B7 AAK68802 Likely Dps Rhodobacter sphaeroides (purple bacterium) 2.4.1 YP_351589 BFR Neisseria gonorrhoea (bacterium) FA 1090 YP_207920 BFR Listeria innocua (bacterium) Clip11262 NP_470279 Dps 1 Non-heme, homopolymer ferritin (FTN); heme-binding bacterioferritin (BFR); DNA binding protein from starved cells (Dps); heme-binding heteropolymer eukaryotic ferritin (Euk). 44       0                                                                    70 P. multiseries_FTN 100%  LDLFNRQVTQEFTASQVYLSASIWFDQNDWEGMAAYMLAESAEEREHGLGFVDFANKRNIPIELQAVPEW Synechococcus_WH5701_FTN 36.1% HQGLCNHLQMELMASYTYFSLAVWFTQRELNGFAAFARAESDGERAHAGLFVDYLVARSQPVELSALSTW Chloroflexus_aggregans 34.0% LQALNRQITYEYAASYTYLATAAYFESLSLTGFAHWFRVQSEEEREHALRFFDYVNDRGGRVMLGAIDEF Macrobrachium_rosenbergii 34.0% EALINKQINMELYASYVYMSMSHYYDDVALPGMSHFFKKSSDEEREHANKLMKYQNSRGGRIVLQAIAEW Candidatus_methanoregula 33.3% EEALNRQINRELYSSYLYLGMAAYFESVNLKGFASWMLVQSNEERGHAMKFYDYVYARQGKVVLDAIEKW Pacifastacus_leniusculus 33.3% E-PINKQINLEFYASYVYMSMGHYFDDISLPGASKFFKDSSDEEREHGQKLMKYQNKRGARIVLQAIAEW P._marinus_MIT9215 32.7% LDNFFEHLTMERFANVQYFSIYLWFQERDLNGFASYFLRESHGEMEHAKKFADYLIARGQTVKLNDIPNW Pyrococcus_furiosus_FTN 32.7% LKALNDQLNRELYSAYLYFAMAAYFEDLGLEGFANWMKAQAEEEIGHALRFYNYIYDRNGRVELDEIPEW Anaerostipes_caccae 32.0% AELMNDQINKEFYSSYLYLDMSNYYVDKNLDGFANWFKIQAQEERDHAILFMEYLQANNCRVTLEAVASF Thermotoga_maritima_FTN 31.3% RKALNEQLNREIYSSYLYLSMATYFDAEGFKGFAHWMKKQAQEELTHAMKFYEYIYERGGRVELEAIENW Synechococcus_sp._RS9916 29.9% VGAIQQHINLELEASMTYLSMSIWCAERELAGFYKFFSAESAEERGHAIQFADYLVARAQRNDLQPLQNW Equus_caballus_Euk 29.3% EAAINRQINLELHASYVYLSMSFYFDDVALKNFAKYFLHQSHEEREHAEKLMKLQNQRGGRIFLQDIKDW Homo_sapiens_Euk 29.3% EAAINRQINLELYASYVYLSMSYYFDDVALKNFAKYFLHQSHEEREHAEKLMKLQNQRGGRIFLQDIKDW Salmo_salar_Euk 29.3% EAAINRQINLELYASYVYLSMAYYFDDQALHNFAKFFKNQSHEEREHAEKLMKVQNQRGGRIFLQDVKEW Rana_catesbeiana_Euk 28.6% EAAINRMVNMELYASYTYLSMAFYFDDIALHNVAKFFKEQSHEEREHAEKLMKDQNKRGGRIVLQDVKEW Helicobacter_pylori_FTN 27.9% IKLLNEQVNKEMNSSNLYMSMSSWCYTHSLDGSGLFLFDHAAEEYEHAKKLIVFLNENNVPVQLTSISKF Phaeodactylum_tricornutum 27.9% ATLLVNQANRELDASRMYLAMDMWFRFHDFPGSASWCQTHSVEERSHAMKIFDHLALRQTEKVLSVSQQN Cyanidioschyzon_merolae 26.5% EEAINSQINVEFTAFYVYYALHAYFDTVALPGFADYFRKQAEEERDHAVKLMHYQNKRGGRVHLKPIAEN Glycine_soja_Euk 26.5% ESAINEQINVEYNASYVYHSLFAYFDNVALKGFAKFFKESSEEEREHAEKLMKYQNTRGGRVVLHPIKEK C._reinhardtii_Euk 25.2% EAAINEQVNIEYNVSYLYHALWAYFDNVALPGLAAFFKAGSEEEREHAELLMEYQNRRGGRVVLGAISEK T._elongatus 22.4% INRLNAQINLEMFSARLYLQMSSWCAHKALEGCATFLGQHADEEMAHMRRLLSYMHETGALAILEGLENF Escherichia_coli_BFR 15.2% INYLNKLLGNELVAINQYFLHARMFKNWGLKRLNDVEYHESIDEMKHADRYIERILFLEGLPNLQDLGIG T._erythraeum_Dps 14.6% CEGMNVALASCQALYLQYEKHHFVVEGAEFNQLHEFFKESYEEVKEHVHEMAERLNGLGGVPSFAKLAGS Pseudomonas_aeruginosa_BFR 13.5% IDYLNTLLTGELAARDQYFIHSRMYEDWGFSKLYERLNHEMEEETQHADALLRRILLLEGTPRMRPDDPG Synechocystis_sp._BFR 12.7% LAQLHKLLRGELAARDQYFIHSRMYQDWGLEKLYSRIDHEMQDETAHASLLIERILFLEETPDLSQQDVG Crenarchaeote_AAK68802 12.2% VKQLITNASVEFTAYYYFTNLRMHCTGMEGEGIKGIIEDARLEDLSHFESCIERIFQLGGSLDAQAFTNP R._sphaeroides_BFR 11.6% IEYLNAALRSELTAVSQYWLHYRLQEDWGFGSIAHKSRKESIEEMHHADKLIQRIIFLGGHPNLQRLNIG Neisseria_gonorrhoea_BFR 11.3% VDYMNELLSGELAARDQYFIHSRLYSEWGYTKLFERLNHEMEEETTHAEDFIRRILMLGGTPKMARSEIG Listeria_innocua_Dps 10.4% KEFLNHQVANLNVFTVKIHQIHWYMRGHNFFTLHEKMDDLYSEFGEQMDEVAERLLAIGGSPTLKEFLKP          71                                                                           147 P. multiseries_FTN SSPEDVWQSILELEQANTRSLLNLAEAASTCHDFAVMAFLNPFHLQQVNEEDKIGSILAKVTDENTPGLLRSLDVVL Synechococcus_WH5701_FTN TNVEDVLVSVFEMEAEVTTSLQQLYSLAERCGDYRSSIFLDPLIKSQVDAESEVAHLLAQVRHCGDFGALLILDQSL Chloroflexus_aggregans ASPLDAFEYALAHEQRVTASINAIYALAAQENDYATMSMLKWFIDEQVEEEKSVDEIIRHLKLVGDGVGLLLLDRQL Macrobrachium_rosenbergii GSALDGLQAALDLEKQVNQSLLDLHGTASTANDPHLTKFLEGYLEEQVESIKELGDMITKLKRAGTGLGEYLFDKEL Candidatus_methanoregula TSSGKVFEEVYAHEQKVTGLINNLVELATKEKDHATFEFLQWFVKEQVEEEANAALIVDKIKTLGIPGHLFYLDHEL Pacifastacus_leniusculus GNLHDALQAALDLENEVNQSLLDLDATASKINDPHLTNMLEEFLEEQVESIEKIGNLITRLKRAGSGLGEFLFDKEL P._marinus_MIT9215 DSIEDLISYSFNMEADLTSSLQQLYSISERNSDTRTNVFLDPIVEAQIKSEDEFANILGKVKFASQPSAIFLIDSDL Pyrococcus_furiosus_FTN ESPLKAFEAAYEHEKFISKSIYELAALAEEEKDYSTRAFLEWFINEQVEEEASVKKILDKLKFAKSPQILFMLDKEL Anaerostipes_caccae EEPADPLKAALEHEQYVTSLINNIYDAAYQCKDFRSMQFLDWFVKEQMEEENNTDSLVQKFEMFGDPKGLYMLDAEL Thermotoga_maritima_FTN NGIKDAFEAALKHEEFVTQSIYNILELASEEKDHATVSFLKWFVDEQVEEEDQVREILDLLEKANQMSVIFQLDRYL Synechococcus_sp._RS9916 SKLEDVVFSSFRMEADTTSSIQHLYSIAERENDVRTTVFLDPLLEQQIGSEDQYAYLHGRIKFADDSTALLVIDGEV Equus_caballus_Euk ENGLKAMECALHLEKNVNESLLELHKLATDKNDPHLCDFLEHYLNEQVKAIKELGDHVTNLRRMGSGMAEYLFDKHT Homo_sapiens_Euk ESGQNAMECALHLEKNVNQSLLELHKLATDKNDPHLCDFIEHYLNEQVKAIKELGDHVTNLRKMGSGLAEYLFDKHT Salmo_salar_Euk GSGVEALESSLQLEKSVNQSLLDLHKVCSEHNDPHMCDFIEHYLDEQVKSIKELGDWVTNLRRMGNGMAEYLFDKHT Rana_catesbeiana_Euk GNTLEAMQAALQLEKTVNQALLDLHKVGSDKVDPHLCDFLEEYLEEQVKSIKQLGDYITNLKRLGNGMGEYLFDKHT Helicobacter_pylori_FTN ESLTQIFQKAYEHEQHISESINNIVDHAIKSKDHATFNFLQWYVAEQHEEEVLFKDILDKIELIGQNHGLYLADQYV Phaeodactylum_tricornutum NRVSEVWKKALDQEVQNSQEYFNMAKVAEEESDYVSREFLNWFLNEQLMEENAVEDLYRKAIKLETGGLYVAIDKDM Cyanidioschyzon_merolae SDAIYAMELALQLEKYVQMKLMEVWKVADRERDANMTDFIEDFLDMQVESIKEISDYVAQLKRVGTGHGVYHFDRVL Glycine_soja_Euk GDALYAMELALSLEKLVNEKLLNVHSVADRNNDPQMADFIEEFLSEQVESIKKISEYVAQLRRVGKGHGVWHFDQRL C._reinhardtii_Euk GDALYAMELALSLEKLNFQKLRQLHSVADEHGDASMADFVEELLNEQVEAVKKVSEYVSQLRRVGQGLGVYQFDKQL T._elongatus GSLKEMFSQVYSHEQLVTRKINELVHLANSEPDYSTLQFLQWYVAEQHQEEFLFKSILDKIDLIGEGQGLFFIDQEI Escherichia_coli_BFR EDVEEMLRSDLALELDGAKNLREAIGYADSVHDYVSRDMMIEILRDEEGHIDWLETELDLIQKMGYLQAQIREEG—- T._erythraeum_Dps FNARNMIEHDLKAEQDVIKLLRRLAAQAESLGDRATRYLYEKILLETEERAYHLDHFLA-------PDTLVVLN--- Pseudomonas_aeruginosa_BFR TTVPEMLEADLKLERHVRAALAKGIALCEQHKDFVSRDILKAQLADEEDHAYWLEQQLGLIARMGYLQSQI------ Synechocystis_sp._BFR KTVPEMLQYDLDYEYEVIANLKEAMAVCEQEQDYQSRDLLLKILADEEDHAYWLEKQLGLIEKIGYLQSQMS----- Crenarchaeote_AAK68802 TDIKAILEKCLKAEQGAIVNWDIVCKMTYN-KDPATYDIAKDILMEEIEHESWFLELLYGRPSGHKFSGERPHTQKH R._sphaeroides_BFR QTLRETLDADLAAEHDARTLYIEARDHCEKVRDYPSKMLFEELIADEEGHIDYLETQIDLMGSIGYLNAKPADEAE- Neisseria_gonorrhoea_BFR TDVVSCLKADLQTEYEVRDALKKGIKLCEEAQDYVTRDLMVAQLKDEEDHAHWLEQQLRLIELIGYYQSQL------ Listeria_innocua_Dps KTMDQLMEDLVGTLELLRDEYKQGIELTDKEGDDVTNDMLIAFKASIDKHIWMFKAFLGKAPLE------------- Figure 3-1 Selectively Degapped MAFFT amino acid sequence alignment of 29 selected ferritin amino acid sequences. Ferroxidase site residues are highlighted in gray; algal taxa are highlighted in green; percentage values indicate amino acid sequence identity to P. multiseries ferritin. 45 Sequence alignment indicates that P. multiseries ferritin is not clearly analogous to any one type of ferritin in particular. The primary structure of P. multiseries ferritin compares with the ferritins of thermophilic bacteria, archaea, cyanobacteria, and even mammals - a mixed collection of FTN, BFR, Dps, and eukaryotic (Euk) ferritins (Table 3-1, Fig. 3-1). A search of the completely sequenced genomes of the centric diatom Thalassiosira pseudonana (US DoE JGI), the marine pennate diatom Phaeodactylum tricornutum (US DoE JGI), and the primitive, extremophilic red alga Cyanidioschyzon merolae (Matsuzaki et al., 2004) indicates that the T. pseudonana genome does not appear to contain a ferritin-encoding gene, while P. tricornutum and C. merolae do, although they share relatively low sequence homology with P. multiseries. The diatom protein has no known equivalent in any Chromalveolate genera. An unrooted phylogenetic tree built from the closest ferritin relatives to P. multiseries, as well as a number of other representative mammalian, bacterial, and plant ferritins shows that the main groupings of FTN, Dps, BFR, and eukaryotic ferritins tend to branch together (Fig. 3-2). It also suggests that P. multiseries ferritin (which does not branch with the other eukaryotic ferritins) is more closely related to P. tricornutum and the cyanobacterial genera examined. There are a number of ferritins from prokaryotic and eukaryotic sources that have similar sequence identity, in particular the thermophilic photoautotrophic bacterium Chloroflexus aggregans, the freshwater prawn Macrobrachium rosenbergii, and the archaeon Candidatus methanoregula. However, conserved similarities are also factored into the MAFFT alignment using a weighted amino acid substitution matrix (Blosum62) prior to the computation of the unrooted tree, which leads to a more informative phylogeny. 46 To further refine the phylogenetic analysis, a maximum likelihood rooted tree was also created (Fig. 3-3). Most bootstrap values are high, indicating that there is a high likelihood that nodes are depicted in their correct locations, and the main classes of ferritin are well differentiated from each other. Within the group of non-heme FTNs, however, only the cyanobacterial ferritins are defined with a high degree of confidence – the nodes differentiating the diatoms from the eukaryotic and prokaryotic ferritins, and the node between eukaryotic and prokaryotic ferritins themselves are less certain, with likelihoods under 50%. Even the terminal node separating the two diatom ferritins is not well defined. 47 Figure 3-2 Unrooted phylogenetic tree of selected ferritin sequences. Tree was generated using PHYLIP (Phylogeny Inference Package version 3.5c; Felsenstein, 1993) with the neighbour-joining method (Saitou and Nei, 1987) and excluding positions with gaps. The degapped MAFFT sequence alignment (Blosum62 substitution matrix) shown in Figure 3-1 was used as the input data. Groupings of major ferritin classes are indicated; arrow indicates potential lateral gene transfer from cyanobacteria. ? 48 Figure 3-3 Maximum likelihood rooted phylogenetic tree. Bootstrap values are based on 100 replicates using PhyML v2.4.4 (Guindon et al., 2005) and Blosum62 weighting matrix. Dps Ferritins (non-heme) Mammalian Euk Ferritins Red/Green Algae, Higher Plant Euk Ferritins Arthropod Euk Ferritins Prokaryote (non-heme) FTN Pennate Diatom (non-heme) FTN Cyanobacteria (non-heme) FTN Bacteria (heme) BFR Synechococcus sp. Uncultured Crenarchaeote A. caccae H. pylori T. elongatus C. aggregans C. merolae C. reinhardtii G. soja E. caballus H. sapiens S. salar R. catesbeiana P. leniusculus M. rosenbergii P. multiseries P. tricornutum . trSynechococcus sp. Prochlorococcus sp. L. innocua T. erythraeum R. sphaeroides E. coli Synechocystis sp. P. aeruginosa N. gonorrhoeae C. methanoregula T. maritima P. furiosus 49 3.3 Protein Purification Heat shock (65˚C for 5 minutes) proved to be an effective first purification step, eliminating roughly half of the total protein while enriching the overexpressed P. multiseries ferritin monomeric band relative to other E. coli proteins (Fig. 2-6). Some ferritin was detected in the heat shock pellet, however, this could represent partially formed heat labile multimers. Following anion exchange purification and the subsequent centrifugal concentration, a ferritin solution of near purity was achieved (Fig. 3-4). Figure 3-4 Denaturing 12% SDS-PAGE of ~1 µg purified ferritin. The MW of the monomer was calculated to be 20.0 kDa based on Page Ruler™ prestained MW ladder (Fermentas International Inc., Burlington, ON, Canada). 170 kDa 130 kDa 100 kDa 70 kDa 55 kDa 40 kDa 35 kDa 25 kDa 15 kDa 20.0 kDa ferritin monomer 50 Gel filtration of purified ferritin (Fig. 3-5) confirmed the high purity of the protein solution. When the elution volumes (Ve) of four independent ferritin runs were compared with the mean column calibration curve generated with two independent runs (Fig. 3-5 inset), the MW of apoferritin was determined to be 529 ± 16 kDa. Unfortunately, no larger standards between the column void volume of 44.0 mL and the elution volume of ferritin 61.1 mL were available for a more accurate size determination. However, given that the ferritin purified in this study falls on the best-fit calibration curve, the size determination is reasonable. Figure 3-5 Gel filtration chromatograph of apoferritin. Injection consisted of 2 mg purified apoferritin in 500 µL 50 mM MES buffer, 100 mM NaCl, pH 6.5. Absorbance was followed at 280 nm (peak at 61.1 mL). Inset shows column calibration curve averaged from two identical runs (y = -0.0246x + 4.2284). The first protein to elute in the calibration series (Ve = 61.1 mL) is the purified ferritin from this study. 51 3.4 Spectroscopic Analysis of Ferritin The UV-visible absorbance spectrum of P. multiseries apoferritin is nearly featureless, demonstrating a broad protein peak at 280 nm, due to the absorbance of aromatic amino acid side chains, and a small, poorly resolved peak at 290 nm (Fig. 3-6). Figure 3-6 UV-visible absorbance spectrum of apoferritin. The blank corrected spectrum of 0.76 µM purified apoferritin in 50 mM MES, 100 mM NaCl, pH 6.5 was collected by scanning from 800 nm to 250 nm. The molar extinction coefficient and absorbance at 280 nm were calculated for the fully assembled apoferritin shell (ε280nm = 655,000 M-1 cm-1 and A280nm = 1.23 for a 1 mg mL-1 ferritin solution, respectively) by generating a linear curve of the absorbance of ferritin (MW of 530 kDa) at 280 nm versus the protein concentration calculated using the 52 DC protein assay (3 replicates). The calculated values are comparable to the theoretical ε280nm = 643,000 M-1 cm-1 and A280nm = 1.33, respectively, obtained by the method of Gill and von Hipple, 1989. These values were used to determine protein concentration in subsequent spectroscopic assays. 3.5 Spectroscopic Determination of Ferroxidase Activity Addition of 6 mM FeSO4•7H2O in 1 or 2 µL increments to a 1 mL solution of apoferritin resulted in an increase in absorbance at 295 nm. After each Fe addition, the change in absorbance was observed within the first minute. Subtracting the initial apoferritin spectrum (Fig. 3-6) from successive scans following each Fe addition yielded the series of difference absorbance spectra shown in Figure 3-7. The growing peak at 295 nm is indicative of the formation of the ferroxidase products (di-ferric peroxo mineral precursors and the mineral core) and can be used to monitor the reaction progress (Treffry, 1992). The addition of Fe(II) to buffer in the absence of protein yielded no significant change in absorbance. The change in absorbance (∆Abs) at 295 nm against the concentration of Fe added per starting concentration of protein (Fig. 3-8), shows that the reaction proceeds by two visible rates demonstrated previously by Yang et al. (2000a) and Bou-Abdallah et al. (2002). The initial slope is due to bound Fe undergoing oxidation at the Fox site (phase 2, Eqns 5 and 12), and increases up to a discontinuity, beyond which an alternate oxidation process, probably ferrihydrite (FeOOH) core formation (phase 3, Eqns 6 and 15), proceeds with a lower molar absorptivity (Yang et al., 1998). Here, the molar absorptivity for the absorption increase at 295 nm per Fe added for the phase 2 reaction was variable, 53 ~5700-6100 M-1 cm-1. As the titration progressed, the value diminished gradually before stabilizing at about 2650 M-1 cm-1, the average value for the phase 3 reaction. For P. multiseries ferritin, the interface between these two processes exists at 22.4 ± 2.7 Fe:protein, and reflects the Fe binding stoichiometry of the 24 ferroxidase sites within the protein. Figure 3-7 Difference absorbance spectra of ferritin titrated with ferrous iron. 0.76 µM purified ferritin in 50 mM MES, 100 mM NaCl, pH 6.5 was titrated with 12 mM FeSO4•7H2O. The UV-visible spectrum of apoferritin before iron addition was subtracted from spectra recorded after each iron addition. Ferroxidase site activity is characterized by increasing absorbance at 295 nm. 54 Figure 3-8 Change in absorbance at 295 nm versus iron added per ferritin shell. The increase in absorbance was calculated by subtracting the absorbance spectrum of apoferritin prior to Fe loading from the spectra recorded after each addition of FeSO4•7H2O for 3 independent titrations. Triangles correspond to ~295 nm peak absorbance values in Figure 3-7; circles and squares represent two additional replicates. Total Fe added is normalized to the starting concentration of ferritin (~0.76 µM). Titrations were performed in 50 mM MES, 100 mM NaCl, pH 6.5. Fe oxidation at ferroxidase site Ferrihydrite core formation Discontinuity at 22.4 ± 2.7 55 3.6 Ferritin Iron Content and Binding Capacity To determine the Fe binding capacity of the ferritin construct, a Ferene S® colourimetric assay was used. To emphasize the inflection point, Figure 3-8 is truncated after the addition of ~100 Fe:protein equivalents. However, the titration can routinely be taken to several hundred Fe:protein, while the curve remains linear and the solution free of precipitate. At about 500 equivalents of Fe, a minimal amount of Fe precipitate could be seen in the reaction cuvette during the titration, and this continued to increase as more Fe was added. The effect appeared to be more pronounced when Fe was added in larger increments. When the precipitate was spun down to form a small, rust coloured pellet, the remaining supernatant was evaluated for protein and iron content and found to exhibit an average Fe:ferritin ratio of 633. Comparison to the sample prior to centrifugation indicated that only 14% of the initial protein precipitated, while 42% of the added Fe was found in the pellet. Table 3-2 Ferene S® determination of Fe bound by ferritin. Sample [Fe]/[Protein]1,2 Ferritin As Purified (1.83 µM, 200 µL sample) 11.78 ± 0.31 Unloaded Apoferritin (4.87 µM, 200 µL sample) 0.052 ± 0.01 Fe Loaded Ferritin (0.5 µM, 200 µL sample) 117.2 ± 1.6 (119 equivalents added)3 Maximum Fe Loaded Ferritin (0.33 µM, 50 µL sample) 633.0 ± 73.0 (1370 equivalents added) 3 1 Background Fe was < 0.5 nmol per 200 µL sample (2.5 µM) 2 [Protein] refers to µM ferritin shell 3 Molar ratio of Fe:apoferritin added during titration 56 3.7 Dissolved Oxygen and Hydrogen Peroxide Measurement To study the characteristics of the ferroxidase reaction in this ferritin, dissolved oxygen was monitored during additions of Fe(II) to 1 mL of a 0.76 µM apoferritin solution similar to that used for spectroscopic assays. Figure 3-9 represents the drop in dissolved oxygen from atmospheric saturation (versus protein-free control) that is associated with O2 consumption during the oxidation of Fe at the Fox active site. The rate of the initial drop in O2 is nearly 0.8 µM  s-1 per µM protein, only slightly lower than the rates inferred from the study of E. coli BFR (Yang et al., 2000a) and mammalian H-chain ferritin (Yang et al., 1998) under similar conditions, and in most cases the reaction is complete in under one minute. To determine whether hydrogen peroxide was evolved during the reaction, the use of an electrode capable of measuring H2O2 was attempted; however, the addition of Fe(II) to the protein solution caused a similar response in the electrode during the protein-free control experiment. Thus, the generation of H2O2 could not be measured directly. Instead, catalase was used to consume any hydrogen peroxide present, regenerating oxygen in the ratio 2 H2O2:1 O2 (Fig 3-9). Computing the mean drop in O2 after 5 replicates yielded an Fe:O2 ratio of 1.94 ± 0.22 and an Fe:H2O2 ratio of 3.04 ± 0.49 after 3 replicate measurements of oxygen regeneration following the addition of catalase. 57 Figure 3-9 Sample oxygen electrode trace showing O2 consumption following ferrous iron addition to apoferritin. A final concentration of 12 µM FeSO4•7H2O was added to 0.76 µM purified apoferritin in 2 mL of 50 mM MES buffer, 100 mM NaCl, pH 6.5 and oxygen consumption was monitored using a Clark type electrode (blue trace). When no further drop in O2 was observed, 1 µL catalase (20µg) was added such that the amount of H2O2 produced during the ferroxidase reaction could be assessed (using a ratio of 2:1 for H2O2:O2). The addition of 12 µM Fe(II) to buffer in the absence of protein serves as the control (pink trace). 12 µM Fe injection catalase added O2 consumed O2 regenerated 58 Chapter Four: Discussion 4.1 Behaviour of Recombinant Ferritin in Solution The solubility, stability and functionality of the mature multimeric ferritin protein generated for this study proved to be adequate to manipulate the protein during all experiments. The omission of the 6x HIS tag was one of the key factors leading to improvement of these three crucial properties, as its presence seemed to reduce protein stability. As the protein procured using the initial construct terminated part way through the 5th helix, residues reported to be involved in core formation were potentially omitted in this clone (Levi et al., 1988). Thus, the extension of the protein sequence by an additional 5 amino acids at the carboxyl terminus was likely beneficial, in that it may have imparted the ability to form a significant Fe core. Included in the added amino acids is a cysteine residue that is uncommon in other ferritins (Fig. 3-1), and may play a role in core formation or Fe release. Here, the extension of the construct at the C-terminus did not have the deleterious effects observed in E. coli BFR by Andrews et al. (1993). Instead, the addition of the residues likely increased the protein’s Fe binding potential from ~40 to over 600 Fe per protein shell. Residues 171-178 within the extended portion of the protein are predicted to encode a 6th helical region (Fig. 2-1). Although few ferritins include a sixth helix, the additional residues may be present in the endogenous Pseudo-nitzschia ferritin protein, given that they occur before the natural stop site after position 180 (Fig. 2-1). Further analyses of the protein after modifying the construct using site directed mutagenesis will be required to elucidate the function of the carboxyl terminus of this unique protein. 59 4.2 Spectroscopic Analysis During the titration of apoferritin with ferrous Fe, precipitate of Fe or protein was only noted near the upper limit of the determined binding capacity of the protein when Fe was added slowly; however, if additions were made in large increments, the appearance of precipitate was pronounced. It is thought that when Fe is added faster than it can be processed by the Fox sites, auto-oxidation of iron in the hydrophilic channels of the protein block further Fox activity, leading to the formation of precipitate especially above pH 7 or when Fe(III) is present (Yang et al., 1998; Yang and Chasteen, 1999). Auto- oxidation of Fe upon a preformed mineral core (crystal growth model) has also been reported above pH 7, and has a stimulatory effect on Fe binding, while at pH 6.5 and below, this effect is greatly reduced (Levi et al., 1988; Yang and Chasteen, 1999). Thus, to study the Fox activity in the absence of autocatalytic core formation while maintaining conditions near physiological pH, all assays were performed at pH 6.5. Baaghil et al. (2003) have also regarded these conditions to be optimal for the study of clonal H-chain ferritins. To study the Fox activity of P. multiseries ferritin, Fe(II) was added in small increments in the section of the spectroscopic titration curve below the discontinuity that differentiates ferroxidase activity from the slower mineral core formation (Fig. 3-8). Although the spontaneous oxidation of Fe(II) to Fe(III) in oxygenated solutions could potentially confound the assays performed here, the half life of Fe(II) is estimated to be on the order of 12 hours under conditions similar to those specified here (Maldonado et al., 2006). Since ferritin begins to bind Fe immediately, and the time between additions was on the order of minutes, it can be assumed that all Fe in solution is present as Fe(II) 60 or is bound to ferritin during the course of the assay, unless Fe is added above the binding capacity of the protein. Although ferritins principally take up Fe(II) from solution, Treffry and Harrison (1979) suggest that a relatively small but significant amount of Fe(III) finds its way into horse ferritin. If this were the case, a conjecture could be made that the presence of a small amount of Fe(III) may not be detrimental. Figure 3-8 shows a clear definition between the oxidation of Fe (phase 2) and its incorporation into the mineral core (phase 3), but the rapid initial binding of Fe(II) to the Fox site (phase 1) as discussed in Yang et al. (2000a) is not apparent. This is because to observe the release of a proton associated with Fe binding to the Fox site the experiment must be performed anaerobically such that oxidation of Fe does not occur; thus it was not measurable spectroscopically or using electrode oximetry employed here. Instead, the use of specialized spectroscopic techniques is generally required to detect the bond formation (Liu and Theil, 2004). At the beginning of the titration, the phase 2 molar absorptivity coefficient varied from 5700-6100 M-1 cm-1 per iron, and was greater than 3380 M-1 cm-1, the value determined for E. coli bacterioferritin by Yang et al. (2000a). However, as the reaction progressed, the absorptivity dropped, approaching the phase 3 molar absorptivity coefficient that stabilized around 2650 M-1 cm-1. The phase 3 absorptivity compares more closely with 2030 M-1 cm-1, the value determined for BFR core formation (Yang et al., 2000a). Yang et al. (2000a) compare their findings with the molar absorptivities published for human H-chain and horse spleen ferritins; the phase 2 values are 2990 and 3540 M-1 cm-1, and the phase 3 values are 2140 and 2285 M-1 cm-1, respectively. While the phase 3 molar absorptivity coefficient determined here is comparable to other 61 published values, it remains unclear why the phase 2 values were variable and significantly higher than expected. Perhaps the more important feature noted here is the reproducible shift in Fe binding between phases 2 and 3. Yang et al. (1998, 2000a) report that the discontinuity between the two phases exists at 48 Fe/protein for H. sapiens recombinant H ferritin and E. coli bacterioferritin, the point at which all ferroxidase sites (2 per monomeric subunit, 48 total) are saturated. Mineral core formation begins once a full complement of Fe is bound to the Fox sites (Le Brun et al., 1993), which may then proceed spontaneously via the crystal growth model if conditions permit. In the current study, given the proximity of the observed discontinuity of the spectroscopic titration curve to 24 Fe/protein, it is possible that the di-iron Fox sites are only half filled before core formation proceeds. The possibility that each Fox centre is a mono-iron site does exist, but is not parsimonious considering that ferritins are widely characterized as being di-iron carboxylate proteins. A more likely explanation is the possibility for some other metal, probably zinc (Zn), to be occupying some of the Fe binding sites. Although the Fox sites of the apoprotein were free of Fe after treatment with dithionite reducing agent and chelation by acetate, ferritin is known to preferentially bind Zn over Fe (Le Brun et al., 1995), and may have remained bound to about half of the di-iron sites, unaffected by reduction and chelation. In the literature, ferritins have been shown to contain Zn as purified, and Zn soaked ferritins have been created for crystallographic studies (Webb et al., 1985; Tatur et al., 2007). The addition of 48 Zn(II) to E. coli BFR prior to titration with ferrous Fe is known to fully inhibit Fox activity, with core formation being affected after the first 24 Zn are bound (Le Brun et al., 1995; Yang et al., 2000a; Baaghil et al., 2003); however, time constraints did 62 not permit the development of methods for Zn measurement or removal in this study. No trace metal analysis of the components of the E. coli growth medium (2YT, formulated from Bacto™ tryptone and yeast extract, BD Biosciences, Franklin Lakes, NJ, USA) could be located, and trace metal analysis was not performed, but for 24 Zn to be bound to the expressed ferritin, Zn content of the bulk medium would have had to be on the order of  9 µM. Although the discontinuity in the titration curve was lower than expected, it is clear that Fe was being bound effectively via the Fox active site, as indicated by the increase in absorbance at 295 nm and associated O2 consumption, as well as the inflection point that marked the transition to core formation. The growing peak at 295 nm and the fact that the phase 2 and 3 slopes are both linear when no precipitate is visible, suggests that protein is catalyzing the hydrolysis of Fe. Furthermore, since Fe measurements from spectroscopic analysis agreed well with the Ferene S® determination of iron content, adventitious binding of Fe to the exterior of the ferritin shell does not appear to be contributing significantly to the iron content of the recombinant ferritin. 4.3 Fe Binding Capacity The level of core formation in P. multiseries ferritin was somewhat lower than expected, yielding maximum Fe binding values of around 633 Fe/protein. In contrast, when maximally loaded, eukaryotic ferritins generally contain up to 4500 Fe/protein, bacterioferritins up to 2000 Fe/protein, and Dps ferritins up to 500 Fe/protein. The inability to load the protein with the maximum complement of Fe generally discussed in the literature could be due the fact that autooxidation of Fe did not occur at pH 6.5. 63 Alternatively, the in vitro Fe binding capacity of this clone is potentially lower than the endogenous protein in P. multiseries, the study of which could better reflect the true maximum under optimal conditions. Although the maximum binding capacity for P. multiseries ferritin remains unclear, it does bind more Fe per multimeric ferritin than the reported values for Dps ferritins, and further optimization of reaction conditions could potentially lead to greater Fe incorporation of recombinant ferritin. There are at least three potential factors related to the experimental conditions that could have imposed limits on the amount of Fe that can be bound by the ferritin protein. First, binding 633 µM Fe per µM ferritin shell would have required ~24 µM O2 for the phase 2 reaction (Fe:O2 = 2) and an additional ~152 µM O2 during core formation (Fe:O2 = 4), for a total of ~176 µM O2. Since the atmospheric oxygen saturation was calculated to be ~250 µM under the conditions employed here, up to 70% of the initial oxygen content of the buffer solution could have been consumed, leading to potentially suboxic conditions that may have limited the rate of Fe oxidation. However, it is unlikely that this level of oxygen depletion occurred, since between each addition of Fe the reaction solution was mixed vigorously. Alternatively, H2O2, which is produced during Fe loading of ferritin may have built up to levels that were detrimental to the stability of the protein. Since electrode oximetery was not performed for the core formation reaction, it is unknown whether this protein is able to eliminate net production of H2O2 as in E. coli BFR and L. innocua Dps ferritin (Yang et al., 2000a; Wiedenheft et al., 2005). Build up of H2O2 could have led to instability of the protein, causing premature precipitation prior to the completion of core 64 formation, and would account for the observed precipitate in the cuvette at higher levels of Fe per ferritin. A third potential explanation for suboptimal Fe binding involves the role of inorganic phosphate (Pi) in the formation and composition of the ferrihydrite core in ferritin proteins. There have been several studies that have examined the Fe:Pi ratios of horse spleen ferritin (Treffry and Harrison, 1978; Treffry et al., 1987), bacterioferritin (Treffry et al., 1987; Andrews, 1998; Aitken-Rogers et al., 2004), and most recently in a Dps ferritin (Castruita et al., 2006). These studies generally report that Fe:Pi is high in animal ferritins, about 8:1, while bacterial and plant ferritins contain more phosphate, with ratios ranging from 3:1 to 1:1. Bacterioferritins range from 2:1 to 1:1, and the Dps ferritin from T. erythraeum is intermediate in phosphate content with 4:1 Fe:Pi. The presence of phosphate during iron titration is reported to increase the rate of ferritin-catalyzed Fe(II) oxidation but stoichiometry associated with the Fox reaction remains unchanged (Aitken-Rogers et al., 2004); however, it remains unclear whether or not the maximum amount of bound Fe increases in the presence of phosphate. Johnson et al. (1999) also note faster phase 3 kinetics, and propose that a layer of Pi floats on the surface of the Fe core, expanding and contracting with it as it changes in size. Treffry et al. (1978) argue that Pi is incorporated but not required for core formation, reporting that much of the Pi is only weakly bound to horse ferritin, probably on the outside of the shell where it is easily displaced. Furthermore, the case has been made for autocatalytic core formation in the absence of Pi in mammalian ferritins (Yang et al., 1998). While no phosphate was added during the titrations performed in this study, a significant degree of 65 core formation was noted, although it is currently unknown if this would increase upon inclusion of Pi in the assay buffer. 4.4 Ferroxidase Fe/O2 and Fe/H2O2 Stoichiometry Addition of <18 Fe(II) per P. multiseries ferritin shell led to the phase 2 oxygen consumption ratio of ~2 Fe(II)/O2, identical to the values generally determined for human H-chain and horse spleen ferritins (Xu and Chasteen, 1991; Yang et al., 1998) as well as E. coli FTN (Treffry et al., 1998). While the properties of core formation are not presented in this study, preliminary findings did show an increasing Fe(II)/O2 ratio (associated with a reduction in the consumption of O2) when Fe was added in excess of the phase 2 to phase 3 transition point (~22.4 Fe/ferritin, Fig. 3-8). It is likely that Fe(II)/O2 would have increased to 4 during core formation, in keeping with the findings of Xu and Chasteen (1991) and Yang et al. (1998). The addition of catalase following the measurement of O2 consumption showed that hydrogen peroxide is generated during the ferroxidation reaction, as in human H- chain and horse spleen ferritins (Xu and Chasteen, 1991; Yang et al., 1998). However, the Fe(II):H2O2 ratio observed in this study was slightly higher, ~3 compared to the ratio of 2 expected from the mechanism proposed by Yang et al. (1998). In EcFTN, production of H2O2 is likely but has not yet been confirmed, although it is known that no net production of H2O2 is generally associated with BFR or Dps ferritins. If H2O2 is consumed during Fe binding in P. multiseries ferritin, either at a proximal Fox site or via the Fe catalyzed Haber-Weiss reaction in the assay solution, the increased Fe:H2O2 ratio could be explained. 66 4.5 Classification of P. multiseries Ferritin Regarding the physical structure of the ferritin protein cloned in this study, its monomeric subunit (20.0 kDa) is intermediate in size between 18.5 kDa BFR (Andrews, 1998) and 21 kDa mammalian H-chain ferritins (Levi et al., 1988; Orino et al., 1997). The fully formed shell is larger, however, more akin to plant ferritins with globular protein sizes of over 500 kDa (Ko et al., 1987; Laulhère et al., 1988). The larger MW of plant ferritins is due to their larger subunits, usually 25-28 kDa (Laulhère et al., 1988; Zancani et al., 2004); interestingly, P. multiseries ferritin seems to be of a similar size though with a significantly smaller subunit. It is possible that this protein folds such that the size of the central core and outer diameter are larger in this ferritin, but this would have to be confirmed with electron microscopy or by solving the crystal structure. From the O2 and H2O2 data presented in this study, it appears that P. multiseries ferritin most closely follows the phase 2 (ferroxidation) and phase 3 (core formation) stoichiometry of eukaryotic ferritins, consuming oxygen and producing some net hydrogen peroxide: Ferroxidation (phase 2):        [Fe(II)2-P] + O2 + 3H2O  [Fe(III)2O(OH)2-P] + H2O2 + 2H+ Core formation (phase 3):    [Fe(III)2O(OH)2-P] + H2O  2FeO(OH)(core) + P + 2H+ Although no proposed reactions from the group of prokaryotic non-heme homopolymer FTNs could be located from a comprehensive literature review, it seems probable that 67 they would also share a chemistry somewhat similar to H-chain ferritins given their structural similarities and phylogenetic proximity to the P. multiseries protein (Figs. 3-2, 3-3). Still, the reason for the elevated Fe:H2O2 ratio measured here remains uncertain, and could indicate that BFR type chemistry is occurring in P. multiseries ferritin, whereby some of the hydrogen peroxide produced during Fe binding may be somehow used up, either by the protein or via reactions within the assay solution. Based on the combination of sequence identity (Fig. 3-1), phylogenetic trees (Figs. 3-2, 3-3), and the properties of P. multiseries ferritin, it seems that the protein belongs to the class of non-heme class of prokaryotic ferritins formed from 24 identical subunits, in particular, those found in cyanobacterial genera. Its next closest relatives include other prokaryotes, in particular thermophilic archaea and bacteria. Thus, the P. multiseries ferritin (PmFTN) studied here can be classified as a non-heme homopolymer ferritin similar to those found in prokaryotes, but that it appears in a eukaryote is novel. 4.6 Evolution of Ferritin in the Marine Environment A current view suggests that the ancient lineage of marine cyanobacteria evolved from thermophilic bacteria and archaea in the Proterozoic ocean, a reducing environment rich in metal-sulfides and ferrous iron (Saito et al., 2003). The primary endosymbiosis of a cyanobacterial cell engulfed by a heterotrophic eukaryote protist was then thought to give rise to the red algae, phylum Rhodophyta (Li et al., 2006). Under this theory, a secondary endosymbiosis event between a red alga and a heterotrophic eukaryote gave rise to the chromalveolates, a lineage that includes the diatoms (Nisbet et al., 2004; Li et al., 2006). After each endosymbiosis event, most plastid genes were relocated to the 68 nucleus, away from oxidative stress, where recombination reduces the impact of accumulated mutations and improves fitness through genetic diversity (Martin, 2003). No putative ferritin genes could be located in any of the chromalveolate genera that were searched, and comprehensive testing of 23 diatoms indicates that only pennate diatoms, not centric diatoms possess ferritin genes (Marchetti et al., in prep). From the phylogenetic analysis performed here (Figs. 3-1 and 3-2), it seems that PmFTN is more similar to prokaryotic ferritins than to those of eukaryotes. This suggests that the ferritin gene of the pennate diatoms could have been acquired via lateral gene transfer from a cyanobacterium, rather than by the classical route of secondary endosymbiosis that is thought to have originally given rise to the diatoms. In this scenario, the gene encoding ferritin could have been incorporated directly into the diatom nucleus, after secondary red algal endosymbiosis occurred. The fact that the ferritin of the red alga C. merolae branches with other eukaryotes such as green algae and higher plants, but not the diatoms (Fig. 3-2), seems to lend support to the secondary endosymbiosis hypothesis, but suggests that the ferritin of pennate diatoms followed a unique evolutionary trajectory. It has been suggested that the evolution of eukaryotic plastid and nuclear symbiotic components may have occurred by unique pathways, and that the potential loss of plastid membranes could have confounded prior evolutionary reconstruction (Stiller and Hall, 1997). The phylogenetic analysis and protein characterization conducted in this study supports this rationale. The question remains, why do members of pennate diatom genera such as Pseudo-nitzschia and Phaeodactylum possess such dissimilar ferritins (sharing only 27.9% identity, Fig. 3-1), while their genetically closest relatives (namely chromalveolates including centric diatoms) do not possess ferritin at all? With the 69 emergence of new genome sequencing projects and the further study of HNLC regions of the ocean, new findings may help to provide some explanation. Given that cyanobacteria are thought to have evolved in an Fe rich environment, a clear selective pressure existed in favour of species with the ability to produce ferritin. Even now, coastal cyanobacteria of the genus Synechococcus retain superior trace metal storage and utilization ability, especially with respect to Fe and Cu (Palenik et al., 2006). However, the body of knowledge pertaining to ferritin in the open ocean is limited; even the strain of Pseudo-nitzschia used to clone ferritin in this study was isolated from a coastal habitat, although a number of other pennate diatoms including oceanic strains are now known to possess ferritin as well (Marchetti et al., in prep). Thus, the potential for ferritin-like proteins to assist marine phytoplankton, particularly pennate diatoms, to capitalize on transient Fe inputs is only now gaining attention. Interestingly, however, diatoms in HNLC areas are not the only algae that might benefit from the use of Fe storage proteins. Members of the genus Trichodesmium that contain Dps ferritins (Castruita et al., 2006) may also prosper, though for slightly different reasons. It is known that the Fe demand of such nitrogen-fixing organisms is high, due in part to the enzyme nitrogenase from which Trichodesmium derives its diazotrophic ability (Raven, 1988). A recent study showed that this cyanobacterium experiences phosphorus limitation in the central Atlantic (Sañudo-Willhelmy et al., 2001), and if its ferritin requires Fe:Pi in a 4:1 ratio as Castruita et al. (2006) suggest, an adaptational trade-off may exist. Ecologically, ferritin is also important to species of phytoplankton that do not express the protein themselves. Castruita et al. (2008) have determined that recycled Dps ferritin from Trichodesmium erythraeum can be utilized as the sole source of Fe by other 70 prokaryotic and eukaryotic phytoplankton. Thus, the role of ferritin in the marine system is more complex than first thought; it has implications for carbon sequestration, nutrient cycling and primary productivity in the ocean. It could also help to account for, or perhaps predict the levels of domoic acid production based on the Fe nutritional status of harmful blooms of Pseudo-nitzschia sp. in coastal areas. Subsequent studies using the framework developed here will be able to determine the occurrence and properties of ferritins found in the open ocean system. The development of a polyclonal antibody using PmFTN is in progress, and this will no doubt prove to be a powerful tool in the discovery of new ferritins in HNLC species during future Fe fertilization experiments. 71 Chapter Five: Conclusions The cloning and expression of ferritin from P. multiseries was successful, and the behaviour of the protein was greatly improved by omitting the N-terminal 6x HIS tag and extending the C-terminus slightly. In the cloning process, hybrid primers were used to amplify the putative ferritin gene using a PCR based, ligase independent methodology – an efficient and powerful technique that is gaining popularity. The methodology employed here can be adapted to clone virtually any ferritin (or other protein), simply by modifying the portion of the hybrid primers that is complimentary to the ends of the gene sequence of interest, and with minimal optimization of the PCR conditions. The ability of the cloned ferritin subunit to self assemble was crucial for the success of this work, and facilitated proper protein folding that created a functional enzyme. The heat stability of the fully formed ferritin shells is a universal property of ferritins, and was instrumental in the high yield of purified protein recovered during purification by anion exchange chromatography. In characterizing PmFTN, it was apparent that the 24 subunit homopolymer exhibits Fox activity, being able to bind and oxidize Fe(II) at rates comparable to other known ferritins. While the determination of the stoichiometery of diferrous binding at the 24 theoretical Fox sites may have been confounded by the presence of Zn, a metal known to inhibit Fe binding in ferritin, it is clear that this protein demonstrates both phase 2 ferroxidation and phase 3 core formation that leads to the storage of a significant number of Fe atoms. The fact that oxygen is consumed in a 2:1 Fe:O2 ratio, and hydrogen peroxide is produced in the Fox reaction indicates that the chemical nature of PmFTN is similar to clonal eukaryotic H-chain ferritins. However, phylogenetic analysis indicates 72 that the pennate diatom ferritin is closely related to cyanobacterial ferritin, which, along with bacteria and archaea, embody the class of non-heme ferritins, and that the gene was acquired laterally from a prokaryotic ancestor. It is also clear that PmFTN is structurally and functionally unique from bacterioferritins, Dps ferritins and heteropolymer mammalian ferritins. 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