Iron-storage mechanism of ferritin
Kourosh Honarmand Ebrahimi
2009-2013
Cover design: Kourosh Honarmand Ebrahimi Printed & published by: Uitgeverij BOXPress, ‘s-Hertogenbosch
Iron-storage mechanism of ferritin
PROEFSCHRIFT
ter verkrijging van de graad van doctor aan de Technische Universiteit Delft,
op gezag van de Rector Magnificus prof. ir. K.C.A.M. Luyben, voorzitter van het College voor Promoties,
in het openbaar te verdedigen op woensdag 1 mei om 15:00 uur door
Kourosh HONARMAND EBRAHIMI
Master of Science in Biochemical EngineeringProf. Dr. W. R. Hagen
Copromotor Dr. P-L. Hagedoorn
Samenstelling promotiecommissie:
Rector Magnificus, Voorzitter
Prof. Dr. W. R. Hagen Technische Universiteit Delft, promotor Dr. P-L. Hagedoorn Technische Universiteit Delft, copromotor Prof. Dr. Ir. A. J. M. Stams Wageningen Universiteit
Prof. Dr. P. D. E. M. Verhaert Technische Universiteit Delft Prof. Dr. S. de Vries Technische Universiteit Delft,
Dr. E. Bill Max Planck Institut für Chemische Energiekonversion, Mülheim, Duitsland
Dr. M. E. Than Leibniz-Institut für Altersforschung, Fritz-Lipmann Institut, Jena, Duitsland
Prof. Dr. J. H. de Winde Technische Universiteit Delft, reservelid
This work was financially supported by a research grant from the Dutch National Research School Combination-Catalysis Controlled by Chemical Design (NRSC-C).
Abstract and outline of the thesis 1
Chapter One, General Introduction 3
Chapter Two, Catalytic oxidation of Fe(II), the ferroxidase activity. Does
β-amyloid precursor protein (APP) have iron oxidation activity? 13 Chapter Three, Catalysis of iron core formation in Pyrococcus furiosus ferritin 31 Chapter Four, Inhibition and stimulation of formation of the ferroxidase center
and iron core in Pyrococcus furiosus ferritin 49
Chapter Five, The catalytic center of ferritin regulates iron storage via
Fe(II)-Fe(III) displacement 67
Chapter Six, A novel mechanism of iron-core formation by Pyrococcus furiosus archaeoferritin, a member of an uncharacterized branch of the ferritin-like superfamily
109
Chapter Seven, Engineering ferritin towards alkane activation 131
Chapter Eight, Conclusions and perspectives 139
References 141 List of abbreviations 153 Appendix 154 Acknowledgment 159 List of publications 161 Curriculum vitae 162
Abstract and outline of the thesis
torage of Fe(III) is a common mechanism by which the cellular machinery controls the availability of Fe(II) and Fe(III) for biosynthesis of iron-containing cofactors of enzymes which are involved in several essential biological processes, including oxidative phosphorylation. The conserved 24-meric iron-storage protein ferritin has been identified in many organisms to control the availability of Fe(II) by oxidizing the excess Fe(II) and storing the Fe(III) oxidation product in a soluble and nontoxic form. A conserved diiron center, the ferroxidase center, is responsible for catalytic oxidation of Fe(II), the ferroxidase reaction. The mechanism by which ferritin stores the Fe(III) is not fully understood, and the current models in the literature suggest different mechanisms for the functioning of ferritins from different Domains of life. Moreover, a structural gene for a 24-meric ferritin has not been found in some organisms including Pyrococcus
abyssi or Pyrococcus horikoshii. Below we first outline methods which can be used to
measure ferroxidase activity of different proteins. As an example we measure the ferroxidase activity of two proteins, human H ferritin and ceruloplasmin, and that of a synthetic peptide. Subsequently, using these techniques we study the mechanism of iron oxidation of a ferritin from hyperthermophilic archaeal anaerobe Pyrococcus furiosus. We then employ new experimental approaches using isotopically labeled 57Fe(II) to compare the iron-storage mechanism of P. furiosus ferritin with that of eukaryotic human H ferritin. We demonstrate that, conflicting with the current models in the literature these proteins employ a common mechanism to store the Fe(III) oxidation product. We suggest that this mechanism is general from archaea to eukaryotes. Finally, we carry out the in-vitro biochemical characterization of a new member of the ferritin superfamily of proteins that unlike the 24-meric ferritin is monomeric in the absence of iron. We name this protein archaeoferritin and we show that monomers oxidize Fe(II) and reversibly assemble to form Fe(III)-storing oligomeric structures comparable to that of ferritin.
Chapter One : General introduction. In this chapter the importance of iron for the functioning of different enzymes is outlined. Then, we explain the structural features of the ubiquitous iron-storage protein ferritin that has an essential role in controlling the concentration of available intracellular Fe(II) and Fe(III).
Chapter Two : Catalytic oxidation of Fe(II), the ferroxidase activity. This chapter is an introduction to the ferroxidase activity and to proteins that possess this activity. As an experimental example the ferroxidase activity is studied for two proteins, i.e. ceruloplasmin, and human H ferritin, and the results are compared with those of a peptide mimetic of the putative ferroxidase site of amyloid precursor protein (APP). We compare different methods that can be used to measure the ferroxidase activity of proteins.
Chapter Three : Mechanism of iron-storage by Pyrococcus furiosus ferritin. In this chapter steady-state kinetics of iron oxidation by P. furiosus ferritin (PfFtn) are studied. Two new possible mechanisms are proposed: (i) the ferroxidase center acts as a cofactor center; alternatively, (ii) the Fe(III) in the ferroxidase center is displaced by Fe(II).
Chapter Four : Inhibition of iron-oxidation by Zn(II). The inhibitory effect of zinc and acceleratory effect of oxoanions such as phosphate on the rate of Fe(II) oxidation by PfFtn is studied to gain more detailed insight in the ferroxidase center functioning.
Chapter Five : The catalytic center of ferritin regulates iron storage via Fe(II)-Fe(III) displacement. In this chapter we study the mechanism of iron-storage of PfFtn and HuHF in parallel, and we show that the Fe(III) in the ferroxidase center of both proteins is metastable. The Fe(III) in the ferroxidase center is displaced by Fe(II).
Chapter Six : A new iron-storage protein from Pyrococcus furiosus. We describe the in-vitro biochemical characterization of a new member of the ferritin superfamily of proteins which can also oxidize Fe(II) and then store the Fe(III)-oxidation product by subunit polymerization.
Chapter Seven: Engineering ferritin towards alkane activation. In this chapter we describe our attempts to change the catalytic activity of the ferroxidase center in order to catalyze oxidation of alkanes.
Chapter One
1. Iron and its importance for biological reactions.Life as we know it today is based on a limited set of essential elements (Figure 1.1). Among these elements metals play many important biological functions (Table 1.1): they are involved in catalysis of biological reactions, they stabilize the structure of biological molecules such as DNA, RNA, or proteins, and they play important roles in signal transduction. A key factor that determines whether a metal may have biological importance is its bioavailability in the habitat of a specific organism. The metals that are useful for life should be well distributed and well soluble in the biological solvent, water (2).
Figure 1.1. Schematic representation of the periodic table of elements. The most common elements that
are essential for organisms to sustain their life are shown in yellow and red. The red color denotes transition metals which are known to be important for the molecular machinery of living organisms.
The essential properties of a metal ion that defines whether it can act as a catalyst in the catalytic center of proteins are its ability to reversibly accept or donate electrons (or partial charge), to form stable complexes with its coordinating ligands in its biological containers, proteins, and to be able to bind to a substrate and convert it into a product. The metal ions that are involved in catalysis have a rate of hydrolysis (the rate of substitution of a water ligand with another water ligand) of ca. 104 to 108 s-1. Metal ions that have a hydrolysis rate faster than 108 s-1 are not used in catalysis because the life-time of the substrate-metal complex will be too short for catalysis to occur. On the other hand, metal ions that have a hydrolysis rate slower than 104 s-1 are not used because uptake and intracellular transport of these metals take way too long as the coordinating water cannot easily be displaced by the coordinating amino acid residues of a protein. The metal ions that we know to catalyze biological reactions as cofactor in the catalytic center of enzymes are listed in Table 1.2. Among these metals, iron is perhaps the mostly used element for catalysis of many biological reactions including, nitrogen fixation (3), photosynthesis (4), oxidative phosphorylation, conversion of methane to methanol (5), and synthesis of deoxyribonucleotides (6) which are the building blocks of DNA. Iron that controls the catalysis of these reactions has been found in several forms as a cofactor that either directly attends the catalysis or acts as an electron path to transfer electrons to/from the catalytic site. The known iron cofactors and prosthetic groups can be divided into five groups: mononuclear Fe2+/Fe3+ centers (7), diiron centers (8), heme prosthetic groups, iron-sulfur clusters (9), and cofactors that consist of iron and another metal such as NiFe cofactor of hydrogenase.
Table 1.1. Biological role of some metals. Biological function Metal ion Structural roles Mg2+, Zn2+ Signal transduction Ca2+, K+, Na+
*Redox reactions Fe2+/Fe3+, Mn2+/Mn3+, Cu1+/Cu2+
Mg2+ and Zn2+ are not redox active metals but they can act as Lewis acid catalyst.
Table 1.2. Metals involved in biological catalysis.
Metal An example of cofactor and enzyme ref
Iron Diiron cofactor; methane monoxygenase (MMO) (8)
Copper Copper/Zinc cofactor; superoxide dismutase (SOD1). (10)
Manganese Manganese cofactor, SOD2. (11)
Molybdate Molybdopterin cofactor; nitrate reductase (12) Tungstate Tungstopterin cofactor, formate dehydrogenase (13) Cobalt Dicobalt cofactor; methionine aminopeptidase (14)
Nickel NiFe cofactor, Hydrogenase (15)
Vanadium Vanadate cofactor, Vanadate-dependent haloperoxidase (2) *Zinc Copper/Zinc cofactor, superoxide dismutase (11)
* Zn(II) is not a redox active metal. In copper/zinc superoxide dismutase the Zn(II) in the catalytic center has a structural role (10). In hydrolases (e.g. carboxypeptidase) Zn(II) is a Lewis acid catalyst.
Iron is the fourth most abundant element in the earth crust and has been spread in all biological habitats. It is mainly found in two oxidation states, ferrous (Fe2+) and ferric (Fe3+). Iron can assume different electronic configurations, i.e. high spin S=5/2 or low spin S=1/2 (and occasionally intermediate spin S=3/2). The spin state reflects the distribution of electrons in d orbitals. For example a high spin ferric species has five unpaired electron in its d orbitals, while a low spin ferric species has one unpaired electron in only one of its d orbitals. With the advent of an oxygenic atmosphere the usage of iron has become a problem for the cell. Fe2+ is soluble at physiological pH but in the presence of molecular oxygen it is oxidized to Fe3+ which is highly insoluble at this pH and precipitates as iron oxide species. Besides the insoluble Fe3+ the reaction of Fe2+ with molecular oxygen produces reactive radicals such as superoxide (O2•‾) and hydroxyl radicals (OH•). These reactive radicals are commonly known as reactive oxygen species (ROS). ROS attack many components of the cell and irreversibly destroy them and thus may initiate series of events that lead to programmed cell death, apoptosis (16). Therefore, it is vital for the molecular machinery of the living organisms to
strictly control iron trafficking. Iron homeostasis in the cell is controlled by several proteins that are involved in import, export, storage, and transport of iron, and biosynthesis of iron cofactor centers. Some of these processes in prokaryotes and eukaryotes are controlled differently while some others are conserved. A conserved process that has been found in all life forms is storage of toxic free Fe(II) in the form of Fe(III) by the ubiquitous iron-storage protein ferritin.
2. Ferritin. Ferritin consists of 24 subunits that form a spherical-shaped structure with 432 symmetry (Figure 1.2a). The Fe(III) is stored inside the cavity of the protein as ferrihydrite-like mineral core. Each subunit of ferritin consists of five helices which are named helix A, helix B, helix C, helix D, and a short C-terminal helix, helix E (Figure 1.2b).
Figure 1.2. Structure of ferritin. (a) The conserved quaternary structure of the 24-meric ferritin (PDB
1MFR). (b) The tertiary structure of a subunit. Each subunit consists of five helices that are shown in the figure with different colors (PDB 2FG4).
2.1. Vertebrate ferritin. In vertebrates generally two subunits of ferritin have been found which have been named based on their molecular weight: H “heavy” with a molecular weight of ca. 21 kDa and L “light” with a molecular weight of 19 kDa. The two subunits assemble in different ratios to form isoferritins of 24-subunits. The ratio of H and L subunits in mammalian ferritin is tissue dependent. In tissues such as liver and spleen the number of L subunits is greater than that of H subunits, while in tissues such as heart and pancreas the number of H subunits increases (17,18). The H subunit can catalytically oxidize Fe(II) using a diiron catalytic center, the ferroxidase center (FC) (Figure 1.3a), which is located in the middle of its four α-helix bundle. A third structural gene that encodes a subunit with molecular weight between H and L, i.e. M ‘middle’ subunit, has been found in bullfrog (19). The M subunit has a diiron binding site similar to the H subunit. The physiological role of the M subunit and why bullfrog encodes two catalytically active ferritins, i.e. M and H subunits, is not known.
Figure 1.3. (a) The location of the conserved diiron catalytic center in the middle of four α-helical
bundles of ferritin (PDB 2JD7). (b) The location of heme in the interface of two subunits in bacterioferritin (PDB 3E1M).
The H subunit appears to regulate the overall balance of iron in specific tissues (20,21). It is involved in rapid oxidation of iron (22,23),and deletion of the H subunit ferritin gene from mouse embryonic cells has been shown to be lethal (24). Down-regulation of H ferritin using siRNA suggested that suppression of H chain ferritin decreases the cellular resistance to the presence of excess amount of iron (25). These studies together suggest that the H subunit directly participates in the iron homeostasis and detoxification. In contrast, the L subunit appears to be required for storage of the Fe(III) rather than oxidation of Fe(II). Higher ratios of L to H subunit prevents formation of protein aggregates after addition of large amount of Fe(II), i.e. more than circa 1000 Fe(II) per 24-mer (22,25-27). These results suggest that L ferritin is involved in efficient storage of Fe(III).
2.2. Bacterial and archaeal ferritin. In bacteria two different forms of ferritin have been found: a non-heme ferritin and a heme containing ferritin which is commonly known as bacterioferritin. The non-heme ferritin of bacteria is similar to the H subunit of vertebrates and it has the dinuclear iron binding site. The subunits of bacterioferritin are similar to that of non-heme ferritins, but in the quaternary structure of bacterioferritin a non-heme group is located in the interface between each pair of subunits with a methionine from each subunit as axial ligands (Figure 1.3b). 24 identical subunits assemble to form the quaternary structure of the bacterial non-heme ferritins and heme-containing bacterioferritins. The function of heme in bacterioferritin is not known. A site-directed mutant of E. coli bacterioferritin that lacks the heme group, i.e., the Met25His variant, was normally assembled and has essentially the same Fe(II) oxidation activity as that of the wild type protein. Therefore, it has been concluded that the heme group is not essential for the assembly of the 24-mer and iron oxidation by bacterioferritin (28). Very recently Yasmin et al (29) have suggested a new role for heme in
EcBFR. Using kinetics studies of wild-type bacterioferritin and heme-free variants it has been suggested that heme facilitates release of iron from EcBFR. Moreover, recent X-ray crystallographic studies have shown that the [2Fe-2S] cluster of bacterioferritin-associated ferredoxin can reduce the Fe(III) mineral core in Pseudomonas aeruginosa bacterioferritin (Pa-BfrB) via an electron transfer mechanism that is dependent on the heme group in Pa-BfrB (30).
Archaeal genomes appear to have only one type of ferritin gene. There are also exceptions in which the ferritin gene has not been found: e.g. Pyrococcus horikoshii and Pyrococcus abyssi. Two archaeal ferritins have been isolated and their X-ray crystal structures have been determined: Pyrococcus furiosus ferritin (PfFtn) (1), and Archeaoglobus fulgidus ferritin (AfFtn) (31). The subunits of these ferritins have a high degree of identity with that of non-heme ferritins of bacteria and with the H subunit of vertebrate ferritins and they possess Fe(II) oxidation activity comparable to those of bacterial ferritin and H subunit of vertebrate ferritin. Like in bacterial ferritins, all subunits of the 24-meric archaeal ferritins are identical.
3. Structural-functional features of ferritin
3.1. The diiron catalytic center, the ferroxidase center. Kinetics of Fe(II) oxidation by horse spleen ferritin (HoSF) in the presence of the Fe(III) transport protein transferrin showed that the Fe(III) product of the ferroxidase reaction was scavenged by transferrin (32). Based on this observation it was predicted that the catalytic centers in ferritin that oxidize Fe(II) to Fe(III) are near the surface of the protein. Further studies with recombinant human H-chain ferritin (HuHF), recombinant human L-chain ferritin (HuLF), and human liver ferritin which is a heteropolymer of H and L subunits showed that the ferroxidase activity of H-chain is considerably higher than that of L-chain (22). These observations suggested that there are specific sites in H-chain ferritin which facilitate catalysis of Fe(II) oxidation. Subsequently, the X-ray crystal structure of HuHF in the presence of Tb(III) was solved and two Tb(III) binding sites in the middle of the four α-helix bundle, site A and site B, were found (33). Comparison of the amino acid sequence of H- and L-chain ferritin showed that the metal binding residues of the two sites were absent in L-chain ferritin (34). Therefore, these sites are possibly the catalytic centers involved in fast oxidation of Fe(II) by H-chain ferritin. The data obtained from the X-ray crystal structure together with the observation that mutation of the residues of the dimetal binding site effectively blocked the ferroxidase activity of HuHF (35,36) strongly suggested that this site is involved in initial catalytic oxidation of Fe(II). Figure 1.4 shows the residues of the ferroxidase center of HuHF.
Since the identification of the ferroxidase center in human H ferritin, the X-ray crystal structures of ferritins from different organisms, e.g. E. coli ferritin A (EcFtaA) (37), Pyrococcus furiosus ferritin (PfFtn) (1), Pseudomonas aeruginosa ferritin (PaFtn) (38), bullfrong M ferritin (BfMF) (19), and Helicobacter pylori ferritin (39), have been solved in the
presence of different metal ions, e.g. Zn(II), Co(II), or the Fe(II) substrate. In all these structures the diiron catalytic center has been identified.
3.2. Threefold channels. The quaternary structure of ferritin has at least two known channels: eight threefold channels and six fourfold channels. Threefold channels have a funnel shape and are more hydrophilic (Figure 1.5). Initial studies using NMR spectroscopy and spin labeling of a cysteine residue of the threefold channel in horse spleen ferritin (HoSF) (40) suggested that threefold channels are possible entry routes of Fe(II) to the ferritin. Mutational analysis of the threefold channels showed that mutation of the hydrophilic residues of these channels in recombinant H-chain ferritin does not affect iron oxidation (41). Subsequently, using spin labeling, and EPR spectroscopy the threefold channels were proposed as a route for entry of Fe(II) (42). These initial studies were followed by several studies using site directed mutagenesis, inhibition studies, electrostatic calculations, and X-ray crystallography which confirmed that threefold channels are responsible for Fe(II) entry into the protein (43-49). The threefold channels in bacterial and archaeal ferritin are more hydrophilic than those in eukaryotic ferritin.
3.3 Fourfold channels. Six fourfold channels have been identified in the structure of all ferritins. The structure of a fourfold channel of PfFtn is shown in Figure 1.6. Mutation of the residues near the fourfold symmetry axis has suggested that the presence of these channels is essential for core formation in HuHF (41). The small E-helix near the fourfold symmetry axis
Figure 1.4. Ferroxidase center of
HuHF. The ferroxidase center consists of two iron binding sites which are referred to as FeA and FeB site (PDB
1FHA).
appears to be important for handling of iron. Disruption of the E-helix at the fourfold channels of human L ferritin caused mishandling of iron by the mutant compared to the wild type (50,51) and appears to be the cause of neuroferritinopathy: a neurodegenerative disease characterized by abnormal accumulation of iron in the brain. Calculation of the electrostatic free energy that is required to transport ions through threefold and fourfold channels of ferritin (L subunit) using the Poisson-Boltzmann equation suggested that in mammalian ferritin the fourfold channels may act as proton transport channel (45). In the case of bacterioferritin the fourfold channel is more hydrophilic. An X-ray crystal structure analysis of Azotobacter vinelandii bacterioferritin (AvBF) has shown iron binding to fourfold channels (52). This observation, in combination with the observation that at intermediate and high iron loading the residues of the fourfold channel in Helicobacter pylori ferritin undergo considerable conformational changes (39), have led to the proposal that fourfold channels in bacterioferritin are involved in iron transport through the protein shell.
Figure 1.5. Threefold channels of PfFtn. (a) The funnel shape of the threefold channels. (b) Amino acid
Figure 1.6. Fourfold channels of PfFtn. (a) Shape of the fourfold channels. (b) Residues of a fourfold
channel in PfFtn (PDB 2JD7).
In summary, although the X-ray crystal structure of several ferritins have been solved, our knowledge about how these proteins capture the Fe(II) substrate and store the resultant Fe(III) oxidation product is limited. It appears that first the Fe(II) enters the protein cavity via the threefold channels and subsequently reaches the ferroxidase center where it is oxidized. The exact route of Fe(II) to the ferroxidase center is not known. After oxidation of Fe(II), the resultant Fe(III)-oxidation product is somehow stored in the ferritin cavity. The residues of the E-helix in eukaryotic ferritin appear to be essential for proper storage of Fe(III) in these ferritins, but the role of these residues in the mechanism of Fe(III) storage is not known.
Chapter two
Catalytic oxidation of Fe(II), the ferroxidase activity.
Does β-amyloid precursor protein (APP) have iron
oxidation activity?
This chapter is adapted from:
Honarmand Ebrahimi K., Hagedoorn P. L., Hagen W. R., A Synthetic Peptide with the Putative Iron Binding Motif of Amyloid Precursor Protein (APP) Does Not Catalytically Oxidize Iron, (2012), PLoS ONE 7(8): e40287
Abstract
he β-amyloid precursor protein (APP), which is a key player in Alzheimer’s disease, was recently reported to possess an Fe(II) binding site within its E2 domain which exhibits ferroxidase activity [Duce et al. 2010, Cell 142: 857]. The putative ligands of this site were compared to those in the ferroxidase site of ferritin. The activity was indirectly measured using transferrin, which scavenges the Fe(III) product of the reaction. A 22-residue synthetic peptide, named FD1, with the putative ferroxidase site of APP, and the E2 domain of APP were each reported to exhibit 40% of the ferroxidase activity of APP and of ceruloplasmin. It was also claimed that the ferroxidase activity of APP is inhibited by Zn(II) just as in ferritin. We measured the ferroxidase activity indirectly (i) by the incorporation of the Fe(III) product of the ferroxidase reaction into transferrin and directly (ii) by monitoring consumption of the substrate molecular oxygen. The results with the FD1 peptide were compared to the established ferroxidase activities of human H-chain ferritin and of ceruloplasmin. For FD1 we observed no activity above the background of non-enzymatic Fe(II) oxidation by molecular oxygen. Zn(II) binds to transferrin and diminishes its Fe(III) incorporation capacity and rate but it does not specifically bind to a putative ferroxidase site of FD1. Based on these results, and on comparison of the putative ligands of the ferroxidase site of APP with those of ferritin, we conclude that the previously reported results for ferroxidase activity of FD1 and – by implication – of APP should be re-evaluated.
Introduction
Catalytic oxidation of Fe(II) to Fe(III) by molecular oxygen is commonly known as ferroxidase activity. The ferroxidase activity has been reported for several proteins including ceruloplasmin (53-55), frataxin (56-58), ferritin (32,59), Dps (DNA binding protein from starved cells) family of proteins (60-62), β-amyloid precursor protein (APP) (63) and rubrerythrin (64).
1. Ferritin. Ferritin is the ubiquitous iron storage protein in all forms of life. Ferritin consists of 24 subunits (Figure 2.1A) forming homopolymers in bacteria and archaea but heteropolymers in vertebrates. This subject was briefly discussed in the previous chapter and is discussed in detail in several review articles by others (65). Ferritin uses the ferroxidase activity to store iron in a ferrihydrite mineral form in a core. It is not known whether the Fe(III) that is stored in the ferritin can be used when it is required and how this iron is released to the solution. Each ferritin molecule is able to store approximately 3000 Fe(III) per molecule. The site of the ferroxidase reaction is located in the center of the four α-helical bundles of each catalytically active ferritin which is commonly known as the ferroxidase center (FC). The mechanism of the FC reaction will be discussed in more detail in the next chapter.
2. Dps protein. Dps and Dpr (Dps-like peroxide resistance protein) proteins have 12 subunits that form a hollow spherical protein with 23 symmetry (66) (Figure 2.1B). They show Fe(II) oxidation activity (60,62) and they can store iron in their internal cavity (62). It has been suggested that the function of Dps protein is protection of DNA from ROS (61,67,68). The ferroxidase activity of the Dps protein has been attributed to six diiron binding sites located at the interface between pairs of subunits of the protein (69,70). 3. Ceruloplasmin. Ceruloplasmin (Cp) is a serum protein and it contains ca. 95% of the
copper found in plasma (71). Ceruloplasmin is a member of the family of multi-copper oxidase enzymes. These proteins are identified by the presence of three spectroscopically distinct copper centers which are generally known as type I, type II, and type III (72,73)1. Ceruloplasmin contains three type I copper centers, a single type II copper center and two antiferromagnetically coupled type III coppers that together with the type II copper center form a trinuclear copper center (Figure 2.1C). This center is the place where oxygen binds and is reduced during catalysis. It has been shown that Cp has ferroxidase activity (53,55). The mechanism of ferroxidase activity of Cp and the physiological role of the ferroxidase activity of Cp is not known. A series of studies demonstrated that copper deficiency resulted in decrease of the level of Cp in the plasma and at the same time in 1 Type I copper centers have a deep blue color because of a Cys S‐Cu(II) charge transfer transition. They have a distorted tetrahedral coordination sphere. Type II copper centers, or non‐blue copper centers, are characterized by a square planar, or a deformed octahedral coordination sphere. Type III copper centers contain two antiferromagnetically coupled coppers.
accumulation of iron in the liver and other tissues. Administration of oxidized active ceruloplasmin resulted in release of iron as circulating Fe(III)-transferrin complex, thus the ferroxidase activity of Cp is suggested to be important in iron homoestasis (74,75).
Figure 2.1. X-ray crystal structure of proteins for which ferroxidase activity has been reported. (A)
Iron-storage protein ferritin (24 subunits, each subunit circa 20 kDa), (B) DNA binding protein from starved cells (Dps) (12 subunits each subunit circa 18 kDa), (C) ceruloplasmin (150 kDa), (D) frataxin (18 kDa), (E) E2 domain of β-amyloid precursor protein (APP) (60 kDa), and (F) rubrerythrin (20 kDa monomer).
4. Frataxin. Frataxin (Figure 2.1D) is a conserved protein among eukaryotes and prokaryotes. It is an important protein in iron homeostasis and its deficiency in human results in Friedreich’s ataxin (FRDA) a lethal neurodegenerative disorder (76). Several functions have been reported for this protein (77) but its exact role has not yet been understood. It has been suggested that frataxin interacts with aconitase, the Krebs cycle enzyme that converts citrate to isocitrate, and prevents the disassembly of the [4Fe-4S]2+ cluster of this enzyme and promotes the reactivation of aconitase activity (78). It has been shown that frataxin acts as iron chaperone to deliver Fe(II) to inactive [3Fe-4S]0 cluster of aconitase and converts it to the active [4Fe-4S]2+ form. More recently frataxin has been shown to function as an iron sensor that acts as regulator of Fe-S cluster formation. Frataxin possesses ferroxidase activity (56,57) and monomers of frataxin assemble to form oligomeric structures in order to store iron (58,79,80). The physiological relevance of the ferroxidase activity of frataxin is not known.
5. β-amyloid precursor protein (APP). In Alzheimer’s disease the amyloid β-protein, which is generated from the β-amyloid precursor protein (APP), oligomerizes and mediates synaptic failure (81). The APP gene consists of 19 exons which are alternatively
spliced predominantly into four different products that are named according to their length, namely, APP695, APP714, APP751, and the full length APP770 (82). The iron binding and transport protein transferrin that was originally used to indirectly measure the ferroxidase activity of ferritin (32) and ceruloplasmin (55) was used to measure possible ferroxidase activity of APP (63). It has been reported in ref (63) that APP has ferroxidase activity which is orders of magnitude higher than the activity of ferritin. The ferroxidase activity of APP has been assigned to a putative ferroxidase site in its E2 domain (Figure 2.1E and Figure 2.2).
6. Rubrerythrin. Rubrerythrin is a member of the ferritin superfamily (Figure 2.1F) and contains a diiron binding site and a rubredoxin-like iron binding site (two CxxC motifs coordinating a mononuclear iron). The role of rubrerythrin is not known but it has been shown that it has ferroxidase activity (83) and peroxidase activity (84).
The putative ferroxidase activity of APP. Human β-amyloid precursor protein (APP) is generally thought to play a key role in Alzheimer’s disease as the source of plaque-forming β-amyloid peptides (Aβ) (85,86). APP is a transmembrane protein made up of a large, multidomain extracellular extension, a small, single-pass transmembrane part, and a small intracellular extension (Figure 2.2A). Alternative exon splicing of the APP gene affords different mRNAs that translate into APP iso-forms whose length range from 365 to 770 amino acid residues (87), with APP695 as the dominant form in the brain. Sequential processing of APP by the proteolytic enzymes β-secretase and γ-secretase liberates Aβ: a sequence of typically 40 or 42 residues originally located partially in the membrane and for the remainder extracellularly. It is not known whether Aβ in Alzheimer’s disease is a causative agent or a resulting product. The physiological role(s) of APP in healthy cells has also not been firmly established. Metal ions, notably iron, copper, and zinc have been implicated in the normal physiological functioning of APP (88-90), in the regulation of APP expression (91,92), in the processing of APP affording Aβ (92,93), and in Aβ-plaque related pathogeny (89,94). The complete APP has not been crystallized, but the 3D structures of two extracellular domains, E1 and E2, have been determined.
Two subdomains of the N-terminal E1 domain, the ‘growth factor like domain’ GFD and the ‘copper binding domain’ CuBD, have separately been crystallized (95-97). In none of these four crystal structures any metal has been found. When the CuBD crystal is soaked in 100 mM CuCl2 a single site is found with Cu2+ square-pyramidal coordination by His147, His151, Tyr168 and two waters (96). A physiological role, if any, of this copper-binding site has yet to be established. Other putative copper (and zinc) binding sites have been proposed in E1 (89), and physiological studies have implicated copper in APP trafficking (88). An interaction of iron and the Aβ domain of APP has been reported (98) but a direct interaction with the E2 domain of APP has not been explicitly considered until the recent proposal that APP exhibits an Fe(II) binding site within the E2 domain (Figure 2.2B-C). It was proposed that this site has
homology with the ferroxidase center in the iron-storing ferritins (Figure 2.2D), and it was claimed that the site possesses catalytic iron oxidation activity (ferroxidase activity) (63). Steady-state kinetics of ferroxidase activity were reported for APP695, for the isolated E2 domain, for a synthetic 22 amino acid peptide named FD1 (duplicating a stretch in the E2 domain with the putative ferroxidase site), and for the human copper protein ceruloplasmin (as a positive control) (63).
Figure 2.2. Topology of β
amyloid precursor protein (APP) and comparison of its putative ferroxidase site with the ferroxidase site of human H-chain ferritin. (A) A schematic representation of APP and its metal binding domains (APP770 isoform).
(B) Structure of the E2 domain of APP751 (PDB 3NYL) and (C) the putative ligands of the ferroxidase site (APP770 numbering). The residues in green show the section of the E2 domain that contains the putative ferroxidase site of APP and the residues that were used to synthesize the FD1 peptide. (D) The diiron catalytic center, the ferroxidase center, of human H-chain ferritin (PDB 1FHA). The numbers in the parenthesis are based on APP770 numbering).
Ferroxidase activity of APP was proposed to be functionally relevant in interaction with the iron exporter ferroportin to prevent intracellular iron accumulation and associated oxidative stress. Like in ferritins (43,44,99,100), the APP ferroxidase activity was reported to be inhibited by binding of zinc to the APP, its E2 domain, or the FD1 peptide (63). The present study, which elaborates on the proposed ferroxidase activity of the FD1 peptide (63), was incited by five distinct aspects of the previous work: (i) the ferroxidase activity was measured with what appears to be an unvalidated procedure involving transfer of Fe(III) to the iron transport protein transferrin, and reported activities for ceruloplasmin, APP, the E2 domain of APP, and the FD1 peptide were claimed to be orders of magnitude higher than ferritin ferroxidase; (ii) reported Michaelis-Menten plots for ferroxidase activity of each of the four studied proteins had shapes that appeared to be incompatible with the hyperbolic shape of Michelis-Menten kinetics, and Vmax values that were read as saturation values from the data in figures differed by circa 40% from the values obtained from fitting the Michaelis-Menten equation to the non-hyperbolic data sets; (iii) the kcat value reported in the figures for the FD1 peptide was identical to the values reported for APP and ceruloplasmin, and it was not 40% of the kcat values reported for APP and ceruloplasmin as claimed in the text; (iv) reported ferroxidase activities of the E2 domain for two experiments performed under the same conditions differed by circa 50% while error bars of <1% were claimed; (v) the proposed homology between APP and human H-chain ferritin appeared to be based on a mistake of assignment of iron coordinating amino acids (63). We have therefore re-evaluated the transferrin-based ferroxidase assay by comparison with direct measurements of O2 consumption, and we have compared ferroxidase activity of the putative ferroxidase peptide FD1 from APP with those of ceruloplasmin and human H-chain ferritin.
Materials and Methods
Chemicals. All chemicals were reagent grade and were purchased from Sigma Aldrich. Human
ceruloplasmin (40 U/mg) was also obtained from Sigma Aldrich.
Expression and purification of human H chain ferritin. The Escherichia coli Top10 strain (Invitrogen)
was transformed with the plasmid containing the coding sequence for expression of human H chain ferritin (HuHF) pET12a. After two hours of growth to an OD600nm of 0.6, production of HuHF was induced by the addition of 0.1 mM IPTG. After eight hours cells were collected and were disrupted using a cell disruptor system (Constant systems). The supernatant was collected using centrifugation and was subjected to a heat step at 85 C for 10 min. Denatured proteins were separated using a centrifugation step. HuHF was made apo as described previously (101) and the final preparation contained less than one iron per 24-meric protein as determined with the ferene assay. The concentration of protein was measured using the bicinchoninic acid (BCA) assay using bovine serum albumin as standard.
Preparation of apo-transferrin. Apo-bovine transferrin (> 98% pure) was purchased from Sigma
Aldrich. The lyophilized powder was dissolved in 10 mM Mops, 100 mM NaCl, pH 7.0 and was dialyzed for at least 3 days against the same buffer using dialysis tube (Spectrumlabs) with a cut-off of 10 kDa. Subsequently consecutive dilution and concentration steps using an ultrafiltration membrane with a cut-off of 30 kDa (Millipore) were used to clean transferrin from possible metal complexing molecules which may have remained in the transferrin lyophilized powder from the manufacturing process and to concentrate the protein. The concentration of transferrin was measured using the bicinchoninic acid (BCA) assay and the purified apo-transferrin was directly used in experiments.
Preparation of the synthetic peptide (FD1). The peptide FD1 (90% pure, HPLC) was purchased from
GenScript. 4 mg of peptide was dissolved in 1 ml of 10 mM Hepes, 100 mM NaCl, pH 7.2, and dilutions were used directly in experiments. The sequence HRERMSQVMREWEEAERQAKNL was confirmed by low energy collision induced dissociation tandem mass spectrometry using a hybrid quadrupole time-of-flight mass spectrometer (Waters QTof Premier).
Preparation of ferrous sulphate solution. To prepare the Fe(II) solution the pH of Milli.Q water was set
to 2.5 using HCl. The solution was made anaerobic by flushing with high purity argon gas (99.999 %), and then it was added to ferrous sulfate salt in an anaerobic glove box (Coy Laboratory Products).
Steady state kinetics of Fe(II) oxidation in the presence of transferrin. Kinetics of Fe(II) oxidation
and Fe(III) uptake by transferrin were measured at 460 nm using a molar extinction coefficient for diferric transferrin of ε460nm= 4.56 mM-1cm-1 (32). Measurements were carried out on a fiber-optics spectrophotometer (Avantes) using 1 ml glass cuvettes with a path length of 1 cm. Measurements were done at 37 C in 100 mM Hepes, 100 mM NaCl, pH 7.2 unless otherwise stated. Progress curves were recorded for circa 5 minutes and initial rates were obtained from the slope of a line that was fitted to the data points for the first 50-100 seconds. Each experiment was repeated at least three times with two different batches of transferrin. For kinetic measurements to each cuvette the following additions were made in order (total volume of 1 ml): Hepes buffer, aliquot of transferrin between 50 to 100 µl to reach the desired final concentration, 3.5 µl of 1.4 mM FD1 peptide (as control experiment no peptide was added), and aliquots of FeSO4 solution, between 0.5 µl and 20 µl. To determine the inhibitory effect of zinc, aliquots of 0.5 µl to 20 µl of ZnSO4 solution were added 5 minutes before addition of FeSO4.
Oxygen consumption measurements. To measure consumption of oxygen a Clark electrode was used as
described before (102). The final reaction volume in the cell was 2 ml. To the cell the following was added in order: buffer, protein or peptide. Subsequently aliquots of 5 µl anaerobic solution of ferrous sulfate were added using a gastight syringe. The buffer was 100 mM Hepes, 100 mM NaCl, pH 7.2. Measurements were performed at 22 C.
Fast kinetics of Fe(II) oxidation. Fe(II) oxidation by human H chain ferritin was followed using a
Scientific PQ/SF-53 preparative quench/stopped-flow instrument with an EG&G Princeton Applied Research 1,024-element photodiode-array detector. The instrument was equipped with a nitrogen gas
flow on top of the sample cells to prevent oxidation of the Fe(II) solution before mixing with protein. The instrument was set to zero with rapid mixing of protein with anaerobic water, pH 2.4, before each measurement. The buffer was 400 mM Mops, 100 mM NaCl, pH 7.0.
Isothermal titration calorimetry. Zn(II) binding to transferrin was measured using isothermal titration
calorimetry with a VP-ITC microcalorimeter (GE healthcare). A control experiment in the absence of protein or peptide was performed to measure the heat generated due to Fe(II) or Zn(II) dilution in the buffer. The conditions for the control experiments were identical to the conditions for titration of Fe(II) or Zn(II) to transferrin or to FD1 peptide. For anaerobic Fe(II) binding to FD1, the ITC machine was placed in a polyethylene bag (Atmosbag, Sigma Aldrich). The bag was made anaerobic by high purity argon gas (99.999%). All solutions were made anaerobic using the same argon gas. The Fe(II) solution was prepared in 200 mM Mops, 100 mM NaCl pH 5.8 and the protein was in the same buffer with pH 7.0. Measurements were performed at 25 C in duplicate. The parameter settings of the ITC machine were number of injections 30 or 60, volume of each injection 3 µl, stirring rate 307 rpm, and time spacing 200 sec. The volume of the first injection was 2 µl and the resulting data point was excluded from the fit of integrated heat of binding. The results of ITC were analyzed using Origin 7.0 software under a two independent binding sites model or a single binding site model.
Results
1. Significance of non-enzymatic oxidation of Fe(II) in the presence of transferrin. Osaki (53,55) used the Fe(III) binding protein transferrin to study the ferroxidase activity, i.e. enzymatic oxidation of Fe(II) by molecular oxygen, of ceruloplasmin. In the absence of ceruloplasmin and for an Fe(II) concentration of 100 µM, the overall initial rate of non-enzymatic oxidation of Fe(II) by molecular oxygen and subsequent incorporation of the Fe(III) product into transferrin at pH 7.3 was significant, namely, 24 µM Fe(III) formed per minute (53). More recently, Duce et al used transferrin in attempts to test the ferroxidase activity of human β-amyloid precursor protein (APP), of the E2 domain of APP, and of a 22-residue segment of E2 in the form of the synthetic peptide, FD1 (63). In that study the initial rate of non-enzymatic oxidation of Fe(II) in the presence of transferrin was reported to be zero for an Fe(II) concentration of 0-200 µM and at pH 7.2. This observation contradicts the original results reported by Osaki. Therefore, we checked the significance of non-enzymatic oxidation of Fe(II) and incorporation of the Fe(III) product into transferrin at pH 7.0 in the absence of any other proteins and at various concentrations of Fe(II) (Figure 2.3). Kinetics with apparent positive cooperativity was observed and the Hill equation was used to fit the data with a Hill constant of 1.9 and a Vmax of 21 µM/min. The observed Vmax is comparable to all previous reported values by others (32,53,55), and is within experimental error comparable to the values reported by Duce et al (63) for the putative ferroxidase activity of the APP protein, the E2 domain, and the FD1 peptide. The rate of non-enzymatic oxidation of Fe(II) and formation of
diferric-transferrin dramatically increased with increasing the pH from 6.2 to 8.2. The initial rate of Fe(III) formation (µM per minute) was 0.080.01 at pH 6.2, 2.6 0.4 at pH 7.2, and 80.2 8.5 at pH 8.2 for an initial Fe(II) concentration of 40 µM. Thus, the initial rate of non-enzymatic oxidation of Fe(II) in the presence of transferrin depends on the concentration of Fe(II) and on the pH.
2. The use of transferrin to study the ferroxidase activity of H-chain ferritin and ceruloplasmin. Before using the transferrin assay for measurement of the ferroxidase activity of the FD1 peptide with the putative ferroxidase site of APP, we tested this assay by measuring the established ferroxidase activities of human H-chain ferritin (HuHF) and ceruloplasmin. Bakker and Boyer measured the iron oxidation activity of horse spleen ferritin (HoSF) at pH 6.2 in the presence of human transferrin (32). They reported the activity as formation of diferric-transferrin (i.e. sequestering of Fe(III) by transferrin from ferritin) under the – then unproven – assumption that the ferroxidase activity of ferritin was the rate-limiting step in the overall assay, and under the assumption that all the 24 subunits of HoSF were catalytically active. Subsequent studies have shown that HoSF consists of two distinct subunits, 80% L ‘light’ and 20% H ‘heavy’ subunits (32); and the catalytic center is only present in the H subunit (22,36). Measurements of kinetics of iron oxidation by HuHF have shown that oxidation of Fe(II) occurs within seconds (46). In this process two Fe(II) bind simultaneously to the catalytic site of ferritin, which is commonly known as the ferroxidase site (FC), and they are oxidized by molecular oxygen at a high rate. This reaction has typically been measured by following the UV-visible absorbance spectrum of ferritin-bound Fe(III) between 300 nm and 500 nm (103). Using this procedure a kcat = 3.3 sec-1 has been reported for HuHF (36) which is comparable to the values reported by Duce et al for ceruloplasmin and not orders of magnitude lower as claimed by them (63). We determined whether the oxidation rate of Fe(II) by the FC of HuHF in one turnover, i.e. oxidation of 2 Fe(II) per FC, is comparable to the rate of diferric-transferrin complex formation. Absorbance at 650 nm was recorded for the formation and
Figure 2.3. Effect of concentration of Fe(II)
on non-enzymatic oxidation of Fe(II) and incorporation of the Fe(III) product into transferrin. Progress curves were recorded at 460 nm for formation of diferric-transferrin and the initial rates were plotted versus concentration of Fe(II). The error bars represent standard deviation of three independent measurements. Concentration of apo-transferrin was 100 µM. Buffer was 100 mM Hepes, 100 mM NaCl, pH 7.2.
decay of a blue intermediate, the proposed µ-peroxodiferric complex, which is an indicator of the oxidation of Fe(II) in the ferroxidase site of HuHF (104,105), and at 460 nm for the formation of diferric-transferrin complex. An amount of 2 Fe(II) per FC was added to apo-HuHF in the presence of transferrin and, using a stopped-flow spectrometer, the spectra were recorded (Figure 2.4A). Within five seconds the blue intermediate with maximum absorbance at 650 nm forms and decays. This rate of formation and decay for the blue intermediate is comparable to the previously reported values for HuHF (104). The rise in absorbance at 460 nm in the first five seconds is due to the broad absorbance spectrum of ferritin-bound Fe(III) from the UV region to circa 500 nm (103). Subsequently, the absorbance at 460 nm slowly increases because transferrin scavenges ferritin-bound Fe(III). Thus the rate limiting step is the transfer of the Fe(III) from ferritin to transferrin, and – contrary to the proposal of Bakker and Boyer (32) - the rate of diferric-transferrin complex formation under these conditions is not equal to the rate of the ferroxidase activity. Therefore, the transferrin assay is not a suitable method to quantitatively measure ferroxidase kinetics of the iron-storage protein ferritin. Then, we measured the ferroxidase activity of ceruloplasmin as a function of Fe(II) concentrations using the transferrin assay (Figure 2.4B).
Figure 2.4. Use of transferrin to measure the ferroxidase activity of ferritin and ceruloplasmin. (A) Effect
of transferrin on Fe(II) oxidation by human H-chain ferritin ( HuHF) measured at 650 nm for formation and decay of the blue intermediate , and at 460 nm for formation of diferric transferrin complex. Concentration of apo-HuHF was 1.7 µM (24-mer) and that of transferrin was 70 µM. Two Fe(II) per
ferroxidase center were added and measurements were done at 34 C in 400 mM Mops buffer, 100 mM
NaCl, pH 7.0. (B) Ferroxidase activity of ceruloplasmin was measured using the transferrin assay at 460 nm. Progress curves were recorded and initial rate of Fe(III) formation was plotted versus concentration of Fe(II). Concentration of ceruloplasmin was 0.1 µM. Measurements were performed at 37 °C in triplicate. Buffer was 100 mM Mops, 100 mM NaCl, pH 7.1.
The specific activity of ceruloplasmin was calculated from the initial rate of diferric-transferrin formation which was obtained from the progress curves at 460 nm using an extinction coefficient of 4.56 mM-1cm-1 for the diferric-transferrin complex. The activity was measured
for Fe(II) concentrations between 0 and 160 µM. Addition of ceruloplasmin significantly increased the rate, and at an Fe(II) concentration of 40 µM a maximum rate of 240 µM Fe(III) formed per minute per µM of ceruloplasmin was observed (Figure 2.4B). The data were fitted to the Michaelis-Menten equation affording Km = 10 3 µM and Vmax = 3.1 0.3 U/mg (µmol of Fe(II) oxidized per minute per mg of ceruloplasmin). It can be concluded that unlike HuHF, ceruloplasmin does not store iron and the resulting Fe(III) product of the ferroxidase reaction is rapidly scavenged by transferrin and therefore the transferrin assay can be used to measure the activity of ceruloplasmin.
3. The FD1 peptide does not have ferroxidase activity in the transferrin-coupled assay. The reported ferroxidase activity of APP was tested using the 22-residue synthetic peptide FD1, which carries the putative ferroxidase active site of APP. Duce et al have reported that this peptide possesses 40% of the activity of APP or ceruloplasmin (63). The initial rate of diferric-transferrin complex formation due to oxidation of Fe(II) by FD1 was measured and was compared (i) to that of the non-enzymatic oxidation in the presence of transferrin, (ii) to that of BSA which is known not to oxidize Fe(II), (iii) to that of HuHF which should decrease the rate because it stores the Fe(III), and finally (iv) to that of ceruloplasmin (Figure 2.5A).
Figure 2.5. The FD1 peptide does not
catalytically oxidize iron as measured by the transferrin assay. (A) Non-enzymatic oxidation of Fe(II) and incorporation of Fe(III) into transferrin was compared with that of BSA, HuHF, ceruloplasmin (CP) and FD1. 52 µM of apo-transferrin was mixed with 2 µM BSA or 0.18 µM HuHF (24-mer) or 2 µM FD1 peptide, or 0.1 µM ceruloplasmin, and then an aliquot of 5 µl anaerobic solution of ferrous sulphate was added. Measurements were done in 100 mM Hepes, 100 mM NaCl, pH 7.2. Final concentration of Fe(II) was 40 µM. (B) Effect of pH on non-enzymatic oxidation of Fe(II) in the presence of transferrin was checked at three different pH values in the presence or absence of FD1 peptide (2 µM): pH 6.2, 100 mM Mes, 100 mM NaCl; pH 7.2, 100 mM Hepes, 100 mM NaCl, and pH 8.2, 100 mM Tris, 100 mM NaCl. Concentration of apo-transferrin was 52 µM. Final concentration of Fe(II) was 40 µM. Measurements were performed at 37 °C in triplicate with two different batches of transferrin.
At an Fe(II) concentration of circa 40 µM the rate of the non-enzymatic reaction is circa 3 µM Fe(III) formed per minute (Figure 2.5A and see Figure 2.3). Ceruloplasmin shows a rate that is 8-fold above that of the non-enzymatic reaction. This rate of 24 µM Fe(III) per minute (i.e. a specific activity of 3.1 U/mg) is comparable to the originally reported value for this protein (53). With the FD1 peptide, however, the rate of diferric-transferrin formation is within experimental error equal to that of non-enzymatic oxidation of Fe(II) by molecular oxygen either in the presence or in the absence of BSA. To test if the pH can have an effect on the rate of Fe(II) oxidation by FD1, the ferroxidase activity of FD1 was examined at three different pH values, 6.2, 7.2, and 8.2 (Figure 2.5B). The results were within experimental error identical to those of the non-enzymatic oxidation of Fe(II). The presence of FD1 did not change the rate of iron oxidation and diferric-transferrin complex formation at any of the tested pH values. 4. The FD1 peptide does not have oxygen-consumption activity in the presence of Fe(II). A second procedure that has been used to follow the ferroxidase activity of proteins such as ceruloplasmin and ferritin, and one that provides information about the stoichiometry of the reaction, is to amperometrically measure the consumption of the substrate molecular oxygen (102,106). Therefore oxygen consumption by FD1 was measured and was compared with that of ceruloplasmin and HuHF as positive controls. BSA does not consume oxygen and was used as a negative control. It has been shown that when two Fe(II) per subunit are added to HuHF, one molecular oxygen is consumed at a high rate for the oxidation of two Fe(II) in the ferroxidase center (36,107). When the amount of Fe(II) added per subunit of ferritin is increased, the Fe(II)/O2 ratio increases, and at Fe(II) per subunit ratios greater than 15 circa 4 Fe(II) are oxidized per molecular oxygen (108). When we added 10 Fe(II) per subunit to HuHF (Figure 2.6A) a stoichiometry of 2.4 Fe(II) oxidized per one O2 consumed was found, which is in agreement with previous studies (36,109). At Fe(II) concentration of 200 µM the rate of oxygen consumption of ceruloplasmin was comparable to that of HuHF (Figure 2.6B). As previously reported (53) a stoichiometry of one O2 consumed per 4 Fe(II) oxidized was obtained for ceruloplasmin, which suggests that molecular oxygen is reduced to water by ceruloplasmin. Comparison of oxygen consumption of FD1 with that in the non-enzymatic reaction (no peptide or protein present) and that of HuHF, ceruloplasmin and BSA (Figure 2.6) provides further independent evidence that FD1 does not have ferroxidase activity. The rate of oxygen consumption in the presence of FD1 is not different from that in the presence of BSA and is equal to non-enzymatic oxidation of Fe(II). From the combined results of iron oxidation and oxygen consumption measurements we conclude that FD1 does not have ferroxidase activity.
Figure 2.6. Oxygen consumption as a monitor of Fe(II) oxidation. (A) Consumption of oxygen as a result
of oxidation of Fe(II) was measured for HuHF, FD1, and BSA and was compared with that of the non-enzymatic oxidation of Fe(II). Concentrations of HuHF (subunit), FD1, and BSA were 14 µM. Buffer was 100 mM Hepes, 100 mM NaCl, pH 7.2. Final concentration of Fe(II) was 140 µM. Each measurement was performed in triplicate. (B) Comparison of oxygen consumption in the ferroxidase reaction of HuHF and of ceruloplasmin. Final concentration of HuHF (subunit) and ceruloplasmin (CP) was 1.1 µM. Concentration of Fe(II) was 200 µM. Measurements were performed in 100 mM Mops buffer, 100 mM NaCl, pH 7.1. For all measurements the temperature was 22C.
5. Fe(II) and Zn(II) binding to transferrin and to the FD1 peptide. Because FD1 does not show any iron oxidation activity, we checked if it binds Fe(II) at all. We measured anaerobic binding of Fe(II) to the FD1 peptide using isothermal titration calorimetry (ITC) (Figure 2.7A). No Fe(II) binding to the FD1 peptide was observed within the detection limit of ITC. Then, using ITC binding of Zn(II) to transferrin and to the FD1 peptide was measured (Figure 2.7B-C). Statistical analysis of the integrated heat of binding data was performed to obtain the thermodynamic parameters. For transferrin a model of two independent binding sites was required to obtain a fit to the data of integrated heat (Figure 2.7B). A binding with Ka1 = (6.0 1.8)·104 (M-1), (Table 2.1) and with a stoichiometry of circa one Zn(II) per transferrin was found. A second site with a stoichiometry of circa one and with a lower affinity Ka2 = (1.3 0.6)·103 (M-1) (Table 2.1) was also observed. Zn(II) has been shown to bind to human serum transferrin as well (110). Previous measurements of Zn(II) binding to transferrin using difference UV monitoring in the presence of competing ligands have suggested two strong binding sites for Zn(II) (111). Unlike the previous study we measured Zn(II) binding to transferrin directly in the absence of any competing ligands.
Figure 2.7. Fe(II) and Zn(II) binding to transferrin and to FD1 measured by isothermal titration
calorimetry. (A) Anaerobic Fe(II) titration of the FD1 peptide. Concentration of FD1 in the cell was 70 µM and that of Fe(II) in the syringe was 2.23 mM. The Fe(II) solution was prepared in 100 mM Mops, 100 mM NaCl pH 5.8, and FD1 was in 100 mM Mops, 100 mM NaCl, pH 7.0. The data for the integrated heat of binding were corrected for the heat of dilution due to titration of Fe(II) to buffer (control). Measurements were performed at 25 °C. (B) Zn(II) binding to transferrin and (C) to the FD1 peptide. For experiments with transferrin concentration of protein in the cell was 126 µM and the concentration of Zn(II) in the syringe was 12 mM. For experiments with the FD1 peptide, concentration of peptide in the cell was 70 µM and that of Zn(II) in the syringe was 3.35 mM. Transferrin, FD1 peptide, and Zn(II) were prepared in 100 mM Mops, 100 mM NaCl, pH 7.0. Measurements were performed at 25 °C. The data for the integrated heat of binding were corrected for the heat of dilution due to titration of Zn(II) to buffer (control).
Table 2.1. Thermodynamic parameters for Zn(II) binding to
transferrin and FD1 peptide.
Transferrin FD1 N1 0.8 0.06 0.5 0.2 Ka1 (M-1) (6.0 1.8)·104 (5.1 1.9)·104 ΔH1 (kJ/mol) -1.2 0.2 -2.5 0.6 ΔG1 (kJ/mol) -27.7 0.5 -26.6 1.0 N2 1 0.5 ---- Ka2 (M-1) (1.3 0.6)·103 ---- ΔH2 (kJ/mol) 45 16 ---- ΔG2 (kJ/mol) -17.4 1.2 ----
Binding of Zn(II) to transferrin and FD1 peptide was measured by isothermal titration calorimetry. N is the stoichiometry of Zn(II) binding per protein. Measurements were performed at 25 °C. A two independent binding site model or a single binding site model was used to fit the data points for integrated heat of binding. Standard deviations were obtained from triplicate experiments.
For analysis of Zn(II) binding to FD1 peptide (Figure 2.7C) a model of a single binding site was sufficient. A binding with Ka = (4.1 1.0)·104 M-1 and stoichiometry of circa 0.5 Zn(II) per FD1 peptide was obtained. This stoichiometry suggests that two FD1 molecules create one coordinating site for a single Zn(II) (Table 2.1). The observation that Zn(II) apparently does not bind to the putative ferroxidase site of a single peptide is consistent with the results of a recent X-ray crystal structure determination of monomeric E2 domain of APP in the presence of Zn(II) or Cd(II) in which no metal binding to the ligands of the putative ferroxidase site has been observed (112).
Figure 2.8. Inhibitory effect of
Zn(II) on diferric-transferrin complex formation. (A) Initial rates of diferric-transferrin formation at 460 nm are plotted versus concentration of Zn(II).
Concentration of apo-transferrin was 80 µM and that of Fe(II) was 200 µM. (B) Comparison of the inhibitory effect of Zn(II) on the diferric-transferrin complex formation for the non-enzymatic oxidation of Fe(II) by molecular oxygen in the presence of transferrin (Tf 70 µM) and for the ferroxidase activity of
ceruloplasmin (CP 0.08 µM) measured with the transferrin assay. Final concentration of Fe(II) was 60 µM and that of Zn(II) was 70 µM. Measurements were done in 100 mM Hepes buffer, 100 mM NaCl, pH 7.2 in triplicate at 37 C.
6. Zn(II) bound to transferrin inhibits binding of the Fe(III) product of Fe(II) oxidation. Because FD1 does not bind Fe(II) and does not catalytically oxidize Fe(II), we surmise that the previously observed inhibitory effect of Zn(II) on the Fe(II) oxidation rate in the presence of transferrin (63) could be due to binding of Zn(II) to transferrin and thus to a decrease in the rate of diferric-transferrin formation. Therefore, we measured the effect of Zn(II) concentration on the rate of non-enzymatic oxidation of Fe(II) (200 µM) and incorporation of the Fe(III) product into transferrin (Figure 2.8A). The overall rate decreased considerably in the presence of Zn(II) presumably because Zn(II) binds to the Fe(III) binding sites of transferrin. The addition of one Zn(II) per transferrin was enough to decrease the rate by approximately 80%. The reason for observation of this residual rate in the presence of Zn(II) is possibly the fact that Fe(III) binds transferrin orders of magnitude stronger than Zn(II). Thus Fe(III) can displace Zn(II) from its binding sites. If Zn(II) inhibition of putative ferroxidase activity of APP would be specific, i.e. because of binding of Zn(II) to the APP and not to transferrin, then Zn(II) should not inhibit ferroxidase activity of ceruloplasmin as indeed claimed by Duce et al. (63). However, we find with the transferrin assay (Figure 2.8B) that in the presence of Zn(II) the ferroxidase activity of ceruloplasmin is diminished and is comparable to the residual rate of non-enzymatic oxidation of Fe(II) in the presence of at least one Zn(II) per transferrin. This decrease in rate is apparently due to binding of Zn(II) to the transferrin and inhibition of Fe(III)-transferrin complex formation.
Discussion
In human H-chain ferritin the ferroxidase reaction occurs in the ferroxidase center made up of five residues (65) that can bind two Fe(II) simultaneously through eight coordination bonds to afford their oxidation by molecular oxygen (Figure 2.2). On the contrary, the crystal structure of the E2 domain of APP (97) shows (Figure 2.2) that the RExxE putative ferroxidase site of APP (63) consists of only two putative metal-binding residues, namely E412 and E415 in APP770. The side chain of these two residues point to the surface of the protein, i.e. there do not appear to be additional residues with which to form a metal binding site. In the previous work of Duce et al. the two residues E412 and E415, have been arbitrarily aligned with two metal-coordinating residues of the ferroxidase center of human H-chain ferritin, namely E62 and H65 (63). However, the two glutamates in the RExxE motif in ferritin are not ligands to the metals in the ferroxidase center. In other words, there is no significant overall similarity between the structure of the ferroxidase centre of ferritin and the RExxE putative ferroxidase activity carrying motif of APP.
We argue that the study reported by Duce et al. (63) on ferroxidase activity of purified APP695, its E2 domain, the 22-residue synthetic peptide FD1, and tissues containing APP, and the inhibition of these activities by Zn(II) has several important deficiencies: (i) The authors
find that non-enzymatic oxidation of up to 200 M Fe(II) in the presence of transferrin is negligible at pH 7.2 (63). This is clearly in disagreement with several previous reports (32,53,55) and with our current results. The rate of non-enzymatic oxidation of Fe(II) by molecular oxygen and incorporation of the resulting Fe(III) product into transferrin is significant at pH 7.2. This rate is not dependent on the presence of the FD1 peptide, indicating that FD1 does not have any ferroxidase activity. Increasing the pH from 6.2 to 8.2 drastically increases the rate of non-enzymatic oxidation of Fe(II) and formation of diferric-transferrin. The effect of pH is crucial since results based on small differences or changes in the pH could easily be mistaken for enzymatic activity. We find that the overall non-enzymatic rate of oxidation of Fe(II) and incorporation of the Fe(III) product into transferrin at pH 7.0 is within experimental uncertainty close to the previously claimed specific activity values for APP, its E2 domain, and FD1 (63). (ii) In the previous study the fact that Zn(II) can bind to transferrin and inhibit the formation of the diferric-transferrin complex has not been considered (63). It has been suggested that Zn(II) binds to the putative ferroxidase site of APP but not to ceruloplasmin and that it inhibits ferroxidase activity of APP only. However, in agreement with other previous studies (111,113) we have shown that Zn(II) binds to transferrin. Binding of Zn(II) to transferrin inhibits formation of the diferric-transferrin complex and therefore decreases the ferroxidase activity of ceruloplasmin in the transferrin assay. (iii) Duce et al. have not measured oxygen consumption by APP as an independent method for ferroxidase activity assay and for determination of the stoichiometry of the ferroxidase reaction of APP, however, they have proposed a model in which APP can catalyze the oxidation of four iron by one molecular oxygen (63). The oxygen consumption measurements reported here show that FD1 does not consume oxygen in the presence of iron. (iv) Duce et al. have not measured Fe(II) binding to APP, its E2 domain or FD1 peptide. Using isothermal titration calorimetry we observed that within the detection limit of this method the putative catalytic site of the FD1 peptide does not bind Fe(II).
We therefore conclude that the FD1 peptide with the RExxE ferroxidase consensus motif of APP does not carry ferroxidase activity. In view of these results and of the observation that Zn(II) binding to transferrin directly interferes with the measurement of ferroxidase activity using this protein, we suggest that the results of Duce et al (63) for the ferroxidase activity of the FD1 peptide and – by implication – of the E2 domain and the full-length APP should be re-evaluated.
Chapter Three
Catalysis of iron core formation in Pyrococcus furiosus
ferritin
This chapter is adapted from:
Honarmand Ebrahimi K., Hagedoorn P. L., Jongejan J. A., Hagen W. R., Catalysis of iron core fomration in Pyrococcus furiosus ferritin, (2009), J. Biol. Inorg. Chem. 14: 1265-1274
Abstract
he hollow sphere-shaped 24-meric ferritin can store large amounts of iron as a ferrihydrite-like mineral core. In all subunits of homomeric ferritins and in catalytically active subunits of heteromeric ferritins a diiron binding site is found that is commonly addressed as the ferroxidase center (FC). The FC is involved in the catalytic Fe(II) oxidation by the protein; however, structural differences among different ferritins may be linked to different mechanisms of iron oxidation. Non-heme ferritins are generally believed to operate by the so-called substrate FC model in which the FC cycles by filling with Fe(II), oxidizing the iron, and donating labile Fe(III)–O–Fe(III) units to the cavity. In contrast, the heme-containing bacterial ferritin from Escherichia coli has been proposed to carry a stable FC that indirectly catalyzes Fe(II) oxidation by electron transfer from a core that oxidizes Fe(II). Here, we put forth yet another mechanism for the non-heme archaeal 24 meric ferritin from Pyrococcus furiosus in which a stable iron-containing FC acts as a catalytic center for the oxidation of Fe(II), which is subsequently transferred to a core that is not involved in Fe(II)-oxidation catalysis. The proposal is based on optical spectroscopy and steady-state kinetic measurements of iron oxidation and dioxygen consumption by apoferritin and by ferritin preloaded with different amounts of iron. Oxidation of the first 48 Fe(II) added to apoferritin is spectrally and kinetically different from subsequent iron oxidation and this is interpreted to reflect FC building followed by FC-catalyzed core formation.
