Characterisation of flavodoxins and ferredoxin/flavodoxin reductases from Bacillus cereus and their interactions

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Characterisation of flavodoxins and ferredoxin/flavodoxin reductases from

Bacillus cereus and their interactions

Thesis for the degree of Philosophiae Doctor

by

Ingvild Gudim

Department of Biosciences

Faculty of Mathematics and Natural Sciences University of Oslo

2018

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© Ingvild Gudim, 2018

Series of dissertations submitted to the

Faculty of Mathematics and Natural Sciences, University of Oslo No. 2000

ISSN 1501-7710

reproduced or transmitted, in any form or by any means, without permission.

Cover: Hanne Baadsgaard Utigard.

Print production: Reprosentralen, University of Oslo.

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Acknowledgments

First and foremost I want to thank my primary supervisor, Hans-Petter Hersleth, for hiring me on his NRC Young Research Talents grant and for his patience, around the clock support, and trust in me during the last four years. I have really appreciated the opportunity to learn structural biochemistry and the freedom I’ve had to manage my own time and take on commitments that have not been related to my project. What is more, I don’t think many other people can rival HP’s kindness and generosity. I am very grateful for our group dinners, (scientific) excursions, and for the unlimited supplies of marzipan eggs in the office during my writing-up period. I am also thankful to my co-supervisor Morten Sørlie for bringing in fresh enthusiasm when joining the project and for contributing with a valuable outsider’s perspective on the work. In addition, I am obliged to Kara Bren for giving me the opportunity to work in her lab in Rochester, NY, and to Sanela Lampa-Pastirk for putting a lot of effort into supervising my project there. It was very educational working for two such committed, sharp, enthusiastic, and accomplished scientists.

Huge thanks also to Marta, Marie, Hedda, and Inger for their efforts in helping me whenever concepts or lab procedures were too biological and, along with Bie, Niels and the BMB-members on the 2^nd floor, for creating a friendly and supportive social environment. I am equally grateful to the Bren lab for warmly welcoming me and for all their help and kind invitations to various outings throughout my stay. In particular, thank you, Vincenzo, for being such a great lab- and housemate. A faccè e chi cè vò male!

In addition, I would like to thank Terry Baine, Espen Mariussen, and Philip Ash, who have been outstanding mentors and sources of inspiration to me in studying science and becoming a researcher.

Moreover, I am indebted to my wonderful friends and family, who have constantly supported me through all the ups and downs the last four years. Thank you for making life outside of lab so enjoyable.

Finally, I owe Øystein a debt of gratitude for his everlasting patience and understanding when I’ve been home late because of lab mess-ups, when I’ve been distracted or frustrated by work, and when I’ve spent time away from Oslo. Above all, thank you for bringing so much joy, happiness and love into my life.

Oslo, May 2018 Ingvild Gudim

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List of papers

This thesis is based on the following Papers I-III, which will be referred to in the text by their Roman numerals.

I. Lofstad, M.*; Gudim, I.*; Hammerstad, M.; Røhr, Å. K.; Hersleth, H.-P.

Activation of the Class Ib Ribonucleotide Reductase by a Flavodoxin Reductase in Bacillus cereus. Biochemistry 2016, 55 (36), 4998.

II. Gudim, I.; Hammerstad, M.; Lofstad, M.; Hersleth, H.-P. Characterisation of flavodoxin reductase-flavodoxin (FNR-Fld) interactions: an efficient FNR-Fld redox pair and identification of a novel FNR subclass. Submitted.

III. Gudim, I.; Lofstad, M.; Van Beek, W.; Hersleth, H.-P. High-resolution crystal structures reveal a mixture of conformers of the Gly61-Asp62 peptide bond in an oxidised flavodoxin from Bacillus cereus. Protein Science. In press. doi: 10.1002/pro.3436

* Joint authorship.

Other related publications by the author:

IV. Gudim, I.; Lofstad, M.; Hammerstad, M.; Hersleth, H.-P. Measurement of FNR-NrdI Interaction by Microscale Thermophoresis (MST). Bio-protocol 2017, 7 (8), e2223.

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Abbreviations

AdR adrenodoxin reductase

cyt cytochrome

EC enzyme commission

Fd ferredoxin

Fld flavodoxin

FNR ferredoxin/flavodoxin reductase FO flavin oxidising

FR flavin reducing

GR glutathione reductase

hq hydroquinone

KD dissociation constant MST microscale thermophoresis NHE normal hydrogen electrode

ONFR oxygenase-coupled NADH-ferredoxin reductase

ox oxidised

PDB protein data bank

QM/MM quantum mechanics/molecular mechanics RNR ribonucleotide reductase

RPKM reads per kilobase per megabase S cysteinyl radical

sq semiquinone

Trx thioredoxin

TrxR thioredoxin reductase

Y tyrosyl radical

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1 Introduction

1.1 Electron transfer proteins

Oxidation-reduction (redox) reactions, reactions which involve the transfer of electrons, are of great biological importance: most living things derive their energy through a flow of electrons from a fuel source to an electron acceptor, an oxidant.^1,2 In photosynthesis, light energy is used to power the reduction of carbon dioxide, oxidising water, to form carbohydrates and dioxygen. Conversely, eukaryotes and many prokaryotes oxidise either organic or inorganic compounds to acquire energy.² In both cases, the electrons flow from the fuel source to the oxidant along an electron transport chain, a sequence of donor and acceptor electron transfer centres. Many chemical processes are coupled to these electron transport chains. An example is the synthesis of ATP by the enzyme ATP-synthase, where over 15 electron transfer reactions occur between the initial oxidation of NADH to ATP being synthesised.^3,4

The proteins that are involved in these, and all other, redox reactions are called redox proteins. A redox protein can either be a redox enzyme, catalysing a redox reaction in a substrate, or it can be an electron shuttling protein, a current carrier that passes electrons to or from the redox enzymes.^5,6 The electrons flow between the redox proteins via catalytic sites that are connected by redox chains (Figure 1-1A).⁷ Common catalytic sites are flavins, quinones, pterins, Cu, and other miscellaneous metals; i.e. multi- electron redox centres that interact with substrates and can donate or accept pairs of electrons.⁷ In contrast, redox chain elements are single-electron redox centres that are more widely separated, such as FeS clusters and heme groups.^5,7 Most commonly, a redox protein will have one catalytic site and a redox chain leading to a binding patch

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on the protein’s surface (Figure 1-1C). For other proteins, the two catalytic sites are within the same protein, and a connecting chain might not be needed (Figure 1-1B).

Others again have only a redox chain, without any catalytic sites, connecting to catalytic sites in other proteins (Figure 1-1D).⁷

Figure 1-1: Examples of redox proteins and different catalytic sites and redox chain elements.

A. Succinate dehydrogenase (PDB ID 1NEK): two catalytic sites, heme and flavin, connected by FeS cluster chain elements. B. ascorbate oxidase (PDB ID 1AOZ): metals as catalytic sites and no chain elements. C. NiFe hydrogenase (PDB ID 3USE): a metal catalytic site and FeS cluster chain elements. D. cytochrome c554 (PDB ID 1FT5): only heme chain elements.

1.2 Electron transfer theory

The electrons have to flow through the redox protein pathway at a biologically useful rate, and the rate of electron transfer is therefore an important characteristic of an electron transfer reaction. For any electron transfer reaction, the observed rate constant depends on the activation energy, ΔG^# (Equation 1), where A is the limiting rate of the reaction.^8,9

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!_!" = A!^!∆!

#

!" [1]

According to Marcus theory, the activation energy depends on only two variables:

driving force (ΔG°) and reorganisation energy (λ) (Equation 2),^10,11

∆!^# =(∆!^°+!)^!

4! [2]

where the reorganisation energy can be defined as the energy input required to reorganise the nuclei from the equilibrium position of the reactants to the equilibrium position of the products.⁹ ΔG° is given by the redox cofactor’s reduction potential, that is, the cofactor’s willingness to acquire electrons. Combining Equation 1 and Equation 2, we see that the rate of electron transfer will depend on both the free energy and the reorganisation energy (Equation 3)^9,12:

!_!" =!_!"^! !^!(∆!^°^!!)^!^/!!"# [3]

where kET0 is the activation less rate constant (ΔG°= −λ).

In biological electron transfer reactions, both λ and ΔG° are affected by the protein structure.^3,9,13 The reduction potentials of the heme cofactor in various cytochromes provide a good example of how the cofactor’s reduction potential varies depending on the protein scaffold. Free heme in solution has an Fe(III)/Fe(II) reduction potential of

−200 mV vs. NHE (normal hydrogen electrode), whereas the reduction potential of cytochrome c is 260 mV and cytochrome f 450 mV.¹³ Similarly, the protein structure is important in lowering both the outer- and inner-sphere reorganisation energies of a biological electron transfer reaction. For example, Cu-proteins often have a heavily constrained Cu binding site, so that the Cu(II) and Cu(I) redox states have the same binding geometry, which reduces the inner-sphere reorganisation energy.^9,13

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Furthermore, the burial of redox-active cofactors within the hydrophobic protein environment not only secures redox selectivity, it also shields the redox cofactor from the aqueous environment, which can reduce the outer-sphere reorganisation energy by as much as 50%.^8,9,13,14

Burying of the protein redox centres within the nonconductive protein medium means that the redox centres cannot come into close contact.¹³ In fact, protein redox centres are often separated by large distances and can be up to 14 Å apart.^7,14,15 To get from one redox centre to another, the electrons travel through a process called electron tunnelling.³ An electron at a redox centre has a wavefunction that extends throughout the insulating protein medium in all directions, decaying exponentially. If the tail of this wavefunction encounters the tail of the wavefunction of another redox centre, the electron can tunnel from one centre to another.^7,9 The probability of tunnelling is given by the electronic coupling strength, HAB2 (Equation 4),

!_!"(!) =!_!"(!_!)!^!^!^!^!(!!!^!⁾ [4]

which predicts the exponential distance dependence for electron tunnelling.¹³ β is the distance-decay constant, and r0 the centre-to-centre distance of the donor-acceptor pair at contact.

Because the electron transfer in proteins takes place over such long distances, the wavefunctions of the electron donor and the electron acceptor are often weakly coupled.^16,17 This means that there is a low probability for product formation (i.e.

electron transfer) and that the transition state might be formed several times before electron transfer takes place. Such a process is called a non-adiabatic process, in contrast to an adiabatic process, where formation of a transition state almost inevitably

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leads to a product.^16,17 Semi-classical theory can be used to derive a rate expression for non-adiabatic electron transfer (Equation 5).^9,13,17-19 From Equation 5, it can be seen that the activation-less kET0 from Equation 3 (the pre-exponential term) is limited by the strength of the electronic coupling.⁹

!_!" = 4!^!

ℎ^!!"#!_!"^! !^!(∆!^°^!!)^!^/!!"# [5]

It is not in the scope of this thesis to go further into the theory behind electron transfer reactions in proteins. Nonetheless, the implications of these equations are useful to have in mind when considering protein electron transfer. In summary, three principal parameters determine intra-protein electron transfer rates: the free energy (ΔG°), the reorganisation energy (λ), and the cofactor distance (which influences the coupling strength).^12,16 Of these three, the distance between the redox centres is the most important factor.^15,16

1.3 Electron transfer complexes

The rate of electron transfer is, however, not necessarily the rate-limiting step in an intra-protein electron transfer reaction. For electron transfer between two proteins to occur, the proteins must first associate to form an active donor-acceptor complex, electron tunnelling between the protein redox centres must take place, and then the proteins must dissociate.¹⁸ Thus, the dynamics of the conformational sampling upon complex formation and the conformational changes in the protein-protein complex will affect the reaction rate, and these will be further discussed below.^7,17,18

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1.3.1 A brief introduction to protein-protein interactions

Protein-protein interactions can be divided into two main groups based on the stability of the protein-protein complex: obligate and non-obligate interactions.²⁰ Proteins in an obligate protein-protein complex only function when they are associated in the complex.²¹ Many homodimers are therefore obligatory complexes. Non-obligate protein complexes are divided into two groups based on the lifetime of the protein complex:

permanent and transient interactions.²⁰ Permanent (or static) protein-protein complexes have a low dissociation constant and bind strongly in a well-defined orientation.²² These complexes are stable, highly selective, and require tight binding to maintain their biological function.²² Antigen-antibody or enzyme-inhibitor complexes are examples of non-obligate, permanent protein-protein complexes.^20,22 Conversely, transient complexes are denoted by low binding affinities and low specificity, and typically form when a high turnover is required.²² These proteins often have several interaction partners at the same binding site, undermining their specificity.^22,23 Signal transduction and electron transfer are examples of reactions where transient protein complexes are formed.^20,22,24

1.3.2 The encounter complex

From the available crystal structures of protein-protein complexes, one can get the impression that the proteins only adopt a unique, stereospecific orientation in the complex.²⁵ Indeed, protein complex formation was in the past regarded as a one-step process where the proteins were either free or bound in a complex. However, the chances of forming a productive protein-protein complex from diffusional collisions alone are minute, because of the small binding patches on the proteins.^22,23 The current view, therefore, is that protein complex formation is actually a two-step process (Figure 1-2). The first step is that the free proteins associate to form an encounter complex. The

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encounter complex consists of an ensemble of weakly associated conformations that often are in equilibrium with the specific protein-protein complex.22,23,25,26 The formation of an encounter complex allows the proteins to sample multiple conformations in search for the ideal binding geometry. The encounter complex is stabilised by long-range electrostatic interactions. The resulting electric field leads to a pre-orientation of the proteins that increases the chances of the proteins colliding with proximate binding sites.^26,27 Yet, the charged binding patches mean that several conformations in the encounter complex have similar energies, counteracting the formation of a specific complex.^25,26 A specific complex requires a single orientation that has much lower energy than the other orientations. This is attained by short-range interactions, such as hydrogen bonding, van der Waals interactions, hydrophobic contacts, and specific salt bridges.²⁶ Thus, the long-range interactions that are important for the formation of the encounter complex at the same time conflict the formation of the specific complex.^22,25,26

Figure 1-2: Formation of a protein-protein complex is a two-step process, where free proteins first associate in a weakly bound encounter complex, sampling many different conformations, before forming a specific protein-protein complex. The figure is adapted from Schilder and Ubbink²⁶.

The encounter complex is significantly populated in electron transfer complexes.²⁷ These protein-protein complexes need, on the one hand, to bind in a specific manner so

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that electron tunnelling between the redox cofactors can take place, and, on the other hand, a high turnover.^22,26,27 To ensure a high turnover, both the association (kon) and the dissociation (koff) rates for the proteins must be fast. A high association rate is achieved by long-range electrostatic interactions, which is why electron transfer proteins often are highly charged.²⁷ A high dissociation rate is attained by limiting the number of favourable short-range interactions.²⁷ Both of these factors stabilise the encounter state. Fewer short-range interactions also mean that the specific protein complex will have a moderate affinity and that the proteins will bind in a less specific manner.²² This makes crystallographic studies of electron transfer complexes inherently difficult.²⁴ Still, the relative weighting of the populations in the encounter and specific states vary from complex to complex. For electron transfer complexes consisting of large proteins, it is only in the specific state that the distance between the redox centres is short enough to be compatible with fast electron transfer, and so the specific state must be populated to a certain extent. For smaller electron transfer proteins, several orientations can lead to productive electron transfer, and the specific state might not form at all.^22,27

1.4 Flavoproteins

It was the German biochemist and Nobel laureate Otto Warburg who in the early 1930s first isolated a flavoprotein, a yellow protein from yeast.²⁸ Not much later, the Swedish biochemist Hugo Theorell, who was Warburg’s student and later received a Nobel Prize of his own, discovered that upon ammonium sulphate precipitation of the yellow enzyme, the precipitate was white and the supernatant yellow.^29-31 Further studies showed that the supernatant was flavin mononucleotide, a riboflavin (vitamin B2) derivative. Now we know of almost 400 different flavoproteins, all of which contain a

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riboflavin-derived cofactor. For most flavoproteins, this flavin cofactor is either a flavin mononucleotide (FMN) or a flavin adenine dinucleotide (FAD) (Figure 1-3)³².

Figure 1-3: Chemical structure of riboflavin and its FMN- and FAD-derivatives.

Flavins are redox-active cofactors. The isoalloxazine ring system is able of both one and two electron transfer reactions, thus, the flavoprotein can exist in three oxidation states: oxidised, semiquinone (one-electron reduced), and hydroquinone (two electron, fully reduced) (Figure 1-4). The semiquinone form can be either neutral or anionic. The multiple oxidation states make the chemistry of the flavin cofactor particularly versatile.

Flavoproteins carry out electron transfer, oxygen activation, photochemistry, and substrate redox reactions.³³ Dehydrogenation, monooxygenation, halogenation, biodegradation, chromatin remodelling, DNA repair, protein folding, and biosynthesis are other examples of biological processes where flavoenzymes are involved.^32,34 Flavoenzymes are thus attractive for use in industrial biocatalysis, metabolic engineering and synthetic biology.^33,34

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Figure 1-4: Overview of the different flavin oxidation states. The oxidised state is yellow, the semiquinone state is either blue (neutral) or red (anionic), and the fully reduced, hydroquinone state is clear. Changes are marked in red (black in the structure of anionic semiquinone flavin).

The largest class of flavoenzymes is oxidoreductases (Enzyme Commission (EC) number 1), which account for 91% of all reported flavoproteins.³² Of these oxidoreductases, the largest subgroup is monooxygenases/hydroxylases (EC 1.14), then enzymes oxidising a CH-OH group (EC 1.1), and then enzymes oxidising a CH-CH group (EC 1.3).³² This thesis is, however, concerned with a less abundant type of flavin oxidoreductase, namely the ferredoxin/flavodoxin NAD(P)⁺ oxidoreductase (EC 1.18.1.2 and EC 1.18.1.3), and one of their substrate proteins, the electron transfer protein flavodoxin. Together, these two proteins form an electron transfer pathway, supplying electrons from NADPH to various redox enzymes.

1.4.1 Ferredoxin/flavodoxin NAD(P)⁺ oxidoreductases (FNRs)

Ferredoxin/flavodoxin NAD(P)⁺ oxidoreductases (FNRs) catalyse the electron transfer between NAD(P)H and the electron transfer proteins ferredoxin (Fd) or flavodoxin (Fld). In photosynthetic organisms or tissues, FNR receives electrons from Photosystem I via Fd, and reduces NADP⁺ to NADPH.^35,36 Conversely, FNRs can also oxidise NADPH and reduce Fds or Flds. These in turn transfer electrons to various redox enzymes such as cytochrome P450 (by Fd)³⁷ or nitric oxide synthase (by Fld)³⁸.

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Ferredoxin/flavodoxin NAD(P)⁺ oxidoreductases can be divided into two main families, plant-type FNRs and glutathione reductase (GR)-like FNRs, which are structurally and phylogenetically unrelated.³⁹ The plant-type FNRs can be further subdivided into plastidic and bacterial FNRs, whereas the GR-like FNRs can be divided into adrenodoxin reductase (AdR)-like FNRs, thioredoxin reductase (TrxR)-like FNRs and oxygenase-coupled NADH-ferredoxin reductase (ONFR)-like FNRs.^39-41 Both the plant-type and the GR-like FNRs have a two-domain organisation, with one FAD- and one NADP-binding domain, but the arrangement of the domains differs between the two classes.^39,40

1.4.1.1 Thioredoxin reductase-like FNRs

This thesis is concerned with thioredoxin reductase-like FNRs, which were first isolated from the green sulfur bacterium Chlorobaculum tepidum in 2002.⁴² Since then, TrxR-like FNRs have been found in several bacteria and archea, for example Firmicutes^43,44, α-proteobacteria³⁶, and thermophilic organisms^41,45. As the name implies, the TrxR-like FNRs are very similar to the TrxRs, both in structure and sequence. However, the TrxR-like FNRs lack the TrxR catalytic CXXC-motif, which reduces the substrate, thioredoxin (Trx). Therefore, the FNRs cannot function as TrxRs.

The structure of a TrxR-like FNR is shown in Figure 1-5. Similarly to the TrxRs, the TrxR-like FNRs are homodimeric. This is in contrast to the other types of FNRs, which, with a few exceptions, are monomeric.³⁶ Each TrxR-like FNR protomer has two domains, with Rossmann nucleotide binding folds for both FAD and NADPH. The NADPH-binding domain is inserted between two sections of the FAD-binding domain, connecting to the FAD-binding domain by a hinge region consisting of an antiparallel β-sheet. The two domains can rotate relative to each other due to lack of interactions

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between them.⁴⁶ Indeed, the available crystal structures of TrxR-like FNRs show that the proteins have crystallised in several different conformations.^40,46

Figure 1-5: The structure of the homodimeric YumC TrxR-like FNR from Bacillus subtilis (PDB ID 3LZW). The NADPH and FAD cofactors are shown in stick representation.

This domain rotation is not found in any other type of FNR, but the structurally similar TrxRs have a domain rotation that enables catalysis. Here, the NADPH- and the FAD- binding domains are found in only two conformational states; one where the FAD cofactor is reduced by NADPH (FR conformation), and one where the FAD cofactor is reoxidised by the active-site disulphide (FO conformation).^47-49 In the FR conformation, the NADPH- and FAD-binding domains are arranged such that the NADPH molecule is close enough to FAD for productive hydride transfer to take place. The catalytic disulphide is exposed to the solvent and can reduce the substrate, Trx. Once FAD is reduced, the NADPH-binding domain rotates away, and this rotation now positions the disulphide close to the flavin (FO conformation). The flavin reduces the disulphide, before the NADPH-binding domain rotates back to the FR conformation, commencing the next catalytic cycle.

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A rotation between the FAD- and NADPH-binding domains is likely to play a role in the TrxR-like FNR catalytic mechanism too. There is only one crystal structure of a TrxR-like FNR with NADP⁺ bound, the YumC FNR from Bacillus subtilis. Here, the NADP⁺ molecule is over 15 Å away from the FAD group, too far away for productive hydride transfer.⁴⁶ Thus, there must be a domain rotation to position NADPH close enough to reduce FAD. This notion is further strengthened through studies by Muraki and co-workers, who have used site-directed mutagenesis on the hinge region to investigate its importance for the FNR function.⁴⁰ They found that when they replaced the conserved hinge region residues Gly260 and Gly266 with proline the catalytic activity of the FNR dropped, again suggesting the importance of a domain rotation for catalysis.

Finally, the C-terminal subdomain is another distinguishing feature of the TrxR-like FNRs. The C-terminal helix of one subunit stretches over the FAD-group in the other subunit and an aromatic residue followed by two hydroxyl-containing residues in the helix stabilise the FAD cofactor by hydrogen bonding and π-π interactions.⁴⁰ This is thought to be a conserved feature of the TrxR-like FNRs, but it is not quite known how it affects their function. In one study, C-terminal deletion mutants from the B. subtilis YumC reduced Fd at a lower rate, suggesting that the C-terminal might be important in Fd binding and/or electron transfer.⁵⁰ In a recent study on C-terminal deletion mutants from C. tepidum, reduced hydride transfer rates were observed, suggesting that the C-terminal is responsible for formation and stability of charge-transfer complexes.⁵¹ The length of the C-terminal helix also varies between the different proteins. Some FNRs have a long enough helix that a dipole field may form, which could also affect reactivity.^46,50 The C-terminal residues probably also modulate the FNR redox

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potentials relative to the potential of free FAD, but there are no reports on the redox chemistry of TrxR-like FNRs. Thus, despite much work that characterises this novel class of FNRs, there are still unanswered questions regarding their catalytic mechanism and the function of their C-terminal domain.

1.4.2 Flavodoxins

Flavodoxins are small electron transfer proteins that contain an FMN cofactor.

Flavodoxins can replace the more ubiquitous electron carrier ferredoxin in many reactions, and are also found in the same subcellular location as ferredoxins; namely the bacterial cytosol or in algal chloroplasts.⁶ In fact, flavodoxins are most known for acting as a substitute for ferredoxin and are typically induced under low-iron conditions.^6,52 However, it should be emphasized that there are several flavodoxin- specific pathways in prokaryotes. Biotin synthase, cytochrome P450 BioI, pyruvate formate-lyase, bacterial nitric oxide synthase, cobalamin-dependent methionine synthase, and ribonucleotide reductase are all examples of proteins that are reduced by flavodoxins.^38,53-58 Flavodoxins are also essential in some organisms, including organisms where ferredoxins are not essential, such as Escherichia coli and Helicobacter pylori.⁶ Interestingly, some of the organisms where flavodoxins are essential are pathogenic, and flavodoxins are therefore potential drug targets.^59,60

Flavodoxins are widely distributed in bacteria, but are not found in plants or higher eukaryotes. However, flavodoxin-like domains are sometimes fused into eukaryotic genes that encode multi-domain proteins, for example mammalian nitric oxide synthase, cytochrome P450 reductase, and cytochrome P450BM-3.⁶¹ Studies on bacterial flavodoxins and their electron transfer partners may therefore increase the understanding of the electron transfer mechanisms in these multi-domain proteins.^61,62

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Figure 1-6: The structure of the short-chain Flavodoxin 1 from B. cereus (PDB ID 6FSG). The FMN cofactor is shown in stick representation. This figure is from Paper III.

Structurally, flavodoxins consist of a central five-stranded β-sheet that is surrounded by α-helices (Figure 1-6). Flavodoxins can be divided into two groups, long-chain and short-chain flavodoxins, where the long-chain flavodoxins have an extra 20-residue loop with unknown function.⁵² As expected, the flavodoxin protein structure modulates the redox potential of the FMN cofactor for both types of flavodoxins. The reduction of free FMN in solution from the oxidised to the semiquinone state has an ox/sq redox potential of −238 mV at pH 7, whereas flavodoxins typically have a more positive redox potential, ranging from −50 mV to −230 mV.⁶³ In contrast, the sq/hq potential of the protein is normally much more negative than that of free FMN; the proteins usually have a potential ranging between −320 mV to −518 mV, whereas free FMN has a redox potential of −172 mV.⁶³ Thus, the protein structure stabilises the flavin semiquinone state and destabilises the flavin hydroquinone state, making flavodoxins efficient low- potential, one-electron carriers.^62-65

The interactions between the FMN cofactor and the surrounding amino acids have been extensively studied to understand what gives rise to the shifts in the flavodoxin redox

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potentials relative to that of free FMN. Both electrostatic and aromatic interactions have been found to contribute,^52,63 but a conformational change involving a glycine peptide carbonyl that is positioned just opposite the flavin isoalloxazine N5 atom plays a particularly important role in the modulation of the flavodoxin redox potentials.^66,67 In the oxidised state, the carbonyl is pointing away from the flavin isoalloxazine ring, in what is known as an “O-down” conformation (Figure 1-7). Upon reduction of the flavodoxin to the semiquinone state, the N5 atom is protonated. The glycine residue subsequently flips to an “O-up” conformation (Figure 1-7), which allows the carbonyl to form a hydrogen bond with N5H, stabilising the semiquinone state.

Figure 1-7: The three different conformations of the glycine peptide bond, as outlined by Ludwig et al⁶⁸. The glycine peptide bond carbonyl can point towards (O-up) or away (O-down) from the flavin N5 atom. In the O-down state, the peptide bond can be in a cis or trans conformation. The figure is made using the C. beijerinckii Fld structures with PDB IDs: 5NLL (cis O-down), 2FAX (trans O-down) and 2FOX (trans O-up). This figure is from Paper III.

In connection with research on flavin-protein interactions, Martha Ludwig and co- workers made an observation that has received little attention. In the crystal structure of the oxidised, wild-type Clostridium beijerinckii flavodoxin, the Gly58-Asp59 peptide carbonyl, which is positioned just opposite the flavin N5 atom, formed a hydrogen bond with Asn137 in the neighbouring protein molecule. Ludwig et al therefore constructed

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an Asn137Ala mutant, and in this flavodoxin, the glycine peptide carbonyl was in both the O-up and the O-down conformation in the oxidised state.⁶⁸ Ludwig and colleagues hypothesised that this is representative for the solution behaviour of the C. beijerinckii flavodoxin, implying that the peptide flip does not necessarily depend on the protein oxidation state, rather, it depends on the conformational energies of the different conformers. However, this has not yet been confirmed for a wild-type flavodoxin.

1.4.3 Ferredoxin/flavodoxin reductase – flavodoxin interactions

A number of studies have characterised the interactions between flavodoxins and the redox enzymes in the various flavodoxin-specific electron transfer pathways. The interactions between flavodoxins and FNRs have, however, been much less studied.

There are some reports on the FNR-Fld interaction with plant-type FNRs, but not with the GR-like FNRs, the group that the TrxR-like FNRs belong to.^61,69 As the FNRs in these two groups differ in both sequence and structure, one cannot expect that the conclusions of the studies done on the plant-type FNR-Fld interaction will also be valid for the TrxR-like FNRs and their interaction with flavodoxins. The interaction studies that have been carried out on the TrxR-like FNRs have been centred on the FNR- ferredoxin interaction and not on the FNR-Fld interaction.36,41,70,71 Furthermore, both the plant- and GR-like FNR interaction studies have often focused on one specific FNR-Fld pair despite that many organisms have several flavodoxins and several FNRs.

It is not given that these different FNR-Fld pairs will interact in a similar fashion, or that the different Flds will reduce the possible redox partner enzymes at the same rate.

In fact, some studies have found a large difference in substrate reduction rates depending on which Fld has been used³⁸, whereas others have found similar reduction rates regardless of the Fld used in the assay⁵⁵. Thus, a better understanding of the FNR-

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Fld interaction is needed; preferably by comparing and contrasting the interaction and electron transfer rates of several endogenous FNR-Fld pairs.

1.5 The ribonucleotide reductase system

An FNR/Fld electron transfer pathway has been suggested to activate a certain class of ribonucleotide reductases (RNRs). All organisms depend on the RNR system to make new deoxyribonucleotides that are needed for DNA repair and replication.^72,73 The reduction of ribonucleotides to deoxyribonucleotides (Figure 1-8) proceeds through a radical mechanism, where a cysteinyl radical (S) in the enzyme active site abstracts the 3’-hydrogen atom of the ribose to make the deoxyribonucleotide. There are three classes of ribonucleotide reductases, class I-III, and these are separated based on their reactivity with oxygen and how the cysteinyl radical is generated.

Figure 1-8: The chemical reaction catalysed by RNRs: the reduction of ribonucleotides to deoxyribonucleotides.

The oxygen-dependent class I RNRs consist of two types of subunits, α and β. The α- subunit contains the active site where the ribonucleotide is reduced, and the β-subunit holds a metal cofactor that, upon activation, generates a tyrosyl (Y) radical. During catalysis, the radical is shuttled through to the α-subunit where the transient S radical forms and reduces the ribonucleotide. The class I RNRs are further divided into three groups, class Ia-c, based on the identity of the metal cluster in the β-subunit.⁷⁴ Recently, a fourth class, Id, has also been identified.⁷⁵

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The class Ib RNRs have a dimanganese metal cofactor in the β-subunit.^76-80 Another feature characteristic of class Ib RNRs is that the genes for the α- and β-subunits, called NrdE (α) and NrdF (β), are often clustered with two other genes, NrdH and NrdI. NrdH is a small redox-active protein with a glutaredoxin-like sequence and a thioredoxin-like structure and activity profile,^81,82 which in 1996 was found to serve as an electron donor to the NrdE subunit.⁸³ It wasn’t until 2010 that the second protein, NrdI, was found to be required to form the active Mn^III2-Y cofactor in the NrdF subunit.⁷⁶ NrdI is a flavodoxin-like protein, and the reduced FMN cofactor of NrdI reacts with dioxygen to produce an oxidant, most likely superoxide.^84,85 The oxidant is then channelled from NrdI to the NrdF dimanganese cofactor,^86,87 initiating oxidation of the metal cluster and subsequent Y radical formation.⁸⁴

NrdI must be reduced to react with dioxygen and produce the reactive oxygen species required to activate NrdF. In vitro, NrdF cofactor assembly is achieved by reducing NrdI with dithionite under anaerobic conditions whilst apo-NrdF is reconstituted with MnÎI, also under anaerobic conditions. Reduced NrdI and reconstituted NrdF are then mixed and bubbled with O2.^44,88 This procedure typically produces low yields of active MnÎII2-Y cofactor, even if many groups have put considerable effort into optimising their cofactor assembly protocols. NrdF is a functional homodimer, and so the maximum value of Y and MnÎII per β2 is 2 Y/β2 and 4 MnÎII/β2. Yet, the highest radical yield reached so far is 0.9-1.0 Y/β288,89 whilst the highest metal loading achieved is 4 MnÎII/β289. The low efficiency of the cluster assembly means that the catalytic activity of NrdF in vitro is low, and this hampers studies of the class Ib RNRs.⁸⁸ In vivo, NrdE, the catalytic Ib RNR α-subunit, is reduced by either NrdH or a thioredoxin, via a thioredoxin reductase and NADPH.^81,90 Analogously, an NrdI

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reductase would allow NrdI to function catalytically during the activation of NrdF in vivo and might also lead to stoichiometric Mn^III2-Y cofactor assembly in vitro.76,77,80,84,88,91 A flavodoxin reductase is a likely candidate as an NrdI reductase.

Indeed, a TrxR-like FNR has been shown to be involved in ribonucleotide reduction in Lactococcus lactis and is proposed to act as an NrdI reductase.⁹² However, no biochemical characterisation has been done of this interaction.

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2 Aims of study

The overarching goal of the research presented in thesis has been to investigate the structure, function, and interaction of the electron transport flavoproteins flavodoxin and ferredoxin/flavodoxin NADP⁺ oxidoreductase in B. cereus. Specifically, we have sought to establish:

- the structural basis for association and electron transfer between the FNR/Fld protein pairs in B. cereus.

- whether electron transfer is more efficient for certain FNR/Fld pairs in B. cereus, and, if so, why that is.

- whether any of the B. cereus FNRs can function as an NrdI reductase in the ribonucleotide reductase system.

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3 Summary of papers I-III

Paper I

Lofstad, M.*; Gudim, I.*; Hammerstad, M.; Røhr, Å. K.; Hersleth, H.-P. Activation of the Class Ib Ribonucleotide Reductase by a Flavodoxin Reductase in Bacillus cereus.

Biochemistry 2016, 55 (36), 4998.

* Joint authorship.

This study identifies a flavodoxin reductase (FNR) as an NrdI reductase, completing the ribonucleotide reductase class Ib activation pathway. We identified three homologous FNRs in the B. cereus genome, FNRs1-3, which belong to the TrxR-like class of FNRs.

We used microscale thermophoresis (MST) to probe the FNR-NrdI interaction and found that all FNRs bound weakly to NrdI, with KDs in the 20-50 µM range. Further, UV-vis spectroscopy was used to demonstrate that all the FNRs are able to reduce NrdI, with FNR2 reducing NrdI at the highest rate. Steady-state reduction kinetics showed that the turnover number, kcat, of the FNR2-NrdI reaction is 100 min^-1, more than 10- fold higher than the turnover number of the second best FNR, FNR1. Finally, we tested whether the FNR/NrdI system could activate NrdF in vitro. Reducing NrdI with NADPH and FNR and performing the NrdF activation entirely under aerobic conditions gave high radical yields. With FNR2 as an endogenous reductant we achieved 0.6 Y/β2, 3 times higher than the radical yield achieved with FNR1. Thus, FNR2 appears to be the best NrdI reductase. The observed radical yield, however, is lower thanthe highest reported radical yield to date (1.0 Y/β2)⁸⁹. Yet, ours is a first attempt at a novel, aerobic activation protocol involving an endogenous NrdI reductant, and it is likely that the radical yield will increase with optimisation of the activation conditions.

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Gudim, I.; Hammerstad, M.; Lofstad, M.; Hersleth, H.-P. Characterisation of flavodoxin reductase-flavodoxin (FNR-Fld) interactions: an efficient FNR-Fld redox pair and identification of a novel FNR subclass. Submitted.

In this article we report on the characterisation of a redox network consisting of three ferredoxin/flavodoxin oxidoreductases (FNR1-3), two flavodoxins (Fld1-2), and one flavodoxin-like protein (NrdI) in B. cereus. Steady-state reduction kinetics showed that the FNR-Fld reduction rates vary widely. FNR2 is the most efficient reductase for all the flavodoxins. In particular, the FNR2-Fld2 pair has the highest turnover rate, with a kcat of 9125 min^-1. Further, the redox potentials of the FNRs and Flds indicate that FNR2/Fld2 is the most favourable electron donor/acceptor pair. Moreover, we solved the crystal structures of Fld2, FNR1 and FNR2. FNR1 and FNR2 have crystallised in different conformations: the NADPH-binding domain of one FNR is rotated relative to the NADPH-binding domain in the other. We believe that the domain rotation will affect both hydride transfer from NADPH to FAD and substrate binding to FNR. We propose that the conformation of FNR1 approaches the conformation where hydride transfer takes place and that the substrate, flavodoxin, will bind to the FNR in the conformation of FNR2. We carried out molecular docking studies to support this.

Finally, in all the published structures of TrxR-like FNRs to date, an aromatic residue is stacking the FAD isoalloxazine ring on the re face. This has been proposed to be a conserved residue, and FNR2 has indeed a His in this position. Surprisingly, FNR1 has a Val, and we show through multiple sequence alignment that Val is in fact a conserved residue in several FNRs. We therefore propose that TrxR-like FNRs should be divided in two sub-classes, based on the identity of the FAD-stacking residue.

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Gudim, I.; Lofstad, M.; Van Beek, W.; Hersleth, H.-P. High-resolution crystal structures reveal a mixture of conformers of the Gly61-Asp62 peptide bond in an oxidised flavodoxin from Bacillus cereus. Protein Science. In press.

doi: 10.1002/pro.3436

In this paper we present high-resolution crystal structures of Flavodoxin 1 from B. cereus in the oxidised and one-electron reduced state. Examination of the residues in the loop that binds the flavin isoalloxazine ring showed that the Gly61-Asp62 peptide bond is in two conformational states; one where the peptide bond carbonyl is pointing towards the flavin N5 atom (termed “O-up”) and one where the peptide bond carbonyl is pointing away from the flavin N5 atom (“O-down”). This was surprising, as the peptide flip from the O-down to the O-up state has been thought to occur upon reduction of the flavodoxin. We therefore used single-crystal UV-vis and Raman spectroscopy to show that the mixture of conformers is not caused by radiation-induced reduction of the flavin cofactor. Finally, we carried out a structural survey of all the published flavodoxin structures in the Protein Data Bank. The survey shows that several flavodoxins display a pronounced conformational flexibility about the same glycine peptide bond. The degree of flexibility is modulated by the interactions with the surrounding amino acid residues that either stabilise or destabilise the possible conformers.

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4 Discussion

Even if the first account of a TrxR-like FNR was published 16 years ago,⁴² TrxR-like FNRs are still wrongly annotated as TrxRs in many published genomes. In Paper I, we show that the B. cereus strain ATCC 14579 contains three genes that encode for FNRs, BC0385 (FNR1), BC4926 (FNR2), and BC1495 (FNR3). These are annotated as either TrxRs (FNR1 and FNR3) or as both TrxR and FNR (FNR2); however, none of these genes have the TrxR catalytic CXXC motif and should therefore instead be classified as encoding for FNR proteins. A multiple sequence alignment shows that the TrxR-like FNRs cluster in three different branches on a phylogenetic tree (Paper I), however, FNR1 and FNR2 share a higher sequence identity than they do with FNR3. B. cereus also contain two flavodoxins, Fld1 and Fld2, as well as the flavodoxin-like protein NrdI (Paper I and Paper II). Thus, B. cereus has a redox network of three FNRs and three Flds/Fld-like proteins; a good starting point to compare and contrast the structure and function of different FNRs and Flds, as well as the FNR-Fld interaction and electron transfer.

4.1 The FNR-Fld complex: a textbook electron transfer complex

4.1.1 Weak binding and high turnover

Electron transfer complexes are characterised by high turnover rates, which necessitates weak binding. The weak FNR-Fld binding that we observe and describe in Paper I and II is therefore not particularly surprising, and the measured KDs in Paper I are comparable to published dissociation constants for other electron transfer complexes.⁹³ Whereas all the FNR-NrdI pairs have similar dissociation constants, the FNR-Fld turnover rates vary dramatically. Steady-state reduction kinetics studies (Paper I and II) show that FNR2 reduces all the Flds at a much higher rate than the other two FNRs; the

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FNR2-Fld2 pair and the FNR2-Fld1 pair have 91-fold and 28-fold higher rates, respectively, than the third fastest electron transfer pair, FNR2-NrdI. Interestingly, the FNR2-Fld2 pair is also able to transfer electrons under aerobic conditions, implying that for this protein pair only, the rate of FNR-Fld electron transfer is faster than the rate of reoxidation of the FNR by O2. In fact, the FNR2-Fld2 reduction rate is similar to that of the E. coli FldR-Fld pair,⁶⁹ making it tempting to presume that FNR2-Fld2 is the workhorse FNR-Fld pair in B. cereus. Moreover, the other B. cereus FNR-Fld pairs are not physiological electron transfer partners. However, one should not lose sight of the fact that electron transfer rates need only be fast enough, and that several enzyme systems work at much lower catalytic efficiencies than the diffusion-limited textbook enzymes.^14,94 Accordingly, the FNR-Fld pairs with lower electron transfer rates might be physiologically relevant electron transfer pairs by donating electrons to slower enzymes. A final point is that the reduction kinetics studies probes in vitro behaviour, so any extrapolation of the results to the actual, in vivo, reduction kinetics is speculation. Thus, we can only conclude that FNR2 is the most efficient reductase for the Flds and NrdI under the measurement conditions, and that the FNR2-Fld2 pair has a much higher turnover rate than the other FNR-Fld pairs in vitro.

4.1.2 Differing driving forces, electrostatic surface potentials, and electron transfer distances

How can the large differences in reduction rate be explained? Obvious parameters to investigate are driving force (i.e. protein redox potentials), electron transfer distance and electrostatic interactions. In Paper II, we measured the reduction potentials of the six different proteins. The reduction potentials corroborate the reduction kinetics studies, with FNR2 being the best electron donor of the three FNRs and Fld2 the best electron acceptor of Fld1, Fld2, and NrdI. The reduction potentials of the B. cereus

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flavodoxins are similar to literature values of flavodoxin reduction potentials.^68,95,96 To our knowledge, there are no literature values for the reduction potential of TrxR-like FNRs, but TrxRs from Thermoplasma acidophilum and E. coli have a FAD/FADH2

(ox/hq) potential of −305 mV and −291 mV, respectively, without the generation of the semiquinone form.⁹⁷ However, of the B. cereus FNRs, it is solely FNR3 that is reduced directly from the oxidised to the hydroquinone state. FNR1 and FNR2 are only reduced to their semiquinone state, both by various redox dyes and also by NADPH. Similarly, FNR1 and FNR2 only reduce the Flds down to the semiquinone state, whereas NrdI is reduced to the hydroquinone state by the FNRs. This is interesting because flavodoxins are thought to cycle between the hydroquinone and semiquinone state in the cell, but from our reduction potential measurements the FNRs will actually not be able to reduce the Flds to the hydroquinone state. However, we have measured the standard reduction potential, E⁰, and factors such as the pH and the concentrations of the proteins in the cell will affect the reduction potential, E, in vivo.

The FNR protein crystal structures presented in Paper II show that FNR1 and FNR2 have crystallised in different conformations. Using DynDom^98,99, we find that their NADPH-binding domains are rotated 60° relative to each other. This rotation has implications for both substrate binding and the hydride/electron transfer between the three different cofactors NADPH, FAD, and FMN. Firstly, despite repeated attempts, we have not managed to obtain a crystal structure of FNR with NADPH bound. In Paper II we have therefore docked NADPH into our structures of FNR1 and FNR2, based on the NADPH coordinates in the B. subtilis YumC structure.⁴⁶ From these structures and the YumC structure, it is clear that the hydride transfer from the NADPH cofactor to the FAD cofactor in the FNRs can only occur in certain conformations of

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the protein. Because the proton is more massive than the electron, proton tunnelling can only occur at distances between 1-2 Å.¹⁴ In FNR1, the NADPH-FAD distance is 10.2 Å (flavin N5 to NADPH C4), whereas in FNR2 the NADPH-FAD distance is 16.9 Å (flavin N5 to NADPH C4). A large-scale conformational rearrangement must therefore take place in both the FNRs for hydride transfer to happen, although the conformation of FNR1 seems to be closer to the required hydride transfer conformation.

Secondly, FNR2 has crystallised in an open conformation that appears to give more room for substrate binding, whereas FNR1 has crystallised in a rather closed conformation. In Paper II we use the ClusPro server^100-102 to dock the flavodoxins to FNR1 and FNR2. In the open FNR2 conformation, the Flds dock close to the FAD cofactor. In contrast, the Flds dock to FNR1 at several spots, all of which are distant to the FAD cofactor. Thus, the computed Fld binding site would allow for electron transfer in the FNR2 conformation, but not in the FNR1 conformation, further supporting the hypothesis that substrate binding occurs in the open conformation of FNR2.

Thirdly, in paper II we present the calculated electrostatic surfaces of FNR1, FNR2, the Flds and NrdI. A fast association rate is assured by long-range electrostatic interactions that guide the proteins towards their respective binding sites, limiting the conformational search in the encounter complex. From the calculated electrostatic surfaces, it is clear that the flavodoxins are acidic, whereas NrdI has a basic patch around the FMN cofactor. Moreover, FNR2 has a basic region around the FAD cofactor, whereas FNR1 has small basic patches distributed across its surface. This can explain the docking results. The acidic Flds are attracted to the basic binding site in FNR2, whereas in FNR1 they dock to the exposed basic patches on the surface, none of

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which are close to FAD. When FNR1 is homology modelled in the conformation of FNR2, more basic residues around the FAD cofactor are exposed to solvent, but the basic area is still smaller than in FNR2. Thus, even when the FNRs are in the same conformation, the FNR2-Fld encounter complex will be stabilised relative to the FNR1- Fld encounter complex. Similarly, the basic NrdI will be less attracted to the basic patches surrounding the FNRs’ FAD cofactors than the acidic flavodoxins.

Future studies should delve deeper into the domain rotation in the FNRs. The rotation probably happens at a slower timescale than the electron transfer events, so it is likely that the rotation is the rate-limiting step in the FNR catalytic mechanism. It would therefore be interesting to compute the energy barrier for rotation in FNR1 and FNR2.

If FNR1 and FNR2 have different rotational energy barriers, it could provide an explanation for the differences in reduction rates.

Finally, the effect of reorganisation energy, one of the two parameters in Marcus theory, is difficult to measure experimentally.⁹ This is especially true for systems where conformational changes make the observed rate slower than the rate of electron transfer.^13,17 In addition, Moser and Dutton argue that reorganisation energy is less important than driving force and distance in determining the electron transfer rate.^7,14,15 Therefore, we have not made any attempt at measuring the reorganisation energy in these systems.

4.2 What are the functions of the different FNRs in B. cereus?

All the FNRs are capable of reducing the Flds and the Fld-like protein NrdI, illustrating the lack of specificity that is often observed in electron transfer complexes.¹⁰³ Yet, given the large differences in the flavodoxin reduction rates between the different

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FNRs, it seems likely that the FNRs have different functions in B. cereus. Finding the physiological electron transfer partner(s) for each FNR is, however, methodologically challenging. Certain methods to investigate protein-protein interactions, such as pull- down assay, phage display, and tandem affinity purification, cannot be used because of the weak FNR-Fld binding. Other methods, for example cross-linking methods, are excluded due to the lack of specificity between the binding partners. Other methods again, for example FRET or gene knockout studies, might work, and studies investigating the physiological electron transfer partners of the FNRs should be attempted in the future.

Quantification of gene expression can also provide clues as to the physiological role of a protein. A study on the gene expression levels and mRNA half-lives in the ATCC 14579 B. cereus strain show that all the FNRs are expressed to approximately the same extent, lower than the median expression level value of 109 RPKM (reads per kilobase per megabase), and that the expression levels of Fld1 are three times higher than those of Fld2 (Table 4-1).¹⁰⁴ However, the half-life of Fld2 is longer than that of Fld1, and the half-life of FNR3 is twice as long as that of FNR1 (Table 4-1).¹⁰⁴ The FNR2 half-life was unfortunately not determined. A long half-life might indicate a role as a housekeeping gene or that the protein is involved in energy metabolism.¹⁰⁴ Some of these half-lives are longer than the median value of 2.6 min, but further studies must be done to make any conclusions about the roles of the different FNRs and Flds in vivo.

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Table 4-1. Half-life and expression levels of FNR1-3, Fld1-2, and NrdI from the B. cereus strain ATCC 14579. All data are from Kristoffersen et al¹⁰⁴. RPKM: reads per kilobase per megabase.

Locus tag Half-life (min) Expression level (RPKM)

BC0385 (FNR1) 3.8 49.0

BC4926 (FNR2) N/A 44.8

BC1495 (FNR3) 6.3 55.7

BC1376 (Fld1) 1.0 665.8

BC3541 (Fld2) 2.7 252.0

BC1353 (NrdI) 2.1 712.2

Our current hypotheses regarding the possible functions of the different FNRs and Flds in B. cereus will be discussed in the following sections. A preliminary conclusion, based on the presently available structural and biochemical data, is that FNR2 seems like the workhorse ferredoxin/flavodoxin reductase in B. cereus. This is strengthened by the phylogenetic analysis in Paper I, which shows that FNR2 is most similar to FNRs in organisms that only express one FNR, suggesting that these FNR2 homologues act as flavodoxin electron donors in these organisms too. It would be interesting to compare the reduction rate of ferredoxin, the other FNR substrate, with the reduction rates presented in Paper II. Maybe FNR1 or FNR3 work better as a Fd reductase?

Intriguingly, Staphylococcus aureus, another Firmicute bacterium, does not have any genes that code for flavodoxins. It has, however, genes that code for Fd and FNR (unpublished results). The S. aureus FNR is most similar to FNR1, which could indicate that FNR1 and its homologues preferentially reduce ferredoxins. Another possibility is that the FNRs function in different growth phases of B. cereus.

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4.2.1 FNR2 functions as a NrdI reductase: Completion of the RNR class Ib activation pathway

When NrdI in its reduced form was discovered to be involved in the assembly of the active NrdF cofactor, the question of how NrdI is reduced in vivo immediately arose.⁷⁶ The existence of an NrdI reductase was promptly suggested, and this idea has since appeared several times in the literature.77,80,84,88 Yet, until recently, no one has been able to identify an NrdI reductase. It seems like this is partly because the TrxR-like class of FNRs have been overlooked in the efforts to identify the NrdI reductase protein. For example, in a study on the RNR system in S. sanguinis, the E. coli flavodoxin reductase sequence was used to search for a flavodoxin reductase in the S. sanguinis genome, but this did not result in any hits.⁸⁸ In fact, S. sanguinis does contain an FNR, a TrxR-like FNR, but the protein is annotated, wrongly, as a TrxR; TrxR2. Ironically, the S. sanguinis FNR (annotated as TrxR2) was tested as a physiological reductant for the thioredoxin-like protein NrdH in the very same study that tried to identify an FNR that could act as an NrdI reductase. Obviously, the S. sanguinis TrxR2 showed no activity as a TrxR, since it does not contain the required CXXC motif.⁸⁸ Thus, the misannotations of the TrxR-like FNR genes have hampered the efforts in identifying an NrdI reductase.

It was only in 2015 that a study demonstrated that a TrxR-like FNR in L. lactis is linked to ribonucleotide reduction and that it might act as an NrdI reductase.⁹²

In Paper I, we show that FNR2 functions as an NrdI reductase in B. cereus, completing the RNR class Ib activation pathway. Interestingly, the multiple sequence alignment in Paper I shows that FNR2 homologues are found in various bacteria that encode for class Ib RNRs, such as B. anthracis, B. subtilis, L. lactis, and S. sanguinis. This suggests that these proteins may also function as NrdI reductases, but this must be verified by in vivo studies.

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Furthermore, using FNR2 as an NrdI reductase, we show in Paper I that we can perform the NrdF cofactor assembly under aerobic conditions and achieve high Y radical yields. This will be a significant help in further studies of class Ib RNR, which have been held back by the low amounts of active protein that are typically achieved with the standard, partly anaerobic activation protocol. Hopefully, optimisation of our novel protocol will lead to even higher radical yields.

4.2.2 FNR1 constitutes a novel class of TrxR-like FNRs

In addition to crystallising in different conformations, FNR1 and FNR2 also differ with regard to their C-terminal sub-domain. A conserved feature of TrxR-like FNRs, which is not seen in TrxRs, is that the C-terminal helix of one monomer stretches over the FAD cofactor in the other monomer.⁴⁰ In particular, an aromatic residue in the C- terminal helix (His, Tyr, or Phe) followed by two aliphatic hydroxyl-containing residues (Ser or Thr) stabilise the FAD cofactor by π-π interactions and hydrogen-bond interactions in all the published structures of TrxR-like FNRs.^40,46 In fact, the presence of these residues have been suggested to be a common structural feature of TrxR-like FNRs.⁴⁰ In FNR1, however, the aromatic residue is replaced by a Val. In Paper II, we present ConSurf^105-108 analyses that show that the Val is actually a conserved residue in several FNRs. Therefore, the TrxR-like FNRs might be divided in two sub-classes; one class where an aromatic residue is stacked opposite the FAD on the FAD re-face, and one where Val is positioned on the FAD re-face. The function of the conventional aromatic FAD-stacking residue is not even established. Mutants where the aromatic residue has been mutated to Phe⁴⁶, Ser^40,46, His⁴⁰, or Tyr⁴⁰, do not show large changes in reactivity or nucleotide specificity. From this, it is suggested that the residue might shield FAD from exposure to solvent during catalysis.⁴⁰ However, none of these studies have mutated the aromatic residue to a hydrophobic, aliphatic residue such as Val.

Characterisation of flavodoxins and ferredoxin/flavodoxin reductases from Bacillus cereus and their interactions