Light-Harvesting Bioconjugates as Chloroplast Mimics

Light-Harvesting Bioconjugates asChloroplast Mimics

A thesis submitted in partial fulfilment of the

requirements for admission to the degree of

Doctor of Philosophy

by

David Hvasanov

School of Chemistry

The University of New South Wales

February 2013

Table of Contents

Preface i

Acknowledgements iv

Abstract vi

1 Introduction1.1 Bioconjugation Background 1-4

1.2 Natural Photosystems for Energy Conversion 1-10

1.3 Nanoreactors and Organelles 1-13

1.4 Artificial Cells 1-17

1.5 Artificial Chloroplasts 1-21

1.6 Project Description and Goal 1-25

1.7 References 1-28

2 Synthesis of Ru(II)-Complexes and Quinone Derivatives2.1 Methods For Functionalising 2,2’:6’,2”-Terpyridines 2-2

2.2 Functionalised 4’-Aryl Terpyridines 2-6

2.2.1 Synthesis of nitro-phenyl-terpyridine 3 2-6

2.2.2 Synthesis of amino-phenyl-terpyridine 4 2-7

2.3 Functionalised Ru(II)-Bisterpyridine Complexes 2-8

2.3.1 Synthesis of [Ru(tpy)]Cl3 5 and [Ru(tpy)2](PF6)2 6 2-10

2.3.2 Synthesis of [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)]

(PF6)2 7 2-12

2.3.3 Synthesis of [Ru(tpy)(maleimide-hexylcarboxamido-

phenyl-tpy)](PF6)2 8 2-14

2.3.4 Synthesis of [Ru(4’-(4-aminophenyl)-2,2’:6’2’’-

terpyridine)2](PF6)2 9 2-16

2.3.5 Synthesis of [Ru(4’-(4-maleimide-hexylcarboxyamido-

phenyl)-2,2’:6’2’’-terpyridine)2](PF6)2 10 2-17

2.3.6 Spectroscopic properties of Ru(II)-bistepyridine complexes 2-18

2.3.7 Crystallography 2-20

2.4 Functionalised Anthraquinone-Based Acceptors 2-25

2.4.1 Synthesis of 1-amino-3-azidopropane 11 2-26

2.4.2 Synthesis of anthraquinone-2-azidopropylamide 13 2-26

2.4.3 Attempted synthesis of anthraquinone-2-propylamido-

triazole-maleimide 15 2-29

2.5 Conclusions 2-30

2.6 References 2-31

3 Bioconjugate Synthesis of Cytochrome c and Related Derivatives3.1 Purification of Cytochrome c 3-2

3.1.1 Purification of iso-1 cytochrome c using cation

exchange chromatography 3-3

3.2 Bioconjugation Methods 3-5

3.2.1 Modification of histidine 3-5

3.2.2 Modification of lysine 3-5

3.2.3 Modification of cysteine 3-6

3.3 Bioconjugation of Ru(II)-cyt c (8-cyt c) 3-7

3.4 Synthesis of Dimeric Bioconjugates 3-13

3.4.1 Synthesis of cyt c-10-BSA 3-14

3.4.2 Synthesis of cyt c-10-cyt c 3-19

3.4.3 Synthesis of cyt c-16-GFP 3-22

3.4.4 Synthesis of BSA-10-BSA 3-25

3.4.5 Effect of charge on protein dimer yield 3-27

3.5 Conclusion and Future Work 3-30

3.6 References 3-31

4 Green Fluorescent Protein as a Light-Induced Electron Donor4.1 Background 4-2

4.2 Synthesis of GFP-Acceptor Bioconjugates 4-5

4.2.1 Attempted synthesis of anthraquinone-triazole-GFP

via click chemistry 4-6

4.2.2 Synthesis of anthraquinone-GFP (12-GFP) via amine modification 4-9

4.2.3 Synthesis of viologen-GFP (16-GFP) 4-12

4.3 GFP Donor-Acceptor Studies 4-14

4.3.1 Steady-state spectroscopy studies 4-16

4.3.2 Fluorescence lifetime studies 4-20

4.4 Conclusion 4-25

4.5 References 4-26

5 Supramolecular Aggregates for Protein Encapsulation5.1 Liposomes 5-2

5.1.1 Liposome formation and characterisation 5-4

5.1.2 Enzyme encapsulation 5-11

5.2 Polymersomes 5-16

5.2.1 Aggregate formation using the ‘thermodynamic trapping’

method 5-18

5.2.2 Polymersome formation using the ‘kinetic trapping’ method 5-19

5.3 Conclusion 5-34

5.4 References 5-36

6 Photoinduced Electron Transfer Studies of Cytochrome c6.1 Background 6-2

6.2 Room Temperature Photoinduced Electron Transfer Studies 6-4

6.2.1 Biological activity using cytochrome c oxidase assay 6-12

6.3 Nitrite Reductase Mimics 6-14

6.3.1 Photoinduced nitrite reductase activity of cytochrome c 6-16


6.5 References 6-22

7 Self-Assembled Light-Driven Proton Pumping Studies7.1 Background 7-2

7.2 Photosynthetic-Respiratory Hybrid System 7-6

7.2.1 Polymersome morphologies and membrane reconstitution 7-9

7.2.2 Photoinduced pH gradient 7-12

7.2.3 Orientation of reconstituted cytochrome c oxidase 7-19

7.2.4 Dependency of proton translocation rates on pH 7-20

7.2.5 Proton pumping quantum efficiencies ( ) 7-21


7.4 References 7-24

8 Experimental8.1 Chemicals, Equipment and General Methods 8-2

8.2 X-ray Crystallography 8-9

8.3 Synthesis of Terpyridine Chromophores 8-10

8.4 Synthesis of Anthraquinone-Based Acceptors 8-17

8.5 Synthesis and Purification of Bioconjugates 8-20

8.6 Vesicle Formation and Encapsulation 8-27

8.7 Enzyme Activity and Photoreaction Experiments 8-31

8.8 References 8-36

9 Conclusions and Future Work9.1 References 9-9

AppendicesAppendix A – X-ray Crystallography

Appendix B – Bioconjugate Yield Determination

Appendix C – Standard Nitrite Curve

Appendix D – Encapsulation Efficiency

Appendix E – 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS)

Appendix F – Quantum Yield and Proton Pumping Rates

Appendix G – Abbreviations

i

Preface

The work described in this Thesis was performed in the School of Chemistry at the

University of New South Wales during the candidature. All results discussed herein are

my own work except where specific reference to the work of others is made in the text.

Some of the work in this Thesis has been published in the following journals/books:

"Induced polymersome formation from a diblock PS-b-PAA polymer via encapsulation

of positively charged proteins and peptides", David Hvasanov, Jörg Wiedenmann, Filip

Braet and Pall Thordarson, Chem. Commun., 2011, 6314-6316.

"Light-activated Bioconjugate Complexes", David Hvasanov, Daniel C. Goldstein and

Pall Thordarson, In Molecular Solar Fuels, The Royal Society of Chemistry, 2012, p

426-447.

Some of the work included in this Thesis has also been presented at the following

scientific conferences:

"Membrane-Bound Light Harvesting Bioconjugates as Chloroplast Mimics", David

Hvasanov, Joshua R. Peterson and Pall Thordarson, Light in Life Sciences Conference

(LILS2009), Melbourne, Australia, 24-27 November 2009. (Poster Presentation).

ii


Hvasanov, Joshua R. Peterson and Pall Thordarson, The Royal Australian Chemical

Institute Annual One-Day Symposium, The University of Sydney, Sydney, Australia, 3

December 2009. (Poster Presentation). Awarded poster prize.


Hvasanov, Joshua R. Peterson and Pall Thordarson, International Conference on

Nanoscience and Nanotechnology (ICONN2010), Sydney, Australia, 22-26 February

2010. (Poster Presentation).

"Light-harvesting Bioconjugates as Chloroplast Mimics", David Hvasanov, Joshua R.

Peterson, Jörg Wiedenmann and Pall Thordarson, The Royal Australian Chemical

Institute Annual One-Day Symposium, The University of Wollongong, Wollongong,

Australia, 1 December 2010. (Oral Presentation).

"Light-harvesting Bioconjugates as Chloroplast Mimics", David Hvasanov, Joshua R.

Peterson, Jörg Wiedenmann and Pall Thordarson, The 2010 International Chemical

Congress of Pacific Basin Societies (Pacifichem2010), Honolulu, USA, 15-20

December 2010. (Poster Presentation). Awarded poster prize.

"Photoinduced membrane proton pumping via polymersomes as chloroplast mimics",

David Hvasanov, Joshua R. Peterson, Filip Braet, Jörg Wiedenmann and Pall

Thordarson, The Royal Australian Chemical Institute Annual One-Day Symposium, The

University of New South Wales, Sydney, Australia, 30 November 2011. (Poster

Presentation).

iii



Thordarson, 2012 International Symposium on Macrocyclic and Supramolecular

Chemistry (ISMSC-7), Dunedin, New Zealand, 29 January-2 February 2012. (Poster

Presentation). Awarded poster prize.



Thordarson, 243rd American Chemical Society National Meeting, San Diego, USA, 25-

29 March 2012. (Poster Presentation).

iv

Acknowledgements

Firstly, I would like to thank my supervisor, Dr. Pall Thordarson, for all of his guidance,

support and enthusiasm through my PhD candidacy. I can't thank him enough for giving

me inspiring ideas during difficult periods of my candidature and always willing to look

over a presentation or article. I would also like to thank my co-supervisor, Prof. Martina

Stenzel, who provided guidance in polymer aggregate formation and access to her lab.

A special mention to Dr. Joshua R. Peterson and Dr. Shiva Prasad for their

extensive help and advice during the first year of this project. Thanks to all the members

of the Thordarson group for their helpful contributions, current members - Ethan Howe,

Scott Jamieson, Warren Truong, Ekaterina Nam, Andrew Robinson, James Webb, Alex

Mason, Lev Lewis and Alistair Laos - and past members - Dr. Sabrina Dehn, Dr. Daniel

C. Goldstein, Dr. Joris Meijer, Dr. Katie Tong, Luuk Olijve, Scott Jones, Lip-son Chin,

Katrine Qvortrup, Grace Lim, Haythim Hassanein, Dithepon Pornsaksit, Ben Lewis,

Tommy Pieszko, Dedrick Song and Carol Hua. Thanks also to Dr. Alex Falber for

adding colour to the lab.

Thanks to Ethan Howe, Scott Jamieson, Warren Truong and Dr. Danmar Gloria

(surrogate group member) for letting me release stress when going out to coffee or rock

climbing. William Zhang, it's always interesting getting into discussions about quirky

topics.

Thanks to Dr. Ron Haines for keeping the office in order and Ian Aldred from

the Store. Thanks to the Centre for Macromolecular Design for letting me use their

dynamic light scattering instrument. Thanks to all the staff at the Mark Wainwright

Analytical Centre including, Lewis Alder, Dr. Leanne Stephenson and Dr. Anne Poljak

from the bioanalytical mass spectrometry facility; Dr. Mohan Bhadbhade (X-ray

crystallographer) from the solid state and elemental analysis unit; Katie Levick from the

v

electron microscope unit; Dr. Renee Whan and Dr. Henry Haeberle from the biomedical

imaging facility and all the staff at the NMR facility. I would like to thank Dr. Dylan

Owen for initial fluorescence microscopy studies. I thank A/Prof. Filip Braet and

Delfine Cheng at the Australian Centre for Microscopy and Microanalysis, The

University of Sydney for cryo-TEM analysis. I thank A/Prof. Timothy W. Schmidt, Dr.

Raphaël G. C. R. Clady and Mr. Murad Tayebjee at The University of Sydney for

time-resolved fluorescence lifetime studies and Dr. Jörg Wiedenmann for providing the

GFP and plasmids.

I am grateful to both the University of New South Wales and the Australian

Research Council for financial support of this work. I would also like to thank the

Graduate Research School for a Research Travel Award, which allowed me to travel to

Honolulu to present my work at an international conference.

Finally, I must thank my family and friends for all of their love and support

during my candidature.

vi

Abstract

Cells are highly complex bio-nanoreactors comprised of a complex medium where

multiple multistep reactions occur simultaneously across the cell. To prevent

interference and degradation of these catalytic pathways, cells compartmentalise.

Compartmentalisation achieves control of synthetic pathways at specific sites in the cell

(organelles). In nature, compartmentalisation is demonstrated in plants by the organelle,

chloroplast, which is responsible for photosynthesis. Photosynthesis is one of the most

important biological reactions, responsible for sustaining life.

This Thesis describes the preparation of light-activated donor-acceptor

bioconjugates for use as artificial chloroplasts. The studies were aimed at the

development of synthetic compartments for the reconstitution of light-harvesting

bioconjugates to construct a semi-synthetic electron transport chain capable of

generating a transmembrane proton gradient using light. To achieve that end,

bioconjugates were formed using the redox metalloprotein yeast iso-1 cytochrome c

from Saccharomyces cerevisiae and photoactive maleimide functionalised

ruthenium(II)-bisterpyridine chromophores for attachment to a free single cysteine in

iso-1 cytochrome c (CYS102). The resulting photoactive donor-acceptor bioconjugate is

capable of undergoing electron transfer upon photoactivation using 480 nm light.

The synthesis of a number of terpyridines, ruthenium(II)-bisterpyridine

complexes and maleimide functionalised ruthenium(II)-bisterpyridine complexes for use

as photoactive electron donors were prepared to investigate light-activated electron

transfer. The synthesis of asymmetric maleimide complex was achieved by initially

preparing 4’-(4-aminophenyl)-2,2’:6’,2’’-terpyridine ligand (4) synthetically based on

the modified Kröhnke method. Following from the amine 4'-aryl functionalisation, the

synthesis of asymmetric complex [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)](PF6)2

vii

(7) was achieved using methods adapted from the synthesis of iridium(III)-bisterpyridine

related complexes, followed by maleimide attachment using peptide coupling

chemistry, producing [Ru(tpy)(maleimide-hexylcarboxamido-phenyl-tpy)](PF6)2

complex (8) to allow modification with iso-1 cytochrome c. The bismaleimide complex

equivalent 10, was prepared using a similar approach to allow the preparation of dimeric

bioconjugates. Asymmetric donor-acceptor bioconjugate was prepared from complex 8.

Similarly, bismaleimide complex 10 and bismaleimide viologen 16 were used to

prepare high molecular weight bioconjugates using combinations of single cysteine

residue proteins including iso-1 cytochrome c (cyt c), bovine serum albumin (BSA) and

green fluorescent protein (GFP) to explore factors affecting yield. The resulting

bioconjugates 8-cyt c, cyt c-10-cyt c, cyt c-10-BSA, cyt c-16-GFP and BSA-10-BSA

were purified using chromatographic techniques including immobilised metal affinity

chromatography (IMAC), size exclusion chromatography (SEC) or strong cation

exchange chromatography (CEX). Interestingly, electrostatic interactions played a

significant role in dimer bioconjugate yields. The 80 kDa heterodimer of

complementary charge, cyt c-10-BSA, resulted in a 30% yield. However, like-charged

homo/heterodimer bioconjugates resulted in low yields of less than 1%.

Characterisation of bioconjugates by UV-Vis, MALDI-TOF mass spectroscopy and gel

electrophoresis were exploited to demonstrate attachment and assess purity.

The alternative biological donor-synthetic acceptor system was also explored

using the light-induced electron donor GFP. Synthesis of a GFP bioconjugate as a

potential covalent donor-acceptor system based on the

N-hydroxysuccinimide-anthraquinone 12 acceptor was prepared using non-specific

lysine modification. Additionally, the GFP was site-specifically functionalised with

viologen 16 as an alternative potential donor-acceptor bioconjugate. Light-induced

viii

electron transfer studies between GFP and p-benzoquinone, anthraquinone and viologen

electron acceptors as both non-covalent and bioconjugate mixtures were monitored

using steady-state techniques including UV-Vis and fluorescence spectroscopy and

performed using a Xenon arc lamp or custom built LED array. Time-resolved

fluorescence lifetime measurements were exploited to measure and confirm

electron/energy transfer processes of GFP-acceptor systems. Typically, a ket of the order

109 s-1 for the rate of electron transfer between GFP and the quinone acceptors was

observed.

In order to compartmentalise the ruthenium(II)-cytochrome c bioconjugates to

construct artificial organelles, polymersomes (polymer vesicles) were formed based on

a novel method developed in this project based on the polyelectrolyte,

polystyrene140-b-poly(acrylic acid)48 (PS140-b-PAA48). Polymersomes could be induced

in the presence of positively charged biomolecules including cyt c and GFP,

concomitantly encapsulating the biomolecules in the membrane. In the presence of

simple salts or negatively charged biomolecules such as calmodulin, micelles form.

Liposomes (lipid vesicles) based on egg L- -phosphatidylcholine or synthetic

L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine were explored, however, their use as

compartments were limited due to poor encapsulation efficiencies.

Photoinduced electron transfer studies between Ru(II)-bisterpyridine complexes

and iso-1 cytochrome c (cyt c) as non-covalent mixtures and asymmetric bioconjugate

8-cyt c were studied. Electron transfer studies were carried out as bulk solution

bioconjugates and membrane encapsulated 8-cyt c. The electron transfer processes

resulting in reduction of the heme of cytochrome c is followed by steady-state UV-Vis

spectroscopy. Electron transfer to the heme was achieved by irradiation using a 465 nm

LED light under anaerobic conditions at room temperature in the presence of sacrificial

ix

electron donor, ethylenediaminetetraacetic acid (EDTA). In addition, light-induced

electron transfer studies between a non-covalent mixture of Ir(III)-bisterpyridine

complex and horse heart cytochrome c was studied to mimic nitrite reductase behaviour.

Excitation and catalysis was performed using a 372 nm LED source under anaerobic

conditions at room temperature in the presence of 100 nm liposomes.

A primitive synthetic chloroplast capable of converting light energy into

chemical energy by generating a proton gradient upon photo-excitation was developed.

An artificial photosynthetic-respiratory hybrid was achieved by replacing chlorophyll

(photosensitiser) with 8-cyt c. This light-harvesting bioconjugate as well as its natural

electron acceptor, cytochrome c oxidase, is encapsulated in the synthetic polymersome

membrane of the diblock copolymer PS140-b-PAA48, reminiscent of a photosynthetic

membrane construct. The successful self-assembly of this complex light-harvesting

enzyme cascade within membranes of polymeric vesicles were characterised using

confocal laser-scanning microscopy (CLSM), TEM and cryo-TEM. Upon

light-activation, cytochrome c oxidase (reduced) is capable of pumping protons across

the membrane to generate an electrochemical gradient which is detected using a

fluorescent pH dye encapsulated within the aqueous core of the polymersome. Upon

irradiation at an initial pH of 7.2, a proton pumping rate of 3.3 × 103 H+/s across the

polymer bilayer generating a gradient up to pH 0.2 was observed.

Chapter 1

Introduction

Chapter 1 Introduction

1-2

Reproduced from New Scientist, 14 April 2012, Turn a new leaf cover story, p 28.


1-3

1 Introduction

This Thesis describes the development of a primitive chloroplast – a system that

converts light energy into chemical energy in the form of an electrochemical gradient

( ). Such a system can in principle be used as a “nanoreactor battery” where the light

induced proton gradient is used to drive chemical reactions or biochemical processes.

In plants, photosynthesis is carried out in organelles known as chloroplasts,

which converts light energy into chemical energy in the form of adenosine triphosphate

(ATP).1 Photosynthesis is highly important as it provides the majority of biomass on

Earth and produces oxygen to sustain life. This process is achieved by photoinduced

generation of an electron-hole pair followed by rapid charge transfer across the electron

transport chain coupled with proton translocation across a membrane, which drives the

production of ATP.2

Biological compartments (organelles) in nature allow chemical reactions to

occur in a confined space which couple reactions in time and space resulting in enzyme

cascade systems.3 This ensures that the product of a chemical conversion is the substrate

or catalyst of the subsequent process. Compartmentalisation achieves well-defined

reaction environments which vary from simple nanometer-sized systems to complex

micrometer-sized assemblies, while isolating catalytic cycles preventing degradation by

proteases and controls the flux of molecules in or out of compartments.4 Synthetic

compartments based on polymer or phospholipid vesicles have been heavily studied in

literature to create nanoreactors.3,5

Inspired by nature, this Thesis aims to develop an artificial photosynthetic

system capable of converting light energy into a stored proton gradient. The system

combines facets of photosynthetic concepts including the construction of a photoactive

hybrid electron transport chain capable of proton translocation which is reconstituted


1-4

using synthetic compartments based on amphiphilic polymer vesicles (polymersomes).6

The key component of the artificial hybrid photosynthetic-respiratory system is a

donor-acceptor bioconjugate linking an electron donating photosensitiser to a redox

protein and electrostatically coupled to mitochondrial cytochrome c oxidase for

vectorial proton translocation.

The development of artificial photosynthetic systems to generate proton

gradients upon photoexcitation have been previously reported using synthesised organic

donor-acceptor triads7 or aryl diimides8 in phospholipid vesicles or reconstitution of a

light-driven transmembrane proton pump (bacteriorhodopsin) in a polymersome

membrane.9 However, issues of these systems still exist including stability,

reproducibility and robustness. This Thesis seeks to address these issues by developing

methods capable of preparing light-activated bioconjugates as a component of a

semi-artificial photosynthetic-respiratory electron transport chain and methods for

concomitant reconstitution of both hydrophilic and hydrophobic enzymes in membrane

domains of polymersomes.

1.1 Bioconjugation Background

Proteins play an important role in nature; they are responsible for the enzymatic

conversions of nearly all processes in living organisms, including DNA replication and

photosynthesis. In addition, they have structural roles, acting as scaffolds in cells to

support and protect tissues and organisms. Furthermore, proteins are involved in cell

signalling within and between cells which regulate the functions of cells via signal

transduction. Commonly, protein functions depend on post-translational modifications


1-5

after expression such as glycosylation which allows for proper folding and increased

stability.†

Scientists have long sought to understand and exploit the properties of proteins

by incorporating synthetic molecules to introduce and enhance novel (unnatural)

functionalities. This has led to the development of biotechnological, biomedical and

pharmaceutical industry related devices such as biosensors10, bioelectronics11, biofuel

cells12 and bioconjugated therapeutic proteins/drugs.13 In this Thesis, this exploitation is

essential towards the development of a primitive chloroplast, as this will allow

researchers to mix and match the best from synthetic and biological light-harvesters

(chromophores) and electron donor/acceptor systems.

Bioconjugation can be defined as the linking of two or more components to form

a novel complex with the combined properties and structural characteristics of the

individual component molecules.14 In general, bioconjugation combines a biological

component (protein, biomolecule) with a synthetic component creating a chimeric

system resulting in a novel class of biomaterials to perform novel functions.

Theoretically, the properties of the bioconjugate is the linear combination of the

properties of the multi-component system.15 This allows bioconjugates to overcome

their intrinsic limitations and to possess properties and functions which otherwise would

not exist in nature.

The advent of bioconjugation methods over the last few decades has led to the

development of different chemoselective conjugation methods.16 Bioconjugation

chemistry has contributed to significant research progress and understanding in the field

of life sciences. Through careful modification of proteins, scientists have been able to

understand protein structure, functions and interactions.17

† Parts of this work have been published: Hvasanov, D.; Goldstein, D. C.; Thordarson, P. In Molecular Solar Fuels; The Royal Society of Chemistry, 2012.


1-6

One of the most extensively studied areas of protein conjugates is PEGylation,

as it allows for enhancement of peptides and proteins for pharmacological and

therapeutic use in vivo and in vitro since it was first demonstrated in 1977.18 Attachment

of the hydrophilic poly(ethylene glycol) (PEG) polymer via PEGylation has led to

improved targeting and tissue penetration for protein therapeutics in the biomedical

industry.19

Biomolecules other than proteins such as low molecular weight co-factors and

ligands are also common targets for bioconjugation and functionalisation. These often

include porphyrins (heme)20, NAD(P)+21, FAD22, lipids23, sugars and oligosaccharides24,

nucleic acids and nucleotides25 and biotin.26 However, these targets are beyond the

scope of this project.

Bioconjugation reactions involving chemical modifications and chemical

cross-linking are often divided into classes of reactions. These can be separated into

three generic types of functionalisation approaches; including direct functionalisation,

indirect functionalisation and functionalisation with cofactors/ligands as shown in

Figure 1.1.15

Direct functionalisation involves chemically modifying a synthetic ligand

possessing a specific functional group with a protein of interest. Indirect

functionalisation requires bioconjugation between the synthetic or biomolecule

components via a spacer containing heterobifunctionality. Finally, indirect

functionalisation is achieved by reaction of a synthetic ligand with a cofactor, which is

then reconstituted into an apoprotein through self-association (ligand binding).


1-7

Figure 1.1. Schematic for the classes of protein functionalisation. (a) direct functionalisation. (b) indirect functionalisation. (c) indirect functionalisation with a cofactor/ligand.27

Proteins and enzymes are most commonly modified by chemical modification

between a synthetic ligand and a targeted amino acid residue. These residues often

include lysine (Lys), cysteine (Cys), histidine (His), tyrosine (Tyr) and glutamine (Gln).

The most common method for chemical modification in bioconjugate chemistry is the

covalent linkage between a ligand and the amines of a protein. Coupling to the -amino

group on a lysine residue is often targeted as they are abundant on the surface of

proteins as well as the -N terminus of a protein. They can be easily modified by

reacting with an N-hydroxysuccimide (NHS) activated ester.15,28 It should be noted,

chemical bond formation between NHS-esters and amines offer poor site-specificity as

the natural occurrence of lysine residues in mammalian proteins is estimated to be

approximately 6%.29


1-8

Another popular target for chemical modification of natural amino acids is

cysteine (cys). Bioconjugation with proteins are generally dependent on nucleophilic

addition or displacement reaction mechanisms of the amino acid residue with an

activated synthetic ligand.17 Based on the theory of nucleophilicity proposed by

Edwards and Pearson30, the thiol group of cysteine is one of the strongest nucleophiles

in proteins. Cysteine residues possess thiol groups with a pKa of approximately 9.14 Due

to these properties, specific chemical modifications of cysteines can proceed selectively

and rapidly under benign conditions.15 Common modification reagents used in the

literature include haloalkyl compounds, thiosulfonates and maleimides. However,

maleimides are ideal as they allow protein modification at neutral pH (6.8-7) coupled

with control of specificity. They can be used to chemically modify cysteines as

maleimide-functionalised ligands are Michael acceptors.31

Cysteine residues are rare in proteins and often only contain a single accessible

residue available for modification. Alternatively, proteins which lack surface cysteine

residues can be introduced via mutagenesis for site-specific modification.32 This is a

major advantage for synthesis of site-specific bioconjugates when targeting natural

amino acids. Furthermore, maleimides are suitable for site-specific bioconjugation as

they specifically react with cysteines at neutral pH and only side-react with other amino

acids above pH 8.15

As a result of the abundance of natural amino acids in proteins, synthetic

biologists and chemists have developed techniques to introduce non-natural amino

acids, allowing for chemoselective bioconjugation reactions. These non-natural amino

acids include azides and alkynes which can be introduced selectively into proteins by

semisynthetic or recombinant methods for [3+2] Huisgen cycloaddition reactions.33


1-9

The methods described above have been used to synthesise light-activated donor

acceptor systems as described in Chapter 3, 6 and 7 for photoinduced electron transfer

measurements. Similarly, electron transfer studies of metal complex modified redox

proteins have been explored over several decades by Millett and Gray.34

One of the approaches to develop a photoactive bioconjugate by Pan and

co-workers is the modification of lysine residues of horse heart cytochrome c with a

Ru(II) bipyridine photoactive chromophore.34e The luminescence decay rates of the

resulting bioconjugates were found to be between (3.5-21) × 106 s-1, indicating possible

electron transfer from the excited triplet state of ruthenium to the ferric heme group of

cytochrome c.

Peterson et al.35 in the Thordarson group have conjugated Ru(II) bisterpyridine

chromophores with yeast iso-1 cytochrome c creating a light-activated donor-acceptor

bioconjugate via a long and short chain spacer as shown in Figure 1.2. Bioconjugates

were modified using maleimide functionalised ligands with the CYS102 residue

through a Michael addition reaction. Electron transfer rates of resulting short and long

chain bioconjugates were 5.95 × 105 and 2.78 × 105 s-1, respectively.

Figure 1.2. Photoinduced electron transfer of Ru(II)-bisterpyridine cytochrome cbioconjugate.27


1-10

1.2 Natural Photosystems for Energy Conversion

Significant breakthroughs have been made in literature to mimic photosynthesis by

extracting natural photosystems from photosynthetic organisms which are covalently

bioconjugated to nanoparticles or semiconductors to produce hydrogen or electricity,

respectively, upon light-activation.

In an elegant approach, Grimme et al.36 have developed a light-harvesting

system resulting in chemical transformation by producing hydrogen as shown in Figure

1.3. They have utilised a biological electron donor, photosystem I (PSI) found in

photosynthetic organisms and conjugated it to a platinum or gold terminal electron

acceptor nanoparticle. Conjugation was initiated by formation of [4Fe-4S] clusters in

solution by reacting sodium sulphide, ferrous ammonium sulphate and mercaptoethanol.

The resulting cluster was then reconstituted into the PSI stromal protein PsaC to yield

holo-C13G/C33S variant PsaC. PSI was then modified by combining reconstituted PsaC

and P700/Fx cores in the presence of PsaD. Finally, 1,6-hexanedithiol was introduced to

displace the mercaptoethanol ligand from the [4Fe-4S] cluster and to covalently link

PSI and Pt/Au nanoparticle. This resulting PSI photoactive bioconjugate evolves

hydrogen upon white light illumination using a Xenon arc lamp. The resulting chemical

transformation is due to the formation of a charge-separated state when PSI absorbs

wavelengths below 700 nm which is then transferred to the nanoparticle with a redox

potential favourable for hydrogen production. In a recent paper, optimisation of this

system has been reported, maximising H2 production in a platinum-based PSI

bioconjugate.37


1-11

Figure 1.3. Photosystem I/nanoparticle bioconjugate evolving hydrogen upon light-activation.36

Iwuchukwu et al.38, have utilised similar concepts by exploiting natural photosystems in

conjunction with nanoparticles to induce hydrogen production after irradiation. The

photosystem I (PSI)/Platinum (Pt) nanoparticle bioconjugate as shown in Figure 1.4 was

prepared by self-assembled platinisation on the stromal surface of PSI. Upon irradiation,

the system allows electron transport from cytochrome c6 to PSI and finally to the Pt

catalyst which produces hydrogen gas with a 25-fold greater yield than current

biomass-to-fuel strategies.


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Figure 1.4. Photosystem I/Platinum nanoparticle bioconjugate produce hydrogen gas due to electron transport from cytochrome c6 to photosystem I and finally platinum nanoparticles upon irradiation.38

Other than hydrogen production, photosystem hybrids have been used to generate

electricity for the application of photoelectrochemical cells. In a sophisticated system,

Ham et al.39, have developed a regenerative photoelectrochemical complex capable of

reversible solar energy conversion. As shown in Figure 1.5, the photoelectrochemical

complex involves single-walled carbon tubes which act as a support for self-assembly

of lipid bilayers with membrane scaffold proteins to form nanodiscs, which allow

reconstitution of the photosynthetic reaction centres for electricity generation through

the carbon nanotubes upon irradiation. This system allows self-healing by induced

disassembly upon addition of surfactants and reassembly upon its removal over an

indefinite number of cycles. Similarly, Govorov has studied photosystems for electricity

production using a simpler system by incorporating PSI to a semiconductor nanoparticle

(CdTe) via bioconjugation.40


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Figure 1.5. Reversible self-assembly of photosynthetic reaction centres on single-walled carbon nanotubes via nanodiscs for photoinduced electrical current generation.39

1.3 Nanoreactors and Organelles

Despite efforts to mimic photosynthetic reactions by exploiting photosystems from

natural sources, these systems are not representative of chloroplast organelles as they

are free floating systems in solution. In contrast, living organisms perform reactions and

chemical conversions in a confined environment in order to create a highly organised

reaction space which organises metabolic processes.6 This shows that

compartmentalisation is essential to life and also as proposed by Szotzak et al.41,

compartmentalisation played a crucial role in the origin of life by allowing molecules to

be kept in close proximity and allowing advantageous mutations during replication,

leading to evolution. Cells and organelles perform crucial roles to maintain life

including (1) chemical conversions using enzymes, (2) self-reproducing vesicles, (3)

self-reproducing genetic information and (4) energy storage. In this section, the

developments made towards artificial compartments for chemical reactions as

nanoreactors are discussed.

Since the introduction of liposomes (lipid vesicles) by Bangham et al.42 in 1965

for the preparation of artificial compartments, enzyme encapsulation within the interior

of liposomes have been predominantly employed for protein therapeutic applications43

and drug delivery.44 However, attempts toward nanoreactors using phospholipid


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vesicles have been limited for compartmentalised chemical reactions due to hindered

bilayer permeability preventing diffusion of substrate/product molecules in and out of

the vesicles.45 Walde et al.46 prepared 1-palmitoyl-2-oleoyl-sn-glycero-3-

phosphocholine (POPC) liposomes encapsulating water soluble proteinase

-chymotrypsin in the aqueous interior. This system demonstrated selectivity for small

external substrates including benzoyl-L-Tyr-p-nitroanilide and

acetyl-L-Phe-p-nitroanilide for which the membrane was permeable, allowing enzyme

hydrolysis of the substrate to produce p-nitroaniline. Subsequently, Blocher et al.47

expanded on the system to model the enzyme reactions for encapsulated

-chymotrypsin.

In recent years, there has been growing interest in polymersomes for

compartmentalisation due to their improved stabilities, allowing vesicles to remain

stable for weeks to months rather than hours to days for liposomes.48 Similar to

liposomal systems, polymersomes have been used for drug delivery49 and protein

therapeutic applications for biomedicine.50 However, polymersomes based on block

copolymers are ideal candidates for nanoreactors due to the tunability of monomer

blocks allowing control of permeability51, glass transition temperature (Tg)52 and

membrane thickness.53

In an elegant system, Vriezema et al.54 described an enzyme cascade system

based on polystyrene40-b-poly(L-isocyanoalanine(2-thiophen-3-yl-ethyle)amide)50

(PS-PIAT) which possessed intrinsic membrane permeability due to PIAT forming rigid

rod-like helices during self-assembly, allowing diffusion of small substrates/products in

and out of the water pool. A three enzyme cascade system was prepared by

encapsulation of glucose oxidase (GOX) in the lumen, horse radish peroxidase (HRP) in

the membrane and Candida antarctica Lipase B (CALB) in the bulk solution. Addition


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of substrate 1,2,3,4-tetra-O-acetyl- -glucopyranose (GAc4) resulted in hydrolysis of the

acetate groups by CALB in the bulk solution to produce glucose. Subsequently, glucose

diffused into the water pool following the tandem reaction as shown in Figure 1.6 and

the cascade reaction produced a coloured radical cation product (ABTS +) which was

detected spectroscopically. Extensions of this work have been attempted including

encapsulation of chloroperoxidase55 or covalently linking CALB to polymersomes via

an azide-alkyne click reaction.56

Figure 1.6. Schematic representation of a three-step enzyme cascade system allowing the multistep reaction in PS-PIAT polymersomes.54

Meier et al.57 have investigated nanoreactors by reconstitution of channel proteins with

significant focus on porin OmpF from outer cell walls of Gram-negative bacteria.

Rather than diblock copolymers, the Meier group use triblock copolymers as they

predominantly favour vesicle formation and the polymer used possess low Tg, allowing

extrusion to size polymersomes analogous to liposomal preparation.58 Porin OmpF was

reconstituted in poly(2-methyloxazoline)-b-poly(dimethylsiloxane)-b-poly(2-

methyloxazoline) (PMOXA-PDMS-PMOXA) polymersome membranes which act as a


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size-selective filter allowing passage of molecules below 400 Da while -lactamase was

encapsulated in the interior volume.59 The nanoreactor containing -lactamase

hydrolysed ampicillin (diffused into the interior from the bulk through OmpF) into

ampicillinoic acid which reduces iodine to iodide as shown in Figure 1.7. The reaction

was monitored by micro-iodometry via the decolourisation of a starch-iodine complex.

Variations of this system have been made including the use of channel protein FhuA60,

pH-switchability61, surface immobilisation62 and reduction triggered release4

nanoreactors.

Figure 1.7. Representation of a nanoreactor containing -lactamase and membrane reconstituted porin OmpF from an amphiphilic triblock copolymer.59

Stimuli responsive polymersome nanoreactors have also been explored to control

membrane permeability. Polymersomes were formed in a two component mixture

poly(ethylene glycol)-b-polystyrene:poly(ethylene glycol)-b-poly(styrene boronic acid)

(PEG-b-PS:PEG-b-PSBA) with the minor component PEG-b-PSBA being sensitive to


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pH and monosaccharides resulting in disassembly. CALB was encapsulated in the

lumen and enzymatic reactions resulting in hydrolysis of DiFMU octanoate or

p-nitrophenyl acetate substrates was initiated by pore formation after addition of

D-glucose or increasing pH.63

Other than liposomes and polymersomes, there has been growing interest in

nanocapsules64 and capsosomes65 as alternative nanoreactors. Of particular interest are

capsosomes pioneered by the Caruso group due to their capsule:liposome hybrid nature

allowing a microreactor with the possibility of thousands of sub-compartments.66 It

should be noted that specific control of individual sub-compartments still remains a

challenge.

1.4 Artificial Cells

Significant research has been undertaken to investigate the use of compartments for

nanoreactors and enzyme cascades. Another facet of mimicking cells and organelles is

the development of artificial cells using liposomes and polymersomes for genetic

information replication and membrane reproduction. These studies have been conducted

to explore the origin of life using low molecular weight amphiphiles.5b,67

In a landmark article by Szostak et al.41, they proposed three requirements for a

model protocell as (1) bearing an informational substance (DNA or RNA), (2) a catalyst

and (3) a compartment. Although not explicitly mentioned, the requirement for energy

production in a model protocell system is essential to drive these processes. To better

understand the origin of life, two strategies can be employed including the top-down

(minimal cell) and the bottom-up (model protocell) approach.5b Hutchinson et al.68 have

attempted the synthesis of a minimal cell by stripping non-essential cellular components

of a Mycoplasma genitalium bacterium while keeping the cell living. It has been


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predicted that the theoretical minimal number of genes required while maintaining

cellular function is approximately 150.69 However, this system is still complex as

hundreds of genes and thousands of different proteins and molecules remain. In order to

synthesise an artificial cell de novo or better understand the origin of life on Earth,

simpler systems should be studied using the bottom-up approach.41

In order to create a living artificial cell from the bottom-up strategy, it requires a

self-replicating vesicle and a replicase (RNA) with RNA-coded activity to allow

synthesis of amphiphilic lipids for compartment division as shown in Figure 1.8.41 This

allows the 'organism' to be self-sustaining and replicate with evolutionary optimisation

as RNA molecules can be encapsulated spontaneously from the surrounding

environment.

Figure 1.8. Outline of a proposed pathway for the synthesis of a living artificial cell. System composed of RNA replicase and self-replicating vesicle with an RNA-coded linking function for lipid synthesis via a ribozyme.41


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Steps towards the development of a living artificial cell have been attempted. To date,

the focus has been mostly on the replication of the vesicle or the genetic information.

Self-reproducing vesicles were first reported by Walde et al.70 in 1994 using vesicles

composed of fatty oleic and caprylic acids. Growth and reproduction was achieved

when a fatty acid precursor (corresponding anhydride) was added to the solution and

subsequently hydrolysed. Sugawara and co-workers have since reported on

self-reproducing vesicles based on artificially developed amphiphiles.71 Giant vesicles

were composed of amphiphiles with an imine group in its hydrophobic chain which are

formed by a dehydrocondensation reaction between the amphiphilic aldehyde and a

lipophilic aniline derivative as shown in Figure 1.9. When the protected aldehyde

precursor was added to the vesicle suspension containing the lipophilic imine and a

catalyst, dehydrocondensation between the two precursors occurred within the interior

pool producing further amphiphiles, contributing to growth of small vesicles and

eventual exocytosis from the parent giant vesicle.

Figure 1.9. Self-reproducing giant vesicles: (i) protected aldehyde precursor (A') is incorporated into the vesicle composed of V and catalyst C produces reactive precursor A; (ii) A reacts with lipophilic imine (B) within the vesicle to form amphiphilic molecule V via dehydrocondensation reaction; (iii) new vesicles composed of V formed; (iv) exocytosis of new vesicles to the bulk water.71


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In a pioneering investigation, Oberholzer et al.72 replicated a RNA strand using all four

nucleosides and a viral RNA polymerase (Q -replicase) in an oleate/oleic acid small

vesicle (diameter <100 nm) capable of vesicle reproduction upon addition of oleic

anhydride precursor. In an elegant system, Kurihara et al.73 have extended this system

by demonstrating self-reproduction of giant vesicles based on artificial amphiphiles

with concomitant amplification of encapsulated DNA (more robust than RNA) as

shown in Figure 1.10. It was shown that amplification of DNA within a

self-reproducible cationic giant vesicle resulted in distribution of the DNA to the

daughter giant vesicles due to electrostatic interactions between DNA and membrane.

Interestingly, it was reported that the amplification of the encapsulated DNA accelerated

the rate of vesicle division.

Figure 1.10. Schematic representation of the amplification of DNA and self-reproduction of giant vesicles.73

As mentioned earlier, it is essential that an ideal model protocell system can produce

energy to drive processes such as replication of DNA/RNA, catalytic functions or

amphiphile synthesis for vesicle replication. Nearly all organisms on Earth source

energy directly or indirectly from the Sun, with the exception of chemoautotrophs that

live in rocks or deep sea hydrothermal vents.74 In plant based photosynthetic organisms,


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this process occurs in the chloroplast and is therefore also an interesting biomimetic

target for artificial cells.

1.5 Artificial Chloroplasts

To a lesser extent, the biomimicry of chloroplasts have been investigated using

synthetic compartments for photoinduced energy storage as an electrochemical gradient.

In a pioneering investigation by Steinberg-Yfrach et al.7a, a synthetic photosynthetic

reaction centre was prepared by linking a tetraarylporphyrin to a naphthoquinone

moiety fused to a norbornene system bearing a carboxylic acid group and a carotenoid

polyene. This molecular 'triad' was incorporated into the bilayer of a

L- -phosphatidylserine and dioleoylphosphatidylcholine (2:3 molar ratio) liposome.

Upon photoexcitation, the triad undergoes photoinduced electron transfer to generate an

intermediate charge-separated species with a reduction potential near the outer surface

and an oxidation potential near the inner surface of the bilayer membrane. A freely

diffusing quinone molecule alternates between its oxidised and reduced forms due to the

induced redox potential allowing passive transport of protons across the membrane

resulting in acidification of the interior as shown in Figure 1.11. An overall quantum

yield of 0.004 was reported. This work has been extended by incorporating CF0F1-ATP

synthase to catalyse the light-driven synthesis of ATP due to the proton gradient.7b


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Figure 1.11. Schematic representation of photoinduced proton translocation across the liposome bilayer.7a

Bhosale et al.8 have investigated transmembrane proton gradients by incorporating rigid

p-octiphenyl rods to create helical tetrameric -stacks of napthalene diimides that can

span egg yolk lipid bilayer membranes as shown in Figure 1.12. The liposomes used

encapsulated quinone molecules as electron acceptors and the bulk solution contained

ethylenediaminetetraacetic acid as electron donors. Upon irradiation, electron transfer

through the helical stacks internally reduced quinone molecules which subsequently

consumed protons leading to basification of the lumen. Of interest, external ligand

intercalation transformed the photoactive scaffolds into ion channels.


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Figure 1.12. Schematic representation of helical tetrameric -stacks of naphthalene diimides in lipid bilayers that can generate a transmembrane proton gradient upon irradiation. Addition of a ligand transforms the photoactive scaffold into ion channels.8

In another elegant system, biosynthesis of ATP was achieved by reconstitution of a

light-driven proton pump (bacteriorhodopsin) and F0F1-ATP synthase in

PEtOz-PDMS-PEtOz triblock copolymer polymersomes.9 Reconstitution of the proton

pump results in an influx of protons into the vesicle which drives the formation of ATP

as shown in Figure 1.13. Montemagno and co-workers fine-tuned the insertion process

to allow orientational control of bacteriorhodopsin reconstitution.


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Figure 1.13. Schematic representation of proteopolymersomes reconstituted with bacteriorhodopsin (BR) and F0F1-ATP synthase capable of ATP synthesis upon irradiation.9


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1.6 Project Description and Goal

The goal of this project is to develop light-active bioconjugates and incorporate them

into vesicular membranes to create a light-harvesting electron transport chain capable of

generating a transmembrane proton gradient. The light-activation would be a means to

mimic chloroplasts by photoinducing the proton gradient. The design for a

light-activated bioconjugate and the primitive chloroplast is shown in Figure 1.14.

Figure 1.14. Schematic of proposed project goal. (a) a primitive chloroplast capable of generating a transmembrane proton gradient by reconstitution of a light-harvesting bioconjugate with natural electron acceptor and proton pump, cytochrome c oxidase (CcOx). (b) light-harvesting bioconjugate, ruthenium(II)-bisterpyridine cytochrome c(Ru-cyt c).


1-26

The enzyme cytochrome c was chosen for this system due to several advantages it offers

including: (1) a redox metalloprotein, (2) it has been intensively studied and its structure

and photophysical properties are well-known75, (3) it has characteristic UV-Vis

absorption changes corresponding to reduced and oxidised states as well as changes to

protein structure35b, (4) applicable to various bioconjugation techniques because it is

stable in an array of reaction conditions and is highly soluble35b,76, (5) can be purified

readily in large quantities and (6) low cost.

In particular, the iso-1 form of yeast cytochrome c is attractive as a target as it

possesses a single cysteine residue that allows site-specific modification under benign

aqueous conditions using maleimides to form a stable thioester.15 The position of the

cysteine residue is on the opposite side of the active site for cytochrome c oxidase

binding and therefore allows modification without adversely affecting biological and

catalytic function. It has been reported that the cysteine residue is located in a

predominantly hydrophobic pocket which is less accessible to solvents and results in

slower reaction rates, however, reaction conditions have been previously optimised in

the Thordarson group.35b The bioconjugation of iso-1 cytochrome c and a brief review

of bioconjugate techniques and methods have been discussed further in Chapter 3.

Ruthenium(II)-bisterpyridine chromophores have been heavily studied for both

the methods of their synthesis and photophysical properties.77 Ruthenium terpyridine

complexes are often compared to the related bipyridine moieties in terms of their

photophysical properties. Tris(bipyridine)ruthenium(II) complexes are often used for

photophysical studies due to high quantum yields, long-lived luminescence lifetimes

and luminescence can be measured at room temperature.78 Ruthenium bipyridine

complexes have been reported to exhibit room temperature quantum yields and

luminescence lifetimes of 0.04 and 0.6 s, respectively.79 On the other hand, ruthenium


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terpyridine complexes display a lack of room temperature luminescence and low yields

with values of 1 ns and 10-5, respectively, which require quantum yield and lifetime

studies to be performed at 77 K.78

Although ruthenium terpyridine complexes have short-lived room temperature

luminescence lifetimes, the light-harvesting chromophore chosen was based on these

bisterpyridine complexes. The complexes allow (1) assembly of ligands to form linear

complexes without introducing chirality (as observed with bipyridine complexes)77, (2)

the synthesis and photophysical properties of these complexes are heavily studied and

well-known78, (3) photoinduced electron transfer studies involving bisterpyridine

complexes as donor-acceptor systems have been reported80 and (4) electron transfer in

ruthenium(II)-bisterpyridine cytochrome c bioconjugates have been previously

reported.35a A brief review and methods of ruthenium terpyridine complex synthesis is

discussed further in Chapter 2.

As an extension to the synthetic donor-biological acceptor concept used here,

the opposite biological donor-synthetic acceptor motif was also investigated. Following

recent reports by Bogdanov et al.81 that green fluorescent protein (GFP) may act as a

light-induced electron donor, a GFP-synthetic acceptor bioconjugate was constructed to

investigate electron transfer of these donor-acceptor constructs which is discussed

further in Chapter 4.

The use of liposomes and polymersomes has been heavily studied for enzyme

encapsulation.5b,43b In particular, polymersomes based on the diblock copolymer

polystyrene-b-poly(acrylic acid) (PS-b-PAA) was used as the compartments to mimic

chloroplasts in this Thesis. PS-b-PAA offers the following advantages: (1) well studied

polymers for aggregate formation52, (2) improved stabilities and robustness compared to

liposomes48 and (3) morphologies can be fine-tuned by the addition of additives or


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preparation method.82 A brief review of lipid and polymer vesicles and their methods of

preparation are further discussed in Chapter 5.

The photoinduced electron transfer studies of ruthenium(II)-bisterpyridine

cytochrome c bioconjugates and the effect of membrane encapsulation is described in

Chapter 6. Finally, the combination of the light-activated bioconjugate, proton pump

(cytochrome c oxidase) and reconstitution in a polymer membrane to construct a

semi-synthetic electron transport chain for photoinduced transmembrane proton

gradients is described in Chapter 7.

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(56) van Dongen, S. F. M.; Nallani, M.; Schoffelen, S.; Cornelissen, J. J. L. M.; Nolte, R. J. M.; van Hest, J. C. M. Macromol. Rapid Commun. 2008, 29, 321.

(57) Palivan, C. G.; Fischer-Onaca, O.; Delcea, M.; Itel, F.; Meier, W. Chem. Soc. Rev. 2012, 41, 2800.

(58) Nardin, C.; Widmer, J.; Winterhalter, M.; Meier, W. Eur. Phys. J. E 2001, 4,403.

(59) Nardin, C.; Thoeni, S.; Widmer, J.; Winterhalter, M.; Meier, W. Chem. Commun. 2000, 1433.

(60) Nallani, M.; Benito, S.; Onaca, O.; Graff, A.; Lindemann, M.; Winterhalter, M.; Meier, W.; Schwaneberg, U. J. Biotechnol. 2006, 123, 50.

(61) Broz, P.; Driamov, S.; Ziegler, J.; Ben-Haim, N.; Marsch, S.; Meier, W.; Hunziker, P. Nano Lett. 2006, 6, 2349.

(62) Grzelakowski, M.; Onaca, O.; Rigler, P.; Kumar, M.; Meier, W. Small 2009, 5,2545.

(63) Kim, K. T.; Cornelissen, J. J. L. M.; Nolte, R. J. M.; van Hest, J. C. M. Adv. Mater. 2009, 21, 2787.

(64) Kreft, O.; Prevot, M.; Möhwald, H.; Sukhorukov, G. B. Angew. Chem. Int. Ed.2007, 46, 5605.

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Chapter 2

Synthesis of Ru(II)-Complexes and Quinone Derivatives

Chapter 2 Synthesis of Ru(II)-Complexes and Quinone derivatives

2-2

2 Synthesis of Ru(II)-Complexes and Quinone Derivatives

In this Chapter, the synthesis of a number of terpyridine, ruthenium(II)-bisterpyridine

complexes and maleimide functionalised ruthenium(II)-bisterpyridine complexes for use

as photoactive electron donors are discussed. The compounds described herein were

synthesised for the attachment to cytochrome c to form light-activated bioconjugates, as

spacers for homo/hetero dimeric bioconjugates or reference compounds for studies

described elsewhere in this thesis. The 4’-aryl functionalised 2,2’:6’,2”-terpyridines

were synthesised using central ring-assembly methodologies based on literature

procedures.1 For this project, methods that lead to the formation of both

homo/heteroleptic complexes which allow asymmetric and dimeric bioconjugation are

desired.

Additionally, quinone derivatives, with focus on anthraquinone analogues were

synthesised as potential electron acceptors. Anthraquinone derivatives were synthesised

as constructs for covalent attachment to green fluorescent proteins (GFP) to form

potential light-activated bioconjugates. Analogues were synthesised using

N-hydroxysuccinimide/N,N’-dicyclohexylcarbodiimide (NHS/DCC) peptide coupling

chemistry2 to introduce azide functionality for attachment to alkyne-tagged GFP via

Huisgen azide-alkyne 1,3-dipolar cycloaddition.3

The procedures for synthesis of terpyridines, complexation and anthraquinone

derivatives are briefly reviewed in this chapter.

2.1 Methods For Functionalising 2,2’:6’,2”-Terpyridines

Terpyridines (tpy) were first discovered by Morgan and Burstall4 who reported its

synthesis as a byproduct in the synthesis of 2,2’-bipyridine (bpy) which was known 44

years earlier (Figure 2.1).5 The synthesis was achieved by heating pyridine with


2-3

anhydrous iron(III) chloride at 340 oC in an autoclave (50 atm) for 36 h. The authors

reported interesting spectroscopic properties of terpyridine with Fe(II) indicating metal

complex formation.

Figure 2.1. Structures of 2,2'-bipyridine (bpy) and 2,2':6',2"-terpyridine (tpy).

Since the first report of terpyridines, tremendous improvements in synthetic strategy to

optimise yields and routes towards site-specific functionalisation have occurred in the

last 80 years. Modern terpyridine synthesis can be categorised into two basic synthetic

routes, which are central ring assembly and cross-coupling procedures.1b,6

Ring assembly strategies toward terpyridine synthesis have been the most

predominant in recent decades. The most common methods developed within the ring

assembly route are the Kröhnke reaction7, Potts methodology8 and Jameson

methodology.9 The most well-known is the Kröhnke reaction (Scheme 2.1) which

involves Michael addition of N-heteropyridinium salts (L-1) with an , -unsaturated

enone (L-2) to yield a 1,5-dione intermediate. The intermediate is subsequently cyclised

using a nitrogen source such as ammonium acetate.10


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Scheme 2.1. Kröhnke synthesis of 4’-functionalised 2,2’:6’,2”-terpyridine. (a) NH4OAc.

An alternative approach to synthesise functionalised terpyridines is the Potts

methodology (Scheme 2.2), also known as the -oxoketene dithioacetal methodology.

In general, the reaction between the enolate of 2-acetylpyridine with carbon disulfide

and methyl iodide gives the resulting -oxoketene dithioacetal (L-3) product.

Subsequent Michael addition with an additional 2-acetylpyridine gives thiol

functionalised 1,5-dione (L-4). Cyclisation is similarly achieved with a nitrogen source

such as ammonium acetate.11

Scheme 2.2. Potts synthesis of 4'-functionalised 2,2':6',2"-terpyridines. (a) i) t-BuOK, THF, ii) CS2, iii) MeI. (b) t-BuOK, 2-acetylpyridine. (c) NH4OAc, HOAc.

Lastly, a less frequently used method to synthesise terpyridines is the Jameson method

(Scheme 2.3), due to the lack of functionalisation ability. Jameson and Guise first


2-5

reacted 2-acetylpyridine with N,N-dimethylformamide dimethyl acetal to form the

enaminone (L-5). The enaminone (L-5) was condensed with a second equivalent of

2-acetylpyridine resulting in a loss of dimethylamine to yield the 1,5-dione (L-6) and

ring closed with ammonium acetate.9

Scheme 2.3. Jameson synthesis of 2,2’:6’,2”-terpyridine. (a) N,N-dimethylformamide dimethyl acetal. (b) 2-acetylpyridine, t-BuOK. (c) NH4OAc, HOAc.

The disadvantages of these ring assembly methods are that the final condensation step

yields tar-like by-products resulting in purification difficulties.1a Over the last decade,

cross-coupling procedures based on modern Pd(0)-catalysed coupling reactions6 have

improved efficiency and synthetic simplicity with substitution control over traditional

methods.12 Pd(0) catalysed cross-coupling procedures include Suzuki13, Negishi14 and

Stille15 couplings.

In this project, a step-wise modified Kröhnke synthesis is employed which

avoids the common drawback of tarry crude by-products and achieves straightforward

synthesis. Chemical modification at the 4’-position of the terpyridine core with aromatic

groups allows the introduction of functional groups while maintaining achirality,


2-6

improved yields16 and increased excited state lifetimes17 making them suitable

photoactive electron donors when coordinated to metal ions such as Ru(II).

2.2 Functionalised 4’-Aryl Terpyridines

2.2.1 Synthesis of nitro-phenyl-terpyridine 3

In order to synthesise functionalised terpyridines with higher yields and ease of

synthesis, nitro-phenyl-terpyridine 3 was synthesised based on a modified classical

Kröhnke synthesis. The modified Kröhnke synthesis is achieved via the azachalcone,

pryidinium iodide route (Scheme 2.4) for terpyridine 3 which prevents the formation of

crude tarry by-products and allows simplified purification procedures based on previous

alternative methods explored by the Thordarson group.18

As shown in Scheme 2.4, the formation of nitro-substituted terpyridine 3 was

carried out in a three-step convergent synthesis with isolation of the intermediates.

Synthesis of 4-nitro-2’-azachalcone 1 via Claisen-Schmidt condensation between one

equivalent of 4-nitrobenzaldehyde and 2-acetylpyridine in a mixture of aqueous sodium

hydroxide and methanol gave a moderate yield of 28%.19 Subsequently, 1-(2-oxo-2-(2-

pyridyl)ethyl)pyridinium iodide 2 was synthesised by addition of 2-acetylpyridine to a

warm solution (60 oC) of iodine in dry pyridine. The mixture was then heated at 100 oC

and gave acylpyridinium iodide 2 in good yield of 79%.20

Finally, reacting the precursors azachalcone 1 with pyridinium iodide 2 and

subsequent ring closure with ammonium acetate in refluxing dry methanol afforded

terpyridine 3 in 75% yield.20


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Scheme 2.4. Synthesis of 4’-(4-nitrophenyl)-terpyridine 3 by the azachalcone/pyridinium iodide route. (a) MeOH, r.t., 29%. (b) dry pyridine, 60 -100 oC, 79%. (c) NH4OAc, dry MeOH, reflux, 75%.

2.2.2 Synthesis of amino-phenyl-terpyridine 4

In order to further functionalise terpyridine 3, an aniline functional group was

introduced to allow further chemical modification with peptide coupling chemistry to

introduce maleimide functionality. Nitro-phenyl-terpyridine 3 was reduced using

previously reported literature procedures according to Scheme 2.5.21


2-8

Scheme 2.5. Synthesis of 4’-(4-aminophenyl)-2,2’:6’,2”-terpyridine by reduction with hydrazine over a Pd/C catalyst. (a) 10% Pd/charcoal, hydrazine monohydrate, EtOH, reflux, 81%.

Terpyridine 3 was activated in refluxing absolute ethanol over a 10% Pd/charcoal

catalyst for 45 min and subsequently hydrazine monohydrate was added to generate the

reduced aniline terpyridine 4 in 81% yield. Upon reduction, the purple terpyridine 3

changed to a yellow crystalline solid 4. It should be noted that in order to maximise

product yield, workup of the crude should be filtered over celite while the mixture is

warm due to formation of microcrystalline solid 4 upon cooling leading to loss of

product.

2.3 Functionalised Ru(II)-Bisterpyridine Complexes

Mononuclear bisterpyridine complexes are well-known and documented in the

literature.1b,17 In general, metal complexes of the type [M(tpy)2]X2, where X = counter

ion (PF6-, Cl- or ClO4

-) are formed from low oxidation state d-block transition metals.

Traditionally, these complexes are prepared from Zn(II), Co(II), Cu(II), Ni(II) transition

metals and the stability of these metal bis-complexes are a result of the strong

metal-ligand (d- *) back-donation, combined with the dynamic chelate effect.22

Complexation with transition metals results in a distorted octahedral geometry23 due to

the hexa-coordinate nature of most d-block metals.


2-9

Synthetic strategies towards the assembly of complexes are dependent on the

type of desired complex (homo/heteroleptic). Homoleptic complexes can be synthesised

in a straightforward one-pot synthesis, where the terpyridine ligand is treated with the

desired metal ion in a 2:1 (ligand:metal) ratio. The resulting complexes can be purified

in a two-step procedure involving the precipitation of symmetric complex with counter

ions (NH4PF6 or TBA-Cl) followed by recrystallisation.24

On the other hand, the preparation of heteroleptic complexes requires a directed

strategy where the desired ligands are introduced in a two-step procedure to prevent

“scrambling” which involves the exchange of ligands. Directed synthesis requires the

formation of the mono-complex which forms a highly insoluble product and is not

characterised by 1H nuclear magnetic resonance (NMR) spectroscopy due to its

paramagnetic nature. The second ligand is then introduced to the mono-complex under

reductive conditions in an alcoholic solvent to yield the asymmetric complex.25 This

technique is generally only applicable to Ru(II), Ir(III) and Os(II) complexes as a result

of the weaker stabilities of other transition metal complexes at elevated temperatures

leading to scrambling.1b In general, heteroleptic complexes require purification over

silica or alumina using a highly polar mobile phase.25-26

Characterisation of metal bisterpyridine complexes can be achieved using a

combination of techniques including 1H NMR and UV-Vis spectroscopy as well as soft

mass spectrometry techniques (MALDI-TOF or Electrospray). Due to terpyridine

ligands possessing anti orientation, 1H NMR spectroscopy can be used to characterise

complexation by monitoring the upfield shift in the 6,6”-proton resonances as a result of

anti to syn ligand orientation upon complexation and the metal-ligand bond.6 UV-Vis

spectroscopy allows confirmation of complexation by the bathochromic shift of the

ligand-centred (LC) band and appearance of the metal-ligand charge transfer (MLCT)


2-10

band giving rise to intense colours of the compounds.27 Soft mass spectrometry

techniques such as MALDI-TOF or electrospray ionisation are ideal to analyse metal

bisterpyridine complexes as it allows the complexes to be detected without

fragmentation while retaining their isotopic patterns. However, when analysing Ir(III)

complexes, MALDI-TOF may cause fragmentation as the MLCT band overlaps the

excitation wavelength (337 nm) of the MALDI-TOF laser.28

In recent decades, metal bis-complexes of Ru(II), Os(II) and Ir(III) metal ions

have been of primary interest, in particular, Ru(II)-complexes due to their interesting

photophysical, electrochemical and photochemical properties. In this project,

functionalised 4’-aryl Ru(II)-complexes have been selected for study due to the MLCT

band taking place in the visible region allowing photo-excitation by visible light,

straightforward synthesis arising from achirality of 4’-functionalisation and their

improved excited state lifetimes after modification with 4’-aryl groups17 make them

ideal photosensitisers for attachment to cytochrome c as light-activated bioconjugates.

2.3.1 Synthesis of [Ru(tpy)]Cl3 5 and [Ru(tpy)2](PF6)2 6

Synthesis of asymmetric mono-complex precursor 5 and reference complex 6 was

achieved following modified literature methods (Scheme 2.6).29 Ru(II) mono-complex 5

was prepared by refluxing 2,2’:6’,2”-terpyridine and two molar equivalent of

ruthenium(III) trichloride hydrate in absolute ethanol overnight yielding precursor 5 as a

dark black solid in 88%. This compound was selected for forming asymmetric

complexes with maleimide functionalisation via the two-step directed methodology.


2-11

Scheme 2.6. Synthesis of [Ru(tpy)]Cl3. (a) EtOH, reflux, 88%.

In order to synthesise the reference complex 6 for photo-reaction control experiments

(Chapter 6), ruthenium(III) trichloride hydrate and two molar equivalent

2,2’:6’,2”-terpyridine was heated at 110 oC in ethylene glycol overnight in one-pot

(Scheme 2.7). The [Ru(tpy)2]2+ chloride salt was diluted in water and precipitated with

ammonium hexafluorophosphate. Complex 6 was recrystallised with acetonitrile/diethyl

ether to afford the red solid in 50% yield. Synthesis of complex 6 was achieved by a

modified approach, substituting N-ethylmorpholine in refluxing absolute ethanol with

neat ethylene glycol which is discussed further in the next section (Chapter 2.3.2).

Scheme 2.7. Synthesis of [Ru(tpy)2](PF6)2. (a) Ethylene glycol, 110 oC, 50%.


2-12

2.3.2 Synthesis of [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)](PF6)2 7

Asymmetric aniline functionalised complex 7 was prepared using a directed synthesis

approach. Initially, asymmetric aniline complex 7 was prepared using mono-complex

precursor 5 (Scheme 2.8). The synthesis of [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-

tpy)](PF6)2 7 was achieved by reacting aniline 4 with precursor 5 in refluxing absolute

ethanol overnight. The resulting product was filtered over celite and diluted with water.

Subsequently, complex 7 was precipitated with ammonium hexafluorophosphate and

recrystallised with acetonitrile/diethyl ether to afford red complex in 28% yield. 1H

NMR spectroscopy confirmed complexation due to the upfield shift of the 6,6”-protons

as a result of anti to syn conformation change of the terpyridine ligand as shown in

Figure 2.2.6

Scheme 2.8. Synthesis of [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)](PF6)2. (a) EtOH, reflux, 28%. (b) Ethylene glycol, 110 oC, 76%.


2-13

Figure 2.2. 1H NMR (300 MHz) spectra of aniline terpyridine 4 (top-blue-CDCl3) and asymmetric aniline complex 7 (bottom-black-CD3CN) showing upfield shift of the 6,6”-protons upon complexation due to change in anti to syn orientation of the terpyridine ligands.6 Complex 7 NH2 signal suppressed due to slow proton exchange of labile protons with CD3CN. Symbols represent 6,6”-protons.

Generally, the mono-complex 5 is reacted with the second ligand in ethanol (reductant)

containing N-ethylmorpholine30 (catalyst) and refluxed to afford the bisterpyridine

complex. However, previous studies in the Thordarson group showed that in the

presence of N-ethylmorpholine, symmetric complexation of aniline terpyridines resulted

in N-alkylation of the ligand.18

As a result of the poor yield due to the removal of catalyst in the complexation

reaction of aniline 7, the reaction was repeated using an alternative method based on

Ir(III)-bisterpyridine literature.1b Higher yields after complexation were observed when

performing the reaction in ethylene glycol. The synthesis of complex 7 was achieved by

reacting aniline 4 with mono-complex 5 in ethylene glycol at 110 oC. Subsequently, the

solution was filtered over celite and precipitated with ammonium hexafluorophosphate.


2-14

The resulting precipitate was centrifuged, washed with water and collected with

acetonitrile.

Initially, purification by recrystallisation was attempted; however, purification

by crystallisation for complex 7 was not possible due to scrambling of the ligands as a

result of harsher reaction conditions. Purification of the crude mixture was achieved by

separation over silica (gradient from 90:9:1 to 20:3:1, acetonitrile:water:saturated

potassium nitrate (v/v/v)), followed by a second neutral alumina column (gradient from

acetonitrile to 90:9:1, acetonitrile:water:saturated potassium nitrate (v/v/v)) to afford red

crystalline solid in 76% yield.

It was found that purification by chromatography over silica or alumina (neutral)

required highly polar solvents containing salts. This was a result of binding of the

complex to the stationary phase. Additionally, characterisation of purified fractions

proved difficult at times as thin layer chromatography (TLC) analysis showed

appearance of multiple bands due to counter ion exchange after purification.

2.3.3 Synthesis of [Ru(tpy)(maleimide-hexylcarboxamido-phenyl-tpy)](PF6)2 8

Following the preparation of the amino functionalised complex 7, maleimide modified

complexes were prepared for attachment to cytochrome c and other surface cysteine

containing proteins. Maleimide was introduced to complex 7 by peptide coupling

chemistry using a route similar to that of Hovinen31 due to the unreactive nature of

anilines preventing the use of classical N-hydroxysuccinimide/N,N’-

dicyclohexylcarbodiimide (NHS/DCC) coupling conditions.

Maleimide complex 8 was prepared by reacting a solution of

6-maleimidocaproic acid, O-(7-azabenzotriazole-1-yl)-1,1,3,3-tetramethyluronium

hexafluorophosphate (HATU) and N,N-diisopropylethylamine (DIPEA) in dry


2-15

N,N-dimethylformamide was stirred at room temperature for 1 h (Scheme 2.9). Aniline

7 in dry dimethylformamide was then added to the stirring solution and stirred for a

further 26 h at room temperature. Dichloromethane was added to the solution and the

organic phase was washed with aqueous citric acid (10% w/v), water and dried with

anhydrous sodium sulphate. Dichloromethane was removed in vacuo and the

concentrated dimethylformamide phase was precipitated into dry diethyl ether, filtered

and further washed with diethyl ether. The crude mixture was purified over silica using

a gradient from acetonitrile to 70:29:1 acetonitrile:water:saturated potassium nitrate and

precipitated with ammonium hexafluorophosphate yielding the complex 8 as a red

crystalline solid in 41% yield.

Scheme 2.9. Synthesis of [Ru(tpy)(maleimide-hexylcarboxamido-phenyl-tpy)](PF6)2.(a) O-(7-azabenzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU), N,N-diisopropylethylamine, dry N,N-dimethylformamide, r.t., 41%.


2-16

2.3.4 Synthesis of [Ru(4’-(4-aminophenyl)-2,2’:6’2’’-terpyridine)2](PF6)2 9

In order to prepare dimeric bioconjugates, symmetric aniline complex 9 was similarly

prepared using a modified literature method from Ng et al.27 Ruthenium(III) trichloride

hydrate and two molar equivalent of aniline terpyridine 4 in ethylene glycol was heated

at 110 oC overnight (Scheme 2.10). Subsequently, the mixture was diluted with water

and filtered over celite. The product was precipitated using ammonium

hexafluorophosphate and the fine precipitate was collected by centrifugation followed

by washing with water. Purification of the crude mixture was achieved by

recrystallisation in acetonitrile/diethyl ether to yield red crystals in 52% yield.

Scheme 2.10. Synthesis of [Ru(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)2](PF6)2. (a)Ethylene glycol, 110 oC, 52%.


2-17

2.3.5 Synthesis of [Ru(4’-(4-maleimide-hexylcarboxyamido-phenyl)-2,2’:6’2’’-

terpyridine)2](PF6)2 10

Bis-maleimide complex was prepared similarly to compound 8 in order to prepare

dimeric bioconjugates with proteins containing surface exposed cysteines. A solution of

6-maleimidocaproic acid, HATU and DIPEA in dry N,N-dimethylformamide was

stirred at room temperature for 1 h (Scheme 2.11). Aniline 9 in dry dimethylformamide

was then added to the stirring solution and stirred for a further 25 h at room

temperature. Dichloromethane was added to the solution and the organic phase was

washed with aqueous citric acid (10% w/v), water and dried with anhydrous sodium

sulphate. Dichloromethane was removed in vacuo and the concentrated

dimethylformamide phase was precipitated into dry diethyl ether, filtered and further

washed with diethyl ether. The crude mixture was purified over silica using a 20:1:3

acetonitrile:water:saturated potassium nitrate mobile phase and precipitated with

ammonium hexafluorophosphate yielding the complex 10 as a red crystalline solid in

27% yield.


2-18

Scheme 2.11. Synthesis of [Ru(maleimide-hexylcarboxamido-phenyl-tpy)2](PF6)2. (a) O-(7-azabenzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU), N,N-diisopropylethylamine, dry N,N-dimethylformamide, r.t., 27%.

2.3.6 Spectroscopic properties of Ru(II)-bistepyridine complexes

Characterisation of metal bisterpyridine complexes can be achieved by complementary

techniques including 1H NMR spectroscopy and mass spectrometry. However, UV-Vis

spectroscopy can be utilised to characterise complex formation in a straightforward

manner. Complexes of [Ru(tpy)2]2+-type display a characteristic spin allowed d- *

metal-ligand charge transfer (MLCT) band as well as ligand centred (LC) absorption

bands (Figure 2.3).22 Successful formation of terpyridine metal complexes is usually

accompanied by a bathochromic (red) shift of the LC absorption band and appearance

of a new MLCT band.


2-19

Figure 2.3. UV-Vis spectra of (a) terpyridines and 4’-aryl terpyridines in dichloromethane, (b) Ru(II)-bisterpyridine and functionalised complexes in acetonitrileshowing bathochromic shifts of ligand centred (LC) bands and appearance of ametal-ligand charge transfer (MLCT) band.

Comparison of [Ru(tpy)2]2+-type complexes of aniline 7 and 9 as well as maleimide

functionalised complexes 8 and 10 shows bathochromic shift of the LC band at ca.

310 nm compared to the ca. 280 nm absorption band of free terpyridine ligand and

functionalised ligands 3 and 4. These bands correspond to - * transitions with

electrons of the aromatic systems (LC). Additionally, appearance of a new absorption

band of the ruthenium(II) complexes is observed centred ca. 480-500 nm corresponding

to the MLCT bands, which relates to the excitation of an electron from the

metal-centred d-orbitals to an unfilled ligand-centred *-orbital.32

From Table 2.1, it can be seen that both asymmetric and symmetric aniline

complexes 7 and 9 exhibit significant red shifting of the MLCT band, 490 and 504 nm

respectively, compared to prototype complex 6 of 476 nm. This is a result of the

introduction of electron donating substituents which induce red shifting of absorption

bands due to decrease of the gap between the LUMO (ligand *) and HOMO (metal t2g,

distorted octahedral) orbitals after modification.17 Additionally, this effect is observed

with the asymmetric and symmetric maleimide functionalised complexes 8 and 10 with

MLCT bands red shifted at 485 and 494 nm, respectively. Finally, Table 2.1 shows a


2-20

general trend of increasing extinction coefficient after introducing phenyl substituents in

the 4’-positions for complexes 7-10 which is as expected based on simple theoretical

predictions.33

Table 2.1. Absorption data for prototype [Ru(tpy)2]2+ complex and 4’-functionalised analogues

Complex Absorption /nma Extinction coefficient /M-1cm-1

[Ru(tpy)2]2+ 6 476 17 70017

[Ru(tpy)(tpyNH2)]2+ 7 490 17 100[Ru(tpyNH2)2]2+ 9 504 31 800[Ru(tpy)(tpymal)]2+ 8 485 26 000[Ru(tpymal)2)]2+ 10 494 41 50018

a Data from 10-6 M solutions in acetonitrile solution

2.3.7 Crystallography

As further evidence of complex formation, single crystal structures of asymmetric

complexes 7 and 8 were obtained providing detailed structural analysis of resulting

complexes and the effects of 4’-aniline and maleimide substitution on geometries and

bond lengths. X-ray crystallography analysis was performed by Dr. Mohan Bhadbhade

(UNSW) with analysis of aniline 7 performed in-house at the UNSW Analytical Centre

and maleimide 8 at the Australian Synchotron facility.

2.3.7.1 [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)](PF6)2 7

The crystal structure of complex 7 dimethylformamide solvate with the composition

[Ru(C15H11N3)(C21H16N4)](PF6)2 2C3H7NO was crystallised by diffusion of diethyl

ether vapour into a solution of 7 in N,N-dimethylformamide in the triclinic space group,

P 1 as red blades. As shown in Figure 2.4, crystal 7 displays an orthorhombic distortion

from octahedral symmetry (N2A-Ru-N3A 78.8(5)o) which is as expected and observed

in [Ru(tpy)2](PF6)2.23


2-21

Figure 2.4. An ORTEP plot of complex 7 2C3H7NO (20% thermal ellipsoids for non-hydrogen atoms). Hydrogen atoms omitted for clarity. Triclinic space group, P1 .

In Table 2.2, the aniline shows a significant twist with a torsional angle of 29.1(2)o

(C7B-C8B-C18B-C17B) about the interannular bond, greater than the corresponding

twist found in the free 4’-phenyl terpyridine ligand.34 Ru-N bond lengths between

ligands A and B are equivalent, 2.033(7) Å and is within reported ranges for

[Ru(tpy)2](PF6)2.23 The C-C and C-N bond lengths within the aromatic rings are normal

and average 1.380(1) and 1.355(6) Å, respectively. The interannular C-C bond distances

of ligands A and B average 1.464(5) and 1.474(7) Å and are consistent with reported

asymmetric complex [Ru(tpy)(4’-5-carboxypentyl-tpy)](PF6)2 of 1.471(1) Å.29

Table 2.2. Selected bond lengths (Å) and angles/torsions (o) for complex 7 2C3H7NO with estimated standard deviations in parentheses.Ru-N1A 2.061(3) Ru-N1B 2.061(3)Ru-N2A 1.980(3) Ru-N2B 1.971(3)Ru-N3A 2.061(3) Ru-N3B 2.065(3)C5A-C6A 1.463(4) C5B-C6B 1.472(4)C10A-C11A 1.466(4) C10B-C11B 1.483(4)

C8B-C18B 1.469(5)

N2A-Ru-N3A 78.8(5) C7B-C8B-C18B-C17B 29.1(2)


2-22

From the packing diagram in Figure 2.5, it shows the complex cation forming

anti-parallel columnar stacks along the c-axis while forming alternating layers with

hexafluorophosphate anions and N,N-dimethylformamide solvent molecules filling the

voids. Intermolecular contact within the columns are face-to-face and edge-to-face

-stacking at a distance of 3.8 Å (N1B and N3B) and 3.6 Å (N1A and N1B),

respectively, in adjacent unit cells.

Figure 2.5. Packing diagram obtained from single-crystal X-ray diffraction analysis of complex 7 2C3H7NO as a capped stick representation. Counter-ions and dimethylformamide omitted for clarity. Representation viewed along b-axis.

2.3.7.2 [Ru(tpy)(maleimide-hexylcarboxamido-phenyl-tpy)](PF6)2 8

The crystal structure of complex 8 dimethylformamide water solvate with the

composition [Ru(C15H11N3)(C31H27N5O3)](PF6)2 2C3H7NO H2O was crystallised by

diffusion of diethyl ether vapour into a solution of 8 in N,N-dimethylformamide in the

monoclinic space group, P21/c as thin red blades. Structural characteristics of complex

8 are similar to complex 7. The main notable difference is the reduction in the torsion

angle between 4’-aryl maleimide group and the central pyridyl ring of ligand B with an

angle of 23.9(8)o as shown in Table 2.3 and Figure 2.6a.


2-23

Table 2.3. Selected bond lengths (Å) and angles/torsions (o) for complex 8 2C3H7NO H2O with estimated standard deviations in parentheses.Ru-N1A 2.076(4) Ru-N1B 2.061(4)Ru-N2A 1.976(4) Ru-N2B 1.985(4)Ru-N3A 2.068(4) Ru-N3B 2.060(4)C5A-C6A 1.458(5) C5B-C6B 1.481(5)C10A-C11A 1.467(5) C10B-C11B 1.462(5)

C8B-C18B 1.486(6)

N2A-Ru-N3A 78.7(9) C7B-C8B-C16B-C17B 23.9(8)

Based on the packing diagram as shown in Figure 2.6b, it can be seen that complex 8 is

stacked in an interlaced anti-parallel motif along the b-axis, possibly to accommodate

the steric hindrance of the flexible maleimide spacer group. Intermolecular contact

within the columns along the a-axis of the crystal structure is face-to-face and

edge-to-face -stacking at a distance of 3.7 Å (N1A and N3A) and 3.3 Å (N1A and

N1B), respectively, coupled with twisted parallel-displaced -stacking between the

outer pyridyl ring (N3B) and 4’-phenyl ring at a distance of 3.8 Å in adjacent unit

cells. Hexafluorophosphate anions form alternating layers with complex 8 along the

a-axis and solvent molecules fill the voids in the packed structure.


2-24

Figure 2.6. Single-crystal X-ray diffraction structures of complex 8 2C3H7NO H2O. (a) an ORTEP plot (20% thermal ellipsoids for non-hydrogen atoms). Hydrogen atoms omitted for clarity. (b) Packing diagram as a capped stick representation. Counter-ions and solvent molecules omitted for clarity. Representation viewed along a-axis.Monoclinic space group, P21/c.


2-25

2.4 Functionalised Anthraquinone-Based Acceptors

In addition to light-activated electron donors based on metal bisterpyrdines, azide

functionalised anthraquinones were prepared as potential ‘clickable’ electron acceptors

to light-activated biological electron donors such as green fluorescent proteins (GFP).

Electron acceptors of the quinone family are one of the most widely studied

acceptors17,26 due to the electron transfer roles of quinones in photosynthesis and

respiration.35 Quinones are highly suitable as electron acceptors due to their electron

shuttling behaviour allowing reversibility between both stable redox states via a two

electron process as shown in Scheme 2.12.36 In these studies, anthraquinones were

selected due to their higher stabilities compared to p-benzoquinones and functionalised

p-benzoquinones which can degrade to form quinhydrones37 when isolated or stored in

solution for prolonged periods.

Scheme 2.12. Stable reversible conversion of oxidised/reduced anthraquinone (AQ) states via a two electron redox process.

Modifying proteins chemically can be difficult due to an array of potential functional

groups including amines, carboxylic acids, histidines and thiols. The Cu(I)-catalysed

[3+2] Huisgen cycloaddition of azides and terminal alkynes, also known as the

azide/alkyne click reaction, is ideal due to benign reaction conditions as well as

chemoselectivity.38 As such, azide functionalised anthraquinones were selected as

synthetic targets in this project to functionalise light-activated biological electron

donors.


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2.4.1 Synthesis of 1-amino-3-azidopropane 11

In order to introduce azide functionality to anthraquinone, a short chain spacer

containing amide and azide functional groups was synthesised to allow attachment to

anthraquinone via peptide coupling chemistry. The 1-amino-3-azidopropane spacer 11

was synthesised based on a previously reported method.39

The azide spacer 11 was prepared by reacting 1-amino-3-bromopropane

hydrobromide with four molar excess of sodium azide in refluxing water via a

displacement reaction for 23 h (Scheme 2.13). Subsequently, the aqueous mixture was

cooled to 0 oC, extracted with diethyl ether and the organic phase was dried with

anhydrous sodium sulphate. The organic phase was removed in vacuo to afford the pure

yellow oil 11 in 43% yield.

Scheme 2.13. Synthesis of 1-amino-3-azidopropane. (a) NaN3, H2O, reflux, 43%.

2.4.2 Synthesis of anthraquinone-2-azidopropylamide 13

Following the synthesis of azide spacer 11, the azide modified anthraquinone 13 was

then prepared using peptide coupling methods based on NHS/DCC chemistry. The

purpose of either maleimide modification using an N-propargyl maleimide spacer to

attach to GFP or direct attachment to an alkyne-tagged GFP using click chemistry is the

formation of a light activated donor-acceptor system based on GFP-anthraquinone,

which is further discussed in Chapter 2.4.3 and Chapter 4.

Azide 13 was prepared by initially preparing the N-hydroxysuccinimide (NHS)

ester of anthraquinone-2-carboxylic acid (Scheme 2.14). Anthraquinone-2-carboxylic

acid and NHS was cooled to 0 oC in dry dichloromethane and

N,N’-dicyclohexylcarbodiimide (DCC) was added to the mixture. The reaction mixture


2-27

was stirred overnight at room temperature and protected from light, the resulting yellow

solution was filtered to remove the insoluble urea by-product. The organic phase was

removed in vacuo to afford the crude NHS-anthraquinone 12 in quantitative yields that

was used in the next step without further purification.

Scheme 2.14. Synthesis of anthraquinone-NHS. (a) N-hydroxysuccinimide, N,N’-dicyclohexylcarbodiimide, CH2Cl2, r.t.

Azide 13 was prepared using a similar method developed by Zhang et al.40 (Scheme

2.15). Crude anthraquinone-NHS 12 was suspended in a mixed acetone/ethanol solvent.

Azide spacer 11 in an aqueous solution of sodium bicarbonate was added to the reaction

mixture and allowed to react for 23 h. The organic phase was removed in vacuo and

product was resuspended in dichloromethane. Product was washed with basic followed

by acidic water and purified on silica using dichloromethane to afford the yellow

product in 35% yield. CAUTION: Small organic azides can be potentially explosive

and handling of the spacer 11 should be treated with care. 1H NMR spectroscopy

confirmed formation of azide 13, as an upfield shift from 2.27 to 1.97 ppm of proton

signals adjacent to the azide group was observed in comparison to bromide 14 as shown

in Figure 2.7.

An alternative approach to prepare 13 was attempted using an indirect route

which is potentially safer due to the formation of a higher molecular weight azide in

comparison to azide 11. Azide 13 was attempted by initially preparing the bromide

precursor 14, followed by azide substitution under mild heating conditions at 60 oC for

3 days. However, it was found that 50% reduction of the anthraquinone moiety was


2-28

observed compared to bromide 14, as well as formation of unidentifiable by-products by

1H NMR spectroscopy and TLC analysis. Oxidation by

2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) of the reduced species was

attempted with no success.

Scheme 2.15. Synthesis of anthraquinone-2-azidopropane using the direct/indirect methods. (a) Anthraquinone-NHS 12, acetone/EtOH/NaHCO3 (aq), 35%. (b)Anthraquinone-NHS 12, acetone/EtOH/NaHCO3 (aq), 56%. (c) NaN3, CH3CN, 60 oC.

Figure 2.7. 1H NMR (300 MHz, CDCl3) spectra of bromide 14 (top-blue) and azide 13(bottom-black) showing upfield shift of protons after sodium azide displacement reaction indicated by the asterisk.


2-29

2.4.3 Attempted synthesis of anthraquinone-2-propylamido-triazole-maleimide

15

Initially, the synthesis of maleimide modified anthraquinone was attempted using

azide/alkyne click chemistry to allow direct modification of the single cysteine residue

of Acropora millepora green fluorescent protein. Attempted maleimide anthraquinone

15 was prepared by reacting N-propargyl maleimide with azide 13 in a

tetrahydrofuran:water (5:2, v/v) solution containing copper(II) sulphate pentahydrate

and L-ascorbic acid (Scheme 2.16). The solution was reacted overnight and the organic

phase was removed in vacuo and resuspended in dichloromethane, washed with water,

dried with anhydrous sodium sulphate and isolated in vacuo to yield a yellow oil.

Based on 1H NMR spectroscopy, it was observed that the formation of the

triazole was successful with resonances at 8.05 ppm consistent with triazole formation.

However, characteristic maleimide resonance at ca. 6.8 ppm or corresponding

hydrolysed maleimide at ca. 6.3 ppm was not observed.28 Furthermore, mass

spectrometry analysis shows a signal at 596.10 m/z compared to the expected signal for

protonated molecular ion of maleimide 15 at 470.14 m/z. The resulting compound was

not identified; however, it is proposed to be a product of polymerisation.

Scheme 2.16. Attempted synthesis of anthraquinone-triazole-maleimide. (a) CuSO4,L-ascorbic acid, tetrahydrofuran:water (5:2, v/v).


2-30

2.5 Conclusions

Terpyridine ligands bearing amines with 4’-aryl functionalisation were synthesised

based on a modified Kröhnke synthesis with isolation of the azachalcone/pyridinium

iodide precursors. Light-activated chromophores of Ru(II)-bisterpyridine complexes

were prepared based on conditions adapted from Ir(III)-bisterpyridine literature in

ethylene glycol. Homo and heteroleptic aniline complexes 7 and 9 were synthesised to

allow chemical modification by introducing maleimide functionality using peptide

coupling chemistry. Asymmetric and symmetric maleimide complexes 8 and 10 were

synthesised such that these electron donating photosensitisers could be attached to

cysteine bearing proteins/enzymes which are discussed further in Chapter 3. As a

reference compound, complex 6 was prepared for control room temperature

photoreduction studies. Single crystals of complexes 7 and 8 were isolated and analysed

by X-ray crystallography to confirm their structural properties. At the time of writing,

complex 8 is the first reported single crystal X-ray structure of a Ru(II)-bistepyridine

complex with maleimide functionality.

Anthraquinone derivatives were also prepared to function as electron acceptors

after functionalising to a light-activated biological electron donor such as green

fluorescent protein (GFP). Azide bearing anthraquinone 13 was prepared for potential

Cu(I)-catalysed [3+2] Huisgen cycloaddition reactions (click reactions) of azides with

terminal alkynes. Attempts to functionalise azide 13 with maleimides using click

chemistry were unsuccessful and subsequent click chemistry studies of azide 13 with

alkyne-tagged GFP will be discussed further in Chapter 4.


2-31

2.6 References

(1) (a) Heller, M.; Schubert, U. S. Eur. J. Org. Chem. 2003, 947. (b) Hofmeier, H.; Schubert, U. S. Chem. Soc. Rev. 2004, 33, 373.

(2) Anderson, G. W.; Zimmerman, J. E.; Callahan, F. M. J. Am. Chem. Soc. 1964,86, 1839.

(3) (a) Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. Angew. Chem. Int. Ed. 2002, 41, 2596. (b) Dirks, A. J.; van Berkel, S. S.; Hatzakis, N. S.; Opsteen, J. A.; van Delft, F. L.; Cornelissen, J. J. L. M.; Rowan, A. E.; van Hest, J. C. M.; Rutjes, F. P. J. T.; Nolte, R. J. M. Chem. Commun. 2005, 4172.

(4) Morgan, G. T.; Burstall, F. H. J. Chem. Soc. 1932, 20.(5) Blau, F. Ber. Dtsch Chem. Ges. 1888, 21, 1077.(6) Schubert, U. S.; Hofmeier, H.; Newkome, G. R. Modern Terpyridine Chemistry;

1st ed.; WILEY-VCH Verlag: Wienheim, Germany, 2006.(7) Kröhnke, F. Synthesis 1976, 1976, 1.(8) Potts, K. T.; Usifer, D. A.; Guadalupe, A.; Abruna, H. D. J. Am. Chem. Soc.

1987, 109, 3961.(9) Jameson, D. L.; Guise, L. E. Tetrahedron Lett. 1991, 32, 1999.(10) Constable, E. C.; Lewis, J. Polyhedron 1982, 1, 303.(11) Cargill Thompson, A. M. W. Coord. Chem. Rev. 1997, 160, 1.(12) (a) Uenishi, J.; Tanaka, T.; Wakabayashi, S.; Oae, S.; Tsukube, H. Tetrahedron

Lett. 1990, 31, 4625. (b) Parks, J. E.; Wagner, B. E.; Holm, R. H. J. Organomet. Chem. 1973, 56, 53.

(13) Miyaura, N.; Suzuki, A. Chem. Rev. 1995, 95, 2457.(14) Negishi, E. Current Trends in Organic Synthesis; 1st ed.; Pergamon, Oxford,

1983.(15) Stille, J. K. Angew. Chem. Int. Ed. 1986, 25, 508.(16) Fallahpour, R.-A. Synthesis 2003, 155.(17) Sauvage, J. P.; Collin, J. P.; Chambron, J. C.; Guillerez, S.; Coudret, C.; Balzani,

V.; Barigelletti, F.; De Cola, L.; Flamigni, L. Chem. Rev. 1994, 94, 993.(18) Peterson, J. R. PhD Thesis, The University of Sydney, 2009.(19) Mukkala, V.-M.; Helenius, M.; Hemmilä, I.; Kankare, J.; Takalo, H. Helv.

Chim. Acta 1993, 76, 1361.(20) Mikel, C.; Potvin, P. G. Polyhedron 2002, 21, 49.(21) Lainé, P.; Bedioui, F.; Ochsenbein, P.; Marvaud, V.; Bonin, M.; Amouyal, E. J.

Am. Chem. Soc. 2002, 124, 1364.(22) Hofmeier, H.; El-ghayoury, A.; Schenning, A. P. H. J.; Schubert, U. S.

Tetrahedron 2004, 60, 6121.(23) Lashgari, K.; Kritikos, M.; Norrestam, R.; Norrby, T. Acta Cryst. C 1999, 55,

64.(24) Goldstein, D. C.; Cheng, Y. Y.; Schmidt, T. W.; Bhadbhade, M.; Thordarson, P.

Dalton Trans. 2011, 40, 2053.(25) Beley, M.; Collin, J. P.; Louis, R.; Metz, B.; Sauvage, J. P. J. Am. Chem. Soc.

1991, 113, 8521.(26) Storrier, G. D.; Colbran, S. B.; Craig, D. C. J. Chem. Soc., Dalton Trans. 1998,

1351.(27) Ng, W. Y.; Gong, X.; Chan, W. K. Chem. Mater. 1999, 11, 1165.(28) Goldstein, D. C. PhD Thesis, The University of New South Wales, 2011.(29) Hofmeier, H.; Andres, P. R.; Hoogenboom, R.; Herdtweck, E.; Schubert, U. S.

Aust. J. Chem. 2004, 57, 419.


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(30) Maestri, M.; Armaroli, N.; Balzani, V.; Constable, E. C.; Thompson, A. M. W. C. Inorg. Chem. 1995, 34, 2759.

(31) Hovinen, J. Bioconjug. Chem. 2007, 18, 597.(32) Beley, M.; Collin, J.-P.; Sauvage, J.-P.; Sugihara, H.; Heisel, F.; Miehe, A. J.

Chem. Soc., Dalton Trans. 1991, 3157.(33) Phifer, C. C.; McMillin, D. R. Inorg. Chem. 1986, 25, 1329.(34) Constable, E. C.; Lewis, J.; Liptrot, M. C.; Raithby, P. R. Inorg. Chim. Acta

1990, 178, 47.(35) (a) Purves, W. K.; Sadava, D.; Orians, G. H.; Heller, C. H. Life: The Science of

Biology; 7th ed.; Sinauer Associates and W. H. Freeman: U.S.A., 2003. (b) Gust, D.; Moore, T. A.; Moore, A. L. Acc. Chem. Res. 1993, 26, 198.

(36) Luo, W.; Chan, E. W. L.; Yousaf, M. N. J. Am. Chem. Soc. 2010, 132, 2614.(37) (a) Wolfenden, R.; Liang, Y. L.; Matthews, M.; Williams, R. J. Am. Chem. Soc.

1987, 109, 463. (b) Holmes, T. J.; John, V.; Vennerstrom, J.; Choi, K. E. J. Org. Chem. 1984, 49, 4736.

(38) Hvasanov, D.; Goldstein, D. C.; Thordarson, P. In Molecular Solar Fuels; The Royal Society of Chemistry, 2012; p 426.

(39) Lewis, W. G.; Magallon, F. G.; Fokin, V. V.; Finn, M. G. J. Am. Chem. Soc.2004, 126, 9152.

(40) Zhang, Y.; Gu, H.; Yang, Z.; Xu, B. J. Am. Chem. Soc. 2003, 125, 13680.

Chapter 3

Bioconjugate Synthesis of Cytochrome c and RelatedDerivatives

Chapter 3 Bioconjugate Synthesis of Cytochrome c and Related Derivatives

3-2

3 Bioconjugate Synthesis of Cytochrome c and Related Derivatives

In this Chapter, the synthesis of a number of iso-1 cytochrome c based bioconjugates

derived from the Saccharomyces cerevisiae (yeast) species for use as either

light-activated bioconjugates or dimeric bioconjugates were prepared. The

bioconjugation reactions described herein are based on the Michael addition reaction

between a single free cysteine of a protein and a maleimide ligand.

An asymmetric Ru(II)-bisterpyridine cytochrome c (8-cyt c) bioconjugate was

prepared as a light-harvesting bioconjugate for use as an electron transfer

photosensitiser. Additionally, to explore the factors affecting dimer formation of

bioconjugates using bismaleimide functionalised spacers, dimers were prepared using

combinations of free single cysteine containing proteins including iso-1 cytochrome c

(cyt c), bovine serum albumin (BSA) and mutated Acropora millepora green

fluorescent protein (GFP).

These bioconjugates were purified using fast protein liquid chromatography

(FPLC) and were characterised by UV-Vis spectroscopy, mass spectrometry and gel

electrophoresis. The procedures for bioconjugate synthesis are briefly reviewed in this

chapter.

3.1 Purification of Cytochrome c

Isolation of yeast cytochrome c was first achieved in 1930 by Keilin1 using only

precipitation techniques involving sulphur dioxide and salts, with residual

non-cytochrome proteins present in the final product. Since the initial report of

purification of cytochrome c, modifications to the methodology including fractionation

with ammonium sulphate, boiling in ammonium sulphate solution or boiling in the

presence of chloroform have been made.2 With the advent of ion-exchange


3-3

chromatography3, isolation of the variants of yeast cytochrome c including iso-1 and

iso-2 forms has been achieved.4 For this project, bioconjugation reactions involving the

iso-1 form as shown in Figure 3.1 are desirable due to the presence of a single available

cysteine residue at CYS102 for modification with maleimides allowing site-specific

modification.

Figure 3.1. Solid ribbon representation of iso-1 cytochrome c with heme group and CYS102 residue indicated as stick representations. The structure was derived from the protein data bank file ‘1YCC’.5

3.1.1 Purification of iso-1 cytochrome c using cation exchange chromatography

The purification of crude yeast cytochrome c (Sigma Aldrich) has been optimised by

Peterson et al.4 in the Thordarson group. Reduced protein (2 mg, reduced with

dithiothreitol) is loaded onto a strong cation exchange column (Supelco,

7.5 cmh × 0.75 cmd, 3.3 mL column volume) and eluted with a 328 to 450 mM sodium

chloride gradient over 14.5 mL in a 20 mM sodium dihydrogen phosphate buffer, pH 7.0

at 1 mL/min as shown in Figure 3.2. Pure iso-1 cytochrome c was collected as the main


3-4

peak, approximately eluting from 15.8 to 18 mL with slight variation across different

batches. The pooled fraction is concentrated and dialysed against water prior to storage

at -20 oC. The initial reduction step prior to column loading improves separation from

iso-2 cytochrome c while also generating the free cysteine residue (CYS102) by

disulfide bridge cleavage allowing for maleimide-cysteine chemical modification.

Figure 3.2. Purification of iso-1 cytochrome c (peak A) on a Tosoh SP-5PW strong cation exchange column (Supelco, 7.5 cmh × 0.75 cmd, 3.3 mL column volume, 10 mparticle size) eluted with a gradient of 328 to 450 mM in 20 mM sodium dihydrogen phosphate buffer, pH 7.0 at 1 mL/min. Peak B corresponds to iso-2 cytochrome c. The gradient (red) is shown for illustration.

A batch purification run (12 mg crude dry weight) using the method shown in Figure

3.2 typically yields 54% (506 nmol) of pure iso-1 cytochrome c based on UV-Vis

absorbance. Throughout this chapter and project, yield is based on UV-Vis absorbance

using the molar absorptivity of iso-1 cytochrome c ( 410 =97.6 mM-1cm-1)4 and the yield

is interchangeably expressed as both dry weight and the moles of protein.


3-5

3.2 Bioconjugation Methods

Cytochrome c is one of the most highly studied proteins due to its ease of purification

from multiple species3, use in electron transfer studies as a model redox protein6 and the

important roles they play in the electron transport chain of photosynthesis and

respiration.7 Saccharomyces cerevisiae cytochrome c was the first to be isolated from

the cytochrome c enzyme family by Keilin in 1930.1 Bioconjugation of cytochrome c

has been reported extensively in the literature targeting histidine8, lysine9 and cysteine10

residues and a brief review of these types of reactions will be discussed.

3.2.1 Modification of histidine

Modification of histidine residues of horse heart cytochrome c was achieved by the

formation of coordination complexes via coordinate covalent bonding. This is due to the

lone pair electrons of the histidine nitrogen which can coordinate with transition metal

ions. Winkler et al.8a synthesised a Ru(NH3)5(His-33)3+-ferricytochrome c bioconjugate

to study intramolecular electron-transfer kinetics between the two redox centres by

reacting horse heart cytochrome c (0.2 mM) with fifty-fold excess of

[Ru(NH3)5(H2O)]2+ under argon for 72 h.

3.2.2 Modification of lysine

The most common method for chemical modification in bioconjugate chemistry is the

covalent linkage between a ligand and the amines of a protein. Coupling to the -amino

group on a lysine residue is often targeted as they are abundant on the surface of

proteins as well as the -N terminus of a protein.11 They can be easily modified by

reacting with an N-hydroxysuccimide (NHS) activated ester.12 It should be noted,


3-6

modification of amines offer poor site-specificity as the natural occurrence of lysine

residues in mammalian proteins is estimated to be approximately 6%.13

Pan et al.9a synthesised ten singly labelled horse heart cytochrome c

bioconjugates by reacting with an NHS-activated bipyridine ligand, as well as other

products including unmodified protein as shown in Scheme 3.1.

Scheme 3.1. Reactions of amines with N-hydroxysuccimide activated ester.9a

3.2.3 Modification of cysteine

Another popular target for chemical modification of natural amino acids is cysteine.

Based on the theory of nucleophilicity proposed by Edwards and Pearson14, the thiol

group of cysteine is one of the strongest nucleophiles in proteins. Cysteine residues

possess thiol groups with a pKa of approximately 9.15 Due to these properties, specific

chemical modifications of cysteines can proceed selectively and rapidly under benign

conditions.12a They can be used to chemically modify cysteines as maleimide

functionalised ligands are Michael acceptors.16 In the Thordarson group, Peterson et

al.17 demonstrated site-specific modification of iso-1 cytochrome c from the CYS102

residue and maleimide-functionalised ruthenium-terpyridine complexes to form light-

harvesting donor-acceptor bioconjugates as shown in Scheme 3.2. In this project,

maleimide-cysteine Michael addition reactions with free single cysteine proteins will be

the main focus as it allows site-specific modification under benign conditions

(pH 6.8-7).


3-7

Scheme 3.2. Reactions of cysteine with maleimides.17

3.3 Bioconjugation of Ru(II)-cyt c (8-cyt c)

A light-activated bioconjugate, 8-cyt c was synthesised for electron transfer studies

within a membrane environment and as a component in a photosynthetic-respiratory

system for photoreduction of oxygen as discussed in Chapter 6 and 7, respectively. The

synthesis of bioconjugates followed previously optimised conditions in the Thordarson

group.17b The bioconjugate 8-cyt c was site-specifically reacted by using a single

maleimide group functionalised Ru(II)-bisterpyridine chromophore 8 previously

discussed in Chapter 2.

Iso-1 cytochrome c was added to a solution containing five-fold excess of

complex 8 in a phosphate buffer at pH 7.0 containing ethylenediaminetetraacetic acid

(EDTA) and acetonitrile. The final reaction conditions were iso-1 cytochrome c

(10 M), complex 8 (50 M), phosphate buffer (20 mM), EDTA (20 mM) and acetonitrile

(5% v/v) at pH 7.0 as shown in Scheme 3.3. The mixture was stirred overnight at room

temperature in a plastic reaction vessel in the dark to prevent degradation of maleimide


3-8

and photoreduction of cytochrome c. It should be noted that the addition of EDTA

improves bioconjugation yields by removal of trace copper ions via chelation, as

oxidation of cysteine can occur.1

Scheme 3.3. Synthesis of bioconjugate 8-cyt c. (a) Reduced iso-1 cytochrome c (10 M), phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0, acetonitrile (5% v/v), r.t., 6%.

It was observed that with the hexafluorophosphate salt of complex 8, significant

precipitation of the complex was observed after stirring for 20 h resulting in extremely

low yields <1%. However, it was noticed that after exchange with a chloride salt,

improved water solubility of the complex 8 was observed with slight formation of

precipitates. Any precipitates that formed during the course of the reaction were

removed by syringe filtration using a 0.2 m filter and the solution was concentrated

and dialysed into water. Dialysis was performed to remove acetonitrile and EDTA from

the protein solution. In particular, removal of EDTA is essential as leaching of


3-9

immobilised nickel (Ni2+) can occur from subsequent purification using immobilised

metal affinity chromatography (IMAC).

Purification of conjugate 8-cyt c was achieved using similar conditions as

previously developed in the Thordarson group by Peterson et al.17a for

Ru(II)-bisterpyridine based bioconjugates. Separation of covalently attached

Ru(II)-bisterpyridine bioconjugates from unreacted proteins and ligands can be achieved

using Ni2+ IMAC chromatography, although the mechanism remains unknown. The

crude product was loaded onto an IMAC (Ni2+) column and purified using an imidazole

gradient from 0 to 125 mM over 11.5 mL in 20 mM sodium dihydrogen phosphate, 0.5 M

sodium chloride at pH 7.0 as shown in Figure 3.3.

Figure 3.3. Purification of bioconjugate 8-cyt c by IMAC (Ni2+) chromatography (HisTrapTM HP, GE Healthcare) using a gradient from 0 to 125 mM imidazole in 20 mMphosphate buffer, 0.5 M sodium chloride, pH 7.0 in 11.5 mL at 0.5 mL/min. Peak A and B – bioconjugate 8-cyt c. The gradient (green) is shown for illustration.


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Prior to the imidazole gradient, elution of unreacted iso-1 cytochrome c and ligand

occurs. Product containing fractions A and B is eluted after the imidazole gradient,

pooled, concentrated and dialysed against water. This afforded the bioconjugate 8-cyt c

with high purity and moderate yield of 6%. UV-Vis spectroscopy is a key method for

determination of protein and bioconjugate mass and yields, but also to monitor protein

stability18 and to determine the effect and extent of bioconjugation.9a,19 UV-Vis

spectroscopy confirmed the purified product as shown in Figure 3.4.

Figure 3.4. UV-Vis spectra of 8-cyt c (green, H2O) and approximated by the linear sum (blue dashed) of oxidised iso-1 cytochrome c (red, H2O) and complex 8 (black, CH3CN).

The UV-Vis absorption spectrum of the Ru(II) bioconjugate 8-cyt c shows the

characteristic absorption peaks of both iso-1 cytochrome c (410 nm) and complex 8

(485 nm). Based on the spectra shown in Figure 3.4 and the molar absorptivities of iso-1

cytochrome c ( 410 = 97.6 mM-1cm-1)4 and complex 8 ( 485 = 26.0 mM-1cm-1), a 1:1: ratio


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of iso-1 cytochrome c and complex is observed. Figure 3.4 shows that bioconjugate

8-cyt c can be approximated as the linear sum of oxidised iso-1 cytochrome c and

[Ru(tpy)(tpymal)](PF6)2 8, indicating that there is negligible ground state

communication between the two species of the bioconjugate.

Bioconjugate 8-cyt c was characterised by MALDI-TOF mass spectrometry

using either a matrix of saturated sinapinic acid/ -cyano-4-hydroxycinnamic acid

mixture or caffeic acid as it is a soft ionisation technique which produces predominantly

singly charged molecular ions with minimal fragmentation. It was found that for

optimal signal-to-noise, high protein and bioconjugate concentrations (>10 M) were

required and caffeic acid produced better spectra. The MALDI-TOF mass spectrum of

8-cyt c is shown in Figure 3.5.

Figure 3.5. MALDI-TOF mass spectra of iso-1 cytochrome c (black) and 8-cyt c (red). Peaks corresponding to the calculated masses 12 706 and 13 559 Da, respectively, were detected. Spectra were baseline corrected.


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The spectra shown in Figure 3.5 shows that the measured mass of bioconjugate 8-cyt c

(13 555 Da) is in good agreement with the expected value (13 559 Da) and correlates

with UV-Vis analysis showing that only a single chromophore is attached to the protein,

which is expected based on maleimide-cysteine coupling at neutral pH. From the

spectra of iso-1 cytochrome c, matrix adducts of caffeic acid can be observed which is

consistent with previous reports by Yang et al.20 which reported observation of adducts

with matrix components via MALDI-TOF.

Finally, in order to confirm the purity of the bioconjugate 8-cyt c, analysis by

gel electrophoresis was performed and stained with SimplyBlueTM Safestain as shown

in Figure 3.6. Gel electrophoresis of conjugate 8-cyt c was prepared by denaturation at

70 oC for 10 min and if samples were to be reduced, dithiothreitol (DTT) was added.

Figure 3.6. Gel electrophoresis of SeeBlue® Plus2 molecular weight marker, iso-1cytochrome c (expected 12 706), 8-cyt c (expected 13 559) and a mixture of iso-1cytochrome c and myoglobin (Mb, expected 16 948). Samples on the left are reduced with dithiothreitol, while samples on the right are non-reduced. Samples stained with SimplyBlueTM Safestain.


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As expected, bioconjugate 8-cyt c as shown in Figure 3.6 migrates as a slightly higher

molecular weight species than unmodified iso-1 cytochrome c. It can be seen that in the

non-reduced samples, a cytochrome c disulfide dimer band can be observed which is

absent after reduction with DTT. However, the bioconjugate is unaffected indicating

that the covalent linkage between maleimide and cysteine cannot be reduced and is

stable under strong reducing conditions.

3.4 Synthesis of Dimeric Bioconjugates

Dimeric bioconjugates were prepared using Ru(II)-bismaleimide complex 10 and 4,4’-

bipyridinium-N,N-di(maleimidopropyl) hexafluorophosphate 16 as small organic

spacers. In order to investigate factors influencing protein dimer formation to improve

yields, protein dimers were prepared using combinations of iso-1 cytochrome c, bovine

serum albumin (BSA, EC 232-936-2) and green fluorescent protein (Acropora

millepora, GFP) possessing varying molecular size, geometry and in particular

isoelectric points (pI). However, the concept of utilising supramolecular ionic

interactions to facilitate synthetic protein dimer formation has been neglected.

Control of protein dimerisation using small molecular ligands allows for

applications in gene expression21, signal transduction22, protein therapeutics23 and tumor

therapy.24 Although dimerisation of proteins has wide applicability, synthetic

dimerisation by covalent modification using small ligands have mostly been limited to

low molecular weight enzymes or peptides (<10 kDa) which primarily target terminal

(N/C) residues in the literature.25 There have been limited examples of isolated high

molecular weight protein synthetic dimers in reasonable yields (>20%); a 52 kDa dimer

of a monoclonal antibody single-chain fragment (di-scFv)26, a 128 kDa haemoglobin

dimer (di-Hb)27, a 34 kDa human interleukin 1 receptor antagonist (di-IL-1ra)28, a


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133 kDa human serum albumin (HAS) dimer29 and a 96 kDa lipase-BSA heterodimer.30

It is worth noting that Hb has a pI around 7.1-7.531, close to neutrality, largely

eliminating any unfavourable ionic interactions between the two proteins. For di-scFv

and di-IL-1ra, the reaction was facilitated by targeting the corresponding protein

terminal residues. The di-IL-1ra was synthesised by native chemical ligation (NCL) of

N-terminal modified IL-1ra while di-scFV was synthesied from scFV modified on the

C-terminus by a cysteine—E-TAG sequence.32 Additionally, the HSA dimer yields

were reported based on the non-isolated product resulting in an overestimate. Finally,

for the lipase-BSA heterodimer, the reaction was achieved by an azide-alkyne click

reaction.

3.4.1 Synthesis of cyt c-10-BSA

The cyt c-10-BSA (oppositely charged heterodimer) was prepared by adding a solution

of bismaleimide complex 10 (as chloride salt) in acetonitrile to ten-fold excess of iso-1

cytochrome c in a final phosphate buffer at pH 7.0 containing EDTA and acetonitrile

(5% v/v) for 2 h as shown in Scheme 3.4. This step ensures formation of

mono-functionalised cyt c-10 conjugate due to the CYS102 residue being buried in the

hydrophobic pocket of the protein, resulting in slower reaction rates ( 1 h for

completion).17b Desalting of the intermediate conjugate did not require desalting to

remove excess ligand 10 due to low yields from unfavourable homodimer formation,

further discussed in Chapter 3.4.5. Subsequently, BSA was added to the stirring solution

and allowed to react for an additional 21 h in the dark at room temperature. BSA

possesses a more reactive cysteine residue (single cysteine residue, CYS34) ( 2 min for

completion)17b as it is exposed in the hydrophilic region of the protein. The final

reaction conditions were iso-1 cytochrome c (100 M), BSA (200 M), complex 10


3-15

(10 M) in acetonitrile (5% v/v) and 20 mM sodium dihydrogen phosphate, 20 mM

ethylenediaminetetraacetic acid, pH 7.0. It should be noted that a twenty-fold excess of

BSA was added as it has been reported that the actual free cysteine available for

functionalisation is 0.5 mol of the protein.33

Scheme 3.4. Synthesis of bioconjugate cyt c-10-BSA. (a) i) Reduced iso-1 cytochrome c(100 M), 2 h, ii) Bovine serum albumin (BSA, 200 M), 21 h. Phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0, acetonitrile (5% v/v), r.t., 30%. BSA protein structure aligned onto the human serum albumin X-ray crystallographic structure (PDB code: 1AO6).


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Purification of bioconjugate cyt c-10-BSA was achieved by initial dialysis into water to

remove ethylenediaminetetraacetic acid followed by separation of unreacted complex

10 and unreacted proteins using Ni2+ IMAC chromatography. The crude product was

loaded onto an IMAC (Ni2+) column and purified using an imidazole gradient from 0 to

250 mM over 9 mL in 20 mM sodium dihydrogen phosphate, 0.5 M sodium chloride at

pH 7.0 as shown in Figure 3.7a. The product containing fractions (peaks A and B) were

pooled, concentrated and dialysed against water. Unreacted iso-1 cytochrome c, BSA

and complex 10 were eluted prior to the imidazole gradient. Excess unreacted BSA was

present with the cyt c-10-BSA fraction as it is commonly used for enzyme stabilisation

due to its adhesive properties34 and its difficulty of purification has been reported.35 The

product was further purified using a strong cation exchange column using a sodium

chloride gradient from 320 to 450 mM over 14.4 mL in 20 mM sodium dihydrogen

phosphate, pH 7.0 as shown in Figure 3.7b. Excess BSA and mono-functionalised

conjugates (10-cyt c) were removed prior to the sodium chloride gradient and further

residual BSA was removed at a concentration of 1 M sodium chloride (peak E).

Figure 3.7. Purification of bioconjugate cyt c-10-BSA by fast protein liquid chromatography. (a) IMAC (Ni2+) (HisTrapTM HP, GE Healthcare) using a gradientfrom 0 to 250 mM imidazole in 20 mM phosphate buffer, 0.5 M sodium chloride, pH 7.0in 9 mL at 0.5mL/min. Peak A and B – cyt c-10-BSA. (b) Strong cation exchange (CEX) column (SP-5PW, Supelco) using a gradient from 320 to 450 mM sodium chloride in 20 mM phosphate buffer, pH 7.0 in 14.4 mL at 1 mL/min. Peak C and D - cyt c-10-BSA and peak E – residual BSA. The gradient (green) is shown for illustration.


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The product containing fractions C and D is eluted after the sodium chloride gradient,

pooled, concentrated and dialysed against water. This afforded the bioconjugate

cyt c-10-BSA in 30% yield, estimated by gel electrophoresis and UV-Vis spectroscopy

(see Appendix B).

The UV-Vis absorption spectrum of the heterodimer cyt c-10-BSA shows three

primary absorption bands at 280 nm, 410 nm and 495 nm corresponding to BSA,

cytochrome c, and ligand 10, respectively, as shown in Figure 3.8a.

Figure 3.8. UV-Vis spectra of dimeric bioconjugates. (a) cyt c-10-BSA. (b) cyt c-10-cyt c. (c) cyt c-16-GFP. (d) cyt c-10-BSA. All measurements were made in H2O.

Confirmation of cyt c-10-BSA dimer was observed using MALDI-TOF mass

spectrometry with a m/z 80 609 as shown in Figure 3.9.


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Figure 3.9. MALDI-TOF mass spectrum of cyt c-10-BSA. Peaks corresponding to the calculated masses 66 776 and 80 618 Da, respectively, were detected. Spectrum wasbaseline corrected and noise reduced.

The resulting heterodimer was additionally characterised by denaturing gel

electrophoresis. For 12% Bis-Tris SDS PAGE gels, quantitative molecular weight

determination is accurate for proteins less than 30 kDa. Therefore, SDS PAGE

analysis in this study was used qualitatively to confirm formation of product by

observing new bands corresponding to dimer. Hence, the SDS PAGE gel

electrophoresis in Figure 3.10 shows a new dimer band, further confirming conjugate

formation.


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Figure 3.10. Gel electrophoresis of SeeBlue® Plus2 molecular weight marker, iso-1cytochrome c (expected 12 706), green fluorescent protein (GFP, expected 26 807), bovine serum albumin (expected 66 776), cyt c-10-BSA (expected 80 618), cyt c-10-cyt c(expected 26 550), cyt c-16-GFP (expected 40 236) and BSA-10-BSA (expected 134 688). Samples are reduced with dithiothreitol. Samples stained with SimplyBlueTM

Safestain and contrast corrected. Dimer bands highlighted in boxes.

3.4.2 Synthesis of cyt c-10-cyt c

In order to determine the effect of homodimer formation on bioconjugate dimer yields,

the homodimer cyt c-10-cyt c was synthesised. The bioconjugate was prepared by

adding a solution of complex 10 (as a chloride salt) in acetonitrile to ten-fold reduced

iso-1 cytochrome c (five-fold protein per maleimide group) in a final phosphate buffer

solution, pH 7.0, containing EDTA and acetonitrile (5% v/v) for 27 h in the dark at

room temperature as shown in Scheme 3.5. The product was concentrated, dialysed

against water and purified by IMAC (Ni2+) chromatography.


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It should be noted that for all the subsequent protein dimerisation reactions,

conditions used were similar to that of cyt c-10-BSA as cytochrome c is the least

reactive protein, as the free cysteine residue (CYS102) is buried in a hydrophobic

pocket.17b Reaction time varies between different bioconjugation dimer reactions as it is

not a critical factor for dimerisation due to hydrolysis of the maleimide group35-36 and

only requires a minimum overnight incubation to reach completion.17b

Scheme 3.5. Synthesis of bioconjugate cyt c-10-cyt c. (a) Reduced iso-1 cytochrome c(100 M), 27 h. Phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0, acetonitrile (5% v/v), r.t., 1%.

The crude product was loaded onto an IMAC (Ni2+) column and purified over a gradient

from 0 to 250 mM imidazole over 9 mL in 20 mM sodium dihydrogen phosphate, 0.5 M


3-21

sodium chloride at pH 7.0 as shown in Figure 3.12. Prior to the imidazole gradient,

elution of unreacted iso-1 cytochrome c and ligand was achieved. Product containing

fraction A was eluted after the imidazole gradient, pooled, concentrated and dialysed

against water. This afforded the bioconjugate cyt c-10-cyt c with moderate purity and

low yield of 1%. Due to the low yield of the homodimer, isolation with high purity is

extremely difficult.

Figure 3.11. Purification of bioconjugate cyt c-10-cyt c. IMAC (Ni2+) chromatography (HisTrapTM HP, GE Healthcare) using a gradient from 0 to 250 mM imidazole in 20 mMphosphate buffer, 0.5 M sodium chloride, pH 7.0 in 9 mL at 0.5 mL/min. Peak A –bioconjugate cyt c-10-cyt c. The gradient (green) is shown for illustration.

UV-Vis spectroscopy shows characteristic absorption bands at 410 nm and 495 nm

corresponding to cyt c and complex 10 as shown in Figure 3.8 and dimer was further

confirmed by SDS PAGE gel electrophoresis as shown in Figure 3.10 showing a new

dimer band. The formation of dimer was additionally confirmed by MALDI-TOF mass

spectrometry as shown in Figure 3.12 with a m/z of 26 551.


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Figure 3.12. MALDI-TOF mass spectrum of cyt c-10-cyt c. Peaks corresponding to the calculated masses 25 412 and 26 550 Da, respectively, were detected. Spectrum was baseline corrected.

3.4.3 Synthesis of cyt c-16-GFP

In order to probe the effect of dimer yields based on a heterodimer of similar positive

charge, a cyt c-16-GFP heterodimer was prepared. The spacer chosen for this

bioconjugate was based on a bipyridinium (viologen) moiety 16 which is more water

soluble in comparison to complex 10. The dimer was prepared by addition of a solution

of 4,4’-bipyridinium-N,N-di(maleimidopropyl) hexafluorophosphate 16 (fifteen-fold) in

acetonitrile to a solution of green fluorescent protein (GFP, amFP497) derived from

Acropora millepora in phosphate/EDTA buffer at pH 7.0 for 3 h in the dark as shown in

Scheme 3.6. The functionalisation of recombinant GFP with maleimide could be

achieved due to the single cysteine residue at CYS119 introduced by mutation of the

genetic sequence.37


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Subsequently, the crude product was desalted to remove excess viologen spacer

as shown in Figure 3.13a. Purified reduced iso-1 cytochrome c (four-fold) was added to

the desalted product fraction and reacted for a further 17 h in phosphate/EDTA buffer at

pH 7.0 in the dark.

Scheme 3.6. Synthesis of bioconjugate cyt c-16-GFP. (a) i) Green fluorescent protein (10 M), 3 h, ii) Cyt c-16 (70 M), reduced iso-1 cytochrome c (290 M), 17 h.Phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0, acetonitrile (5% v/v), r.t., 0.1%. Green fluorescent protein PDB code: 2A4638, aligned with engineered protein sequence for amFP497.

It was proposed that the extremely low yield was a result of CYS102 being buried in the

hydrophobic pocket of cytochrome c.5 Therefore, an alternative approach to synthesise

cyt c-16-GFP was attempted by preparing cyt c-16 prior to addition of green fluorescent

protein containing surface exposed CYS119. However, no improvement in heterodimer

yield was observed. It should be noted that a cytochrome c-GFP heterodimer has been

previously prepared via recombinant methods as a fused product in the mitochondria


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rather than using an organic linker.39 Additionally, bioconjugation variants of GFP for

use as potential donor-acceptor systems are further discussed in Chapter 4.

Recombinant GFP was expressed with a hexahistidine-tag (6×His-tag) allowing

purification using IMAC chromatography.40 The desalted intermediate product was

loaded onto an IMAC column and purified over a gradient from 0 to 250 mM imidazole

in 20 mm sodium dihydrogen phosphate, 0.5 m sodium chloride at pH 7.0. Prior to the

imidazole gradient, elution of unreacted cytochrome c (peak C) occurs. During the

imidazole gradient, further elution of unreacted cytochrome c and unreacted GFP (peak

D and F) was achieved. Product containing fraction E of bioconjugate cyt c-16-GFP is

eluted, pooled, concentrated and dialysed against water. This afforded the bioconjugate

in an extremely low yield of 0.1%. Residual cytochrome c and GFP was present in the

product fraction due to difficulty in purification for low yielding species.

Figure 3.13. Purification of bioconjugate cyt c-16-GFP. (a) Purification of bioconjugate GFP-16 by fast protein liquid chromatography using a desalting column (HiTrapTM Desalting, GE Healthcare) in 20 mM phosphate buffer, pH 7.0 at 0.5 mL/min. Peak A – mixed GFP and GFP-16. Peak B – unreacted 16. (b) Purification of bioconjugate cyt c-16-GFP using an IMAC (Ni2+) column (HisTrapTM HP, GE Healthcare) using a gradient from 0 to 250 mM imidazole in 20 mM phosphate buffer, 0.5 M sodium chloride, pH 7.0 in 6 mL at 0.5mL/min. Peak C – unreacted cyt c, peak D and F - unreacted cyt c and GFP, peak E – bioconjugate cyt c-16-GFP. The gradient (green) is shown for illustration.

UV-Vis spectroscopy shows characteristic absorption bands at 410 nm and 476 nm

corresponding to cytochrome c and GFP as shown in Figure 3.8. The heterodimer was


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also confirmed by reduced SDS PAGE gel electrophoresis showing a new dimer band

indicated in Figure 3.10. Finally, the heterodimer was confirmed by MALDI-TOF mass

spectrometry as shown in Figure 3.14 with a m/z of 40 245.

Figure 3.14. MALDI-TOF mass spectrum of cyt c-16-GFP. Peaks corresponding to calculated mass 40 236 Da was detected. Spectrum was baseline corrected and noise reduced.

3.4.4 Synthesis of BSA-10-BSA

A homodimer of negative charge was prepared by forming a BSA homodimer using

bismaleimide complex 10 as shown in Scheme 3.7. A solution of complex 10 (as

chloride salt) in acetonitrile was added to fourty-fold BSA in a final 20 mM

phosphate/EDTA buffer, acetonitrile (5% v/v) at pH 7.0 for 23 h at room temperature in

the dark. The crude was concentrated, dialysed against water and the crude was purified

by IMAC chromatography.


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Scheme 3.7. Synthesis of bioconjugate BSA-10-BSA. (a) Bovine serum albumin (400 M), 23 h. Phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0, acetonitrile (5% v/v), r.t., 1%.

The crude was loaded onto an IMAC column and purified over a gradient from 0 to

125 mM imidazole over 6 mL in 20 mM sodium dihydrogen phosphate, 0.5 M sodium

chloride at pH 7.0 as shown in Figure 3.15. Elution of unreacted ligand 10 and BSA

occurred prior to the gradient (peak A). Product containing fraction B eluted during the

imidazole gradient and was pooled, concentrated and dialysed against water to afford

the homodimer in 1% yield.


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Figure 3.15. Purification of bioconjugate BSA-10-BSA by IMAC (Ni2+)chromatography (HisTrapTM HP, GE Healthcare) using a gradient from 0 to 125 mMimidazole in 20 mM phosphate buffer, 0.5 M sodium chloride, pH 7.0 in 6 mL at 0.5 mL/min. Peak A – unreacted 10 and BSA, peak B – BSA-10-BSA. The gradient (green) is shown for illustration.

UV-Vis spectroscopy shows characteristics absorption bands at 280 nm and 495 nm due

to BSA and spacer 10, respectively, as shown in Figure 3.8. Additionally, reduced gel

electrophoresis shows a new band corresponding to dimer formation as shown in Figure

3.10. Due to the high molecular weight (>130 kDa), the BSA homodimer could not be

detected by MALDI-TOF mass spectrometry.

3.4.5 Effect of charge on protein dimer yield

The synthesis of bioconjugate dimers discussed above showed that yields may be

dependent of protein charge. The role and importance of protein charge to probe protein

function after amino acid modification has been utilised in the literature, such as the use

of protein charge ladders.41 However, the exploitation of global protein charge to


3-28

facilitate dimer bioconjugate synthesis via supramolecular interactions has been

neglected.

In this study it was found that a heterodimer of complementary charge and high

molecular weight (>80 kDa), cyt c-10-BSA, could be prepared in up to 30% yield

overcoming steric hindrance as shown in Table 3.1. Cytochrome c is a small protein

(12.7 kDa) with positive charge at physiological pH 7.4 as it possesses a high positive

pI of 10.63 and BSA is a negatively charged large protein (66.7 kDa) with a negative pI

of 4.7.34b

In addition, a positively charged homodimer cyt c-10-cyt c was synthesised and

resulted in a low yield of 1%. Similarly, the positively charged heterodimer

cyt c-16-GFP, which used green fluorescent protein as a like charged protein with a

molecular weight of 26.8 kDa and a high positive pI of 8.342 resulted in extremely low

yields of less than 1%. Additionally, the negatively charged homodimer, BSA-10-BSA

also resulted in an extremely low yield of 1%. Based on these like-charged homo and

heterodimers, it appears the low yields are due to unfavourable dimer formation due to

electrostatic repulsion.

Table 3.1. Summary of conjugate dimer pairings.

Entry Conjugate Protein 1 (pI) Protein 2 (pI) Charge Yielda

1 cyt c-10-BSA cyt c (10.6) BSA (4.7) +/- 30%2 cyt c-10-cyt c cyt c (10.6) cyt c (10.6) +/+ 1%3 cyt c-16-GFP cyt c (10.6) GFP (8.3)b +/+ 0.1%4 BSA-10-BSA BSA (4.7) BSA (4.7) -/- 1%

a Estimated by gel electrophoresis and UV-Vis spectroscopy. b pI for GFP estimated by theoretical calculations of peptide sequence.

To exclude the possibility that the higher yielding complementary charged heterodimer

cyt c-10-BSA was not induced by localised charge effects, but rather global charge, the

protein electrostatic surface of cytochrome c, green fluorescent protein and bovine


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serum albumin was modelled by PyMol (version 1.3). The model assumes that the

positively charged cytochrome c and green fluorescent protein possess 2-8 positive

charges at physiological pH 7.4 and negatively charged bovine serum albumin has

approximately 13 negatively charged residues. From Figure 3.16, the solvent accessible

single cysteine residues of the proteins do not appear to be in particularly negatively or

positively charged regions on the calculated electrostatic surface.43

Figure 3.16. The three proteins used in this model and their key properties. For bovine serum albumin (BSA), the sequence of BSA (ExPASy code: P02769) has been aligned onto the X-ray crystallographic structure of human serum albumin (PDB code: 1AO6). Similarly, the modified Acropora millepora green fluorescent protein (GFP) sequence37

has been aligned with another Anemonia majano GFP structure (PDB code: 2A46). The sequence alignment was performed using ClustalW2 (http:www.ebi.ac.uk/Tools/clustalw2/). The structure for iso-1 cytochrome c (cyt c) was used without further modification (PDB code: 1YCC). Negative (red), neutral (white)and positive (blue) electrostatic surface features are presented according to the inserted polarised scale. Images were generated with PyMol (Version 1.3, Schrödinger, LLC) using the APBS plug-in to calculate the electrostatic surface potentical.43 The pI values for BSA34b and cyt c3 were obtained from the literature while for GFP, it has been estimated by theoretical calculations based on its sequence.42 The number of + or –charges refers to the net whole charges at pH =7.4 assuming only Lys (+), Arg(+), Glu (–) and Asn (–) are charged at that pH (the N- and C-protein terminus cancel each other out). The target cysteine residues for bioconjugation are coloured in green and indicated by an arrow.

http://www.ebi.ac.uk/Tools/clustalw2/


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3.5 Conclusion and Future Work

In this Chapter, the successful synthesis of an asymmetric terpyridine cytochrome c

bioconjugate 8-cyt c was prepared for electron transfer studies in membranes and for

use as a component in a polymer membrane reconstituted electron transfer chain as a

photosynthetic-respiratory hybrid which is further discussed in Chapter 6 and 7,

respectively. The purification of ruthenium(II)-bisterpyridine based bioconjugate could

be achieved using immobilised metal affinity chromatography (IMAC, Ni2+) in high

purity with 6% yield based on procedures previously developed in the Thordarson

group by Peterson et al.17a although the exact mechanism remains unknown. The

limitation of low yielding asymmetric bioconjugate 8-cyt c is a result of the poor water

solubility, and this issue could be addressed further in the future by preparing more

water soluble ligands based on polyethylene glycol modified

ruthenium(II)-bisterpyridine complexes.

Several bioconjugate dimers have been prepared based on combinations of

cytochrome c, green fluorescent protein (Acropora millepora) and bovine serum

albumin to probe factors affecting dimerisation yield. It was found that a high molecular

weight heterodimer of complementary charge cyt c-10-BSA (>80 kDa) was able to be

prepared in up to 30% yield. It is noteworthy that like-charged homo and heterodimers

were prepared in extremely low yields of less than 1%. Based on these studies, it is

proposed that global protein charge can be utilised to induce higher dimer yields by

electrostatic supramolecular ionic attraction. Additionally, the absolute purification of

dimers proved to be difficult due to the low yields of the dimers. In order to confirm

that electrostatic attraction can affect bioconjugate yields, further experiments including

pH titrations on dimer formation (pI effect), Curtin-Hammett regimes44 and


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involvement of other proteins (site directed mutagenesis to introduce CYS) should be

conducted in the future.

The conjugates were characterised by MALDI-TOF mass spectrometry, UV-Vis

spectroscopy and gel electrophoresis.

3.6 References

(1) Keilin, D. Proc. R. Soc. Lond. B 1930, 106, 418.(2) Keilin, D.; Hartree, E. F. Proc. R. Soc. Lond. B 1937, 122, 298.(3) Minakami, S. J. Biochem. 1955, 42, 749.(4) Peterson, J. R.; Thordarson, P. Chiang Mai J. Sci. 2009, 26, 236.(5) Louie, G. V.; Brayer, G. D. J. Mol. Biol. 1990, 214, 527.(6) (a) Millett, F.; Durham, B. Methods Enzymol. 2009, 456, 95. (b) Geren, L.;

Durham, B.; Millett, F. Methods Enzymol. 2009, 456, 507.(7) (a) Cordes, M.; Giese, B. Chem. Soc. Rev. 2009, 38, 892. (b) Gray, H. B.;

Winkler, J. R. Annu. Rev. Biochem. 1996, 65, 537.(8) (a) Winkler, J. R.; Nocera, D. G.; Yocom, K. M.; Bordignon, E.; Gray, H. B. J.

Am. Chem. Soc. 1982, 104, 5798. (b) Yang, X.-J.; Drepper, F.; Wu, B.; Sun, W.-H.; Haehnel, W.; Janiak, C. Dalton Trans. 2005, 256.

(9) (a) Pan, L. P.; Durham, B.; Wolinska, J.; Millett, F. Biochemistry 1988, 27,7180. (b) Hahm, S.; Durham, B.; Millett, F. Biochemistry 1992, 31, 3472.

(10) Wang, K.; Mei, H.; Geren, L.; Miller, M. A.; Saunders, A.; Wang, X.; Waldner, J. L.; Pielak, G. J.; Durham, B.; Millett, F. Biochemistry 1996, 35, 15107.

(11) Hvasanov, D.; Goldstein, D. C.; Thordarson, P. In Molecular Solar Fuels; The Royal Society of Chemistry, 2012; p 426.

(12) (a) Thordarson, P.; Droumaguet, B. L.; Velonia, K. Appl. Microbiol. Biotechnol.2006, 73, 243. (b) Veronese, F. M.; Pasut, G. Drug Discov. Today 2005, 10,1451.

(13) Wu, G.; Ott, T. L.; Knabe, D. A.; Bazer, F. W. J. Nutr. 1999, 129, 1031.(14) Edwards, J. O.; Pearson, R. G. J. Am. Chem. Soc. 1962, 84, 16.(15) Hermanson, G. T. Bioconjugate Techniques; 2nd ed.; Elsevier Inc.: San Diego,

CA, 2008.(16) Hodgson, D. R. W.; Sanderson, J. M. Chem. Soc. Rev. 2004, 33, 422.(17) (a) Peterson, J. R.; Smith, T. A.; Thordarson, P. Chem. Commun. 2007, 1899. (b)

Peterson, J. R.; Smith, T. A.; Thordarson, P. Org. Biomol. Chem. 2010, 8, 151.(18) Kaminsky, L. S.; Davison, A. J. Biochemistry 1969, 8, 4631.(19) Geren, L.; Hahm, S.; Durham, B.; Millett, F. Biochemistry 1991, 30, 9450.(20) Yang, H.; Liu, N.; Qiu, X.; Liu, S. J. Am. Soc. Mass Spectrom. 2009, 20, 2284.(21) Carlotti, F.; Zaldumbide, A.; Martin, P.; Boulukos, K. E.; Hoeben, R. C.;

Pognonec, P. Cancer Gene Ther. 2005, 12, 627.(22) Spencer, D. M.; Belshaw, P. J.; Chen, L.; Ho, S. N.; Randazzo, F.; Crabtree, G.

R.; Schreiber, S. L. Curr. Biol. 1996, 6, 839.(23) Press, O. W.; Corcoran, M.; Subbiah, K.; Hamlin, D. K.; Wilbur, D. S.; Johnson,

T.; Theodore, L.; Yau, E.; Mallett, R.; Meyer, D. L.; Axworthy, D. Blood 2001,98, 2535.


3-32

(24) Quintarelli, C.; Vera, J. F.; Savoldo, B.; Giordano Attianese, G. M. P.; Pule, M.; Foster, A. E.; Heslop, H. E.; Rooney, C. M.; Brenner, M. K.; Dotti, G. Blood2007, 110, 2793.

(25) (a) Ziaco, B.; Pensato, S.; D’Andrea, L. D.; Benedetti, E.; Romanelli, A. Org. Lett. 2008, 10, 1955. (b) Xiao, J.; Tolbert, T. J. Org. Lett. 2009, 11, 4144. (c) Eger, S.; Scheffner, M.; Marx, A.; Rubini, M. J. Am. Chem. Soc. 2010, 132,16337. (d) Li, X. Chem. Asian. J. 2011, 6, 2606.

(26) Natarajan, A.; Du, W.; Xiong, C.-Y.; DeNardo, G. L.; DeNardo, S. J.; Gervay-Hague, J. Chem. Commun. 2007, 695.

(27) (a) Foot, J. S.; Lui, F. E.; Kluger, R. Chem. Commun. 2009, 7315. (b) Yang, Y.; Kluger, R. Chem. Commun. 2010, 46, 7557.

(28) Xiao, J.; Hamilton, B. S.; Tolbert, T. J. Bioconjug. Chem. 2010, 21, 1943.(29) Komatsu, T.; Oguro, Y.; Teramura, Y.; Takeoka, S.; Okai, J.; Anraku, M.;

Otagiri, M.; Tsuchida, E. Biochim. Biophys. Acta, Gen. Subj. 2004, 1675, 21.(30) Hatzakis, N. S.; Engelkamp, H.; Velonia, K.; Hofkens, J.; Christianen, P. C. M.;

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(44) Seeman, J. I. Chem. Rev. 1983, 83, 83.

Chapter 4

Green Fluorescent Protein as a Light-Induced Electron Donor

Chapter 4 Green Fluorescent Protein as a Light-Induced Electron Donor

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4 Green Fluorescent Protein as a Light-Induced Electron Donor

In this Chapter, the synthesis of a mutant green fluorescent protein (GFP) bioconjugate

as a potential covalent donor-acceptor system based on the N-hydroxysuccinimide

anthraquinone 12 acceptor was prepared using non-specific lysine modification.

Additionally, green fluorescent protein has been mutated to contain a single cysteine

residue (CYS119)1 using recombinant methods and is derived from the Acropora

millepora species. The synthesis of site-specific functionalised GFP as potential

donor-acceptor systems using a 4,4’-bipyridinium (viologen) derivative 16 was

achieved and a Cu(I)-catalysed [3+2] Huisgen cycloaddition using azide functionalised

anthraquinone 13 attempted.

Light-induced electron transfer studies between GFP (electron donor) and

p-benzoquinone, anthraquinone and viologen electron acceptors as both non-covalent

and bioconjugate mixtures were monitored using steady-state techniques including

UV-Vis and fluorescence spectroscopy and performed using a Xenon arc lamp or

custom built LED array. The fluorescence lifetime of = 1.65±0.06 ns was determined

for GFP. Additionally, quenched lifetimes due to energy/electron transfer of 0.14±0.01

and 0.23±0.04 ns with corresponding electron transfer rates of ket = 6.5±0.5 × 109 and

3.7±0.7 × 109 s-1 were determined for a non-covalent mixture of GFP with

p-benzoquinone or anthraquinone-2-carboxylic acid, respectively. The fluorescence

lifetime for bioconjugate 12-GFP indicated electron transfer with a quenched lifetime of

0.28±0.04 ns (ket = 2.9±0.4 × 109 s-1).

4.1 Background

Over the last two decades, the green fluorescent protein (GFP) has become one of the

most utilised fluorescent probes in cell biology and molecular biology.2 This is due to


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the ability to fuse GFP to proteins of interest3 and express the resulting gene in vivo to

function as a visual indicator using standard fluorescence microscopy techniques. GFP

has been used for protein localisation4 in cells or organisms or in protein-protein

interaction studies5 via fragment reassembly or Förster resonance energy transfer.

The prevalent role of GFP as a fluorescent probe in cell biology eventually led

to the award of the 2008 Nobel Prize in Chemistry shared by Professors Osamu

Shimomura, Martin Chalfie and Roger Y. Tsien. Wild type GFP (wtGFP) from the

jellyfish Aequorea victoria was first isolated by Shimomura et al.6 in 1962. Chalfie and

co-workers4 demonstrated the versatility of GFP as a visual probe in organisms and

Tsien et al.7 expanded their applications by introducing chromophore mutations to

create spectroscopic variants.

GFPs can be found in natural sources of jellyfish (Aequorea victoria), sea corals

and sea anemones.2c GFP is an approximately 230 amino acid residue fluorescent

protein with monomer molecular weight of 26.8 kDa as shown in Figure 4.1.8 The high

quantum yield and unique fluorescent properties of the protein is a result of sequestering

of the chromophore from the bulk solvent in an -helix that runs down the centre of an

11-stranded -barrel and prevents eventual nonradiative cis to trans isomerisation of the

central chromophore.9


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Figure 4.1. Solid ribbon representation of green fluorescent protein (Acropora millepora) with tripeptide chromophore (GLN68-TYR69-GLY70) and CYS119 residue indicated as stick representations. The modified protein sequence was aligned against another Anemonia majano GFP structure from the protein data bank file ‘2A46’.10

The wild type GFP chromophore is formed via an intramolecular autocatalytic

generation from the tripeptide SER65-TYR66-GLY67 as a

p-hydroxybenzylidene-imidazolidinone chromophore11 with coplanar cis

conformation.12 The chromophore in wild type GFP and variants from different species

and mutations can exist in two tautomeric forms as shown in Figure 4.2, including the

neutral and anionic form responsible for absorption centred near 395 and 475 nm,

respectively.13 The tautomeric forms of the chromophore result in the same emission

band due to ultrafast excited state proton transfer from the neutral to anionic form upon

photoexcitation.14


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Figure 4.2. Tautomeric forms of the green fluorescent (GFP) chromophore. Neutral form is the protonated phenolic form of the chromophore with a keto oxygen on the imidazole ring responsible for the absorption band centred near 395 nm. Anionic form is responsible for the absorption band centred near 475 nm. The equilibrium ratio is dependent on electrostatic interactions between the chromophore and surroundingresidues and are dependent on GFP variants and mutations.9a

The biological purpose and function of bioluminescence of GFPs in jellyfish and sea

corals remain unknown. However, GFP may participate in photochemical reactions with

oxidants as reported by Lukyanov and co-workers who have shown that GFPs from

different species can act as light-induced electron donors.15 In this chapter, mutant GFP

based bioconjugates derived from Acropora millepora (amFP497) with viologen or

quinones as potential electron acceptors were prepared with a GLN68-TYR69-GLY70

chromophore to explore potential light-induced electron transfer reactions in GFP.1

4.2 Synthesis of GFP-Acceptor Bioconjugates

Viologen and quinone derivatives are the most widely studied electron acceptors.16 In

this project, covalent modification of GFP viologen 16 and NHS-anthraquinone 12 was

achieved using cysteine-maleimide coupling17 or amine (lysine) modification,

respectively.18


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4.2.1 Attempted synthesis of anthraquinone-triazole-GFP via click chemistry

The synthesis of a 13-triazole-GFP bioconjugate as a light-activated donor-acceptor

system was attempted using Cu(I)-catalysed [3+2] Huisgen cycloaddition between a

terminal alkyne and azide, also known as the click reaction. Since the initial report of

the Cu(I)-catalysed click reaction between alkyne and azides in 2002 by Sharpless et

al.19, this approach has been a popular strategy for bioconjugation reactions.20 The click

reaction offers advantages over other bioconjugation techniques such as high coupling

efficiency and high selectivity which prevents side reactions with other functional

groups in a reaction mixture, even allowing reactions within cells.21 Furthermore, due to

the weak acid-base properties of azides and alkynes, it is suitable for protein

modification due to benign reaction conditions as well as its chemoselectivity.20b

In order to perform click chemistry on GFP, the alkyne precursor bioconjugate

was prepared as shown in Scheme 4.1. This bioconjugation was performed by reacting

alkyne-maleimide 17 with mutant GFP containing a single cysteine residue (CYS119)

as discussed in Chapter 3. This strategy using the same alkyne-maleimide has been

reported previously to modify bovine serum albumin (BSA).22 GFP was added to a

solution containing ten-fold excess of alkyne-maleimide 17 in a phosphate buffer at

pH 7.0 containing ethylenediaminetetraacetic acid (EDTA) and

N,N-dimethylformamide. The final reaction conditions were GFP (10 M),

alkyne-malemide 17 (100 M), phosphate buffer (20 mM), EDTA (20 mM), and

N,N-dimethylformamide (5% v/v) at pH 7.0. The mixture was stirred for 16.5 h at room

temperature in the dark. The crude mixture was concentrated to ca. 1 mL and desalted

to remove excess alkyne-maleimide 17. Throughout this chapter, yield is based on

UV-Vis absorbance using the molar absorptivity of GFP (amFP497,


4-7

476 = 31.42 mM-1cm-1) and the yield is interchangeably expressed as both dry weight

and the moles of protein.

Scheme 4.1. Synthesis of bioconjugate 17-GFP. (a) Green fluorescent protein (10 M), phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0,N,N-dimethylformamide (5% v/v), r.t., 50%.

The characterisation of alkyne-tagged GFP, 17-GFP by MALDI-TOF mass

spectrometry showed complete conversion into the modified product after desalting as

shown in Figure 4.3 with a product signal of m/z 26 967. Due to the small change in

molecular weight, gel electrophoresis did not show any significant change in band

migration. It should be noted that the direct bioconjugation approach could not be used

as the synthesis of anthraquinone-maleimide 15 was unsuccessful which was discussed

previously in Chapter 2.


4-8

Figure 4.3. MALDI-TOF mass spectrum of alkyne-tagged GFP (17-GFP). Peak corresponding to the calculated mass of 26 965 Da was detected. Spectrum was baseline corrected and noise reduced.

The attempted synthesis of anthraquinone functionalised GFP was attempted using

anthraquinone-azide 13 via click chemistry. Anthraquinone was selected due to

improved stabilities compared to p-benzoquinone which can degrade into quinhydrones

when isolated or stored in solution.23 The conditions used were based on conditions

optimised previously in the Thordarson group by Goldstein et al.24 A solution of a

preformed Cu(I) source, tetrakis(acetonitrile)copper(I) hexafluorophosphate and azide

13 in N,N-dimethylformamide was added to alkyne functionalised GFP (17-GFP) in

4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer at pH 7.0 for 19 h

in the dark at 30 oC with a nitrogen overlay. Significant precipitation was observed after

incubation. The filtrate was characterised after syringe filtration using a 0.2 m

membrane, however, UV-Vis spectroscopy and size exclusion chromatography (HiTrap


4-9

DesaltingTM, GE Healthcare) showed no trace of protein, indicating complete

denaturation of protein.

It was proposed that protein denaturation may have been a result of the

formation of a histidine-Cu(II) chelation complex25 (recombinant GFP expressed with

hexahistidine-tag) or Cu(I)-acetylide intermediate.26 In order to address these issues, a

control experiment under identical reaction conditions containing all components was

repeated using unmodified GFP. Precipitation was observed within 30 min of stirring

indicating protein denaturation due to presence of Cu(II) from oxidation. Based on this

control experiment, the primary factor causing denaturation is a result of the

histidine-Cu(II) chelate as GFP has been previously modified using click chemistry with

polymersomes which did not possess a hexahistidine-tag.27

4.2.2 Synthesis of anthraquinone-GFP (12-GFP) via amine modification

The anthraquinone-GFP bioconjugate (12-GFP) was prepared by non-specific lysine

modification using an NHS-functionalised anthraquinone 12 as shown in Scheme 4.2. A

solution of GFP was added to crude anthraquinone 12 in aqueous sodium bicarbonate

and N,N-dimethylformamide at pH 8.3. The final reaction conditions were GFP

(10 M), anthraquinone 12 ( 100 M), aqueous sodium bicarbonate (50 mM),

N,N-dimethylformamide (5% v/v) at pH 8.3. The mixture was reacted for 16.5 h,

concentrated to ca. 1 mL and desalted by size exclusion chromatography affording the

bioconjugate in 27% yield, based on total protein concentration.


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Scheme 4.2. Synthesis of anthraquinone-GFP (12-GFP). (a) Green fluorescent protein (10 M), aqueous sodium bicarbonate (50 mM), pH 8.3, N,N-dimethylformamide (5% v/v), r.t., 27%.

Characterisation of the desalted product by MALDI-TOF mass spectrometry shows a

mixture of unmodified GFP and singly and doubly labelled 12-GFP in an approximate

ratio of 1:1:1 based on peak intensity as shown in Figure 4.4.

Figure 4.4. MALDI-TOF mass spectrum of anthraquinone-GFP (12-GFP). Peaks corresponding to the calculated masses of 26 807, 27 041 and 27 275 Da, respectively, were detected. Spectrum was baseline corrected and noise reduced.


4-11

The UV-Vis absorption spectrum of the anthraquinone bioconjugate 12-GFP shows the

characteristic absorption peaks of both GFP (476 nm) and the anthraquinone moiety 12

(258 nm). Based on the spectra shown in Figure 4.5 and the molar absorptivities of GFP

(amFP497, 476 = 31.42 mM-1cm-1) and anthraquinone 14 ( 258 = 29.62 mM-1cm-1),

deconvolution of the spectra indicated that the desalted product was composed of 25%

bioconjugate 14-GFP relative to unmodified GFP. UV-Vis spectroscopy provides a

more accurate approach to determine composition in contrast to MALDI-TOF mass

spectrometry as the ionisation efficiency of different molecular weight species varies. It

should be noted that anthraquinone-bromide 14 was chosen as the reference compound

as anthraquinone-2-carboxylic acid displays poor solubility in organic solvents and

water.

Figure 4.5. UV-Vis spectra of 14-GFP (green, H2O) and approximated by the linear sum (blue dashed) of green fluorescent protein (red, H2O) and anthraquinone 14(black, CH2Cl2).


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4.2.3 Synthesis of viologen-GFP (16-GFP)

The potential donor-acceptor system based on viologen (16-GFP) was prepared using

cysteine-maleimide chemistry previously discussed in Chapter 3. GFP was added to a

solution containing ten-fold excess of bismaleimide viologen 16 in a phosphate buffer at

pH 7.0 containing EDTA and acetonitrile. The final reaction conditions were GFP

(10 M), viologen 16 (100 M), phosphate buffer (20 mM), EDTA (20 mM) and

acetonitrile (5% v/v) at pH 7.0 as shown in Scheme 4.3. The mixture was stirred

overnight at room temperature in the dark and quenched with 2-mercaptoethanol.

Scheme 4.3. Synthesis of viologen-GFP (16-GFP). (a) Green fluorescent protein (10 M), phosphate buffer (20 mM), ethylenediaminetetraacetic acid (20 mM), pH 7.0,acetonitrile (5% v/v), r.t., 11%.

The crude was concentrated to ca. 1 mL and purified by Ni2+ immobilised metal affinity

chromatography using an imidazole gradient from 0 to 500 mM over 16 mL in 20 mM

sodium dihydrogen phosphate, 0.5 M sodium chloride at pH 7.0 as shown in Figure 4.6.


4-13

Prior to the imidazole gradient, elution of unreacted ligand 16 (peak A) occurred. Both

unreacted GFP (peak B) and product containing fraction (peak C) as a mixture of

unmodified protein and bioconjugate 16-GFP were eluted after the imidazole gradient.

The product containing fraction was concentrated and dialysed against water affording

the product in 11% yield.

Figure 4.6. Purification of bioconjugate 16-GFP by IMAC (Ni2+) chromatography (HisTrapTM HP, GE Healthcare) using a gradient from 0 to 500 mM imidazole in 20 mMphosphate buffer, 0.5 M sodium chloride, pH 7.0 in 16 mL at 0.5 mL/min. Peak A –unreacted ligand 16, peak B – unmodified GFP and peak C – bioconjugate 16-GFP (mixture of unmodified GFP and bioconjugate).

Characterisation of bioconjugate 16-GFP by MALDI-TOF mass spectrometry spiked

with unmodified GFP showed the single site-specific attachment of ligand 16 with a

measured mass of 27 246 Da and is in agreement with the expected value of 27 239 Da

as shown in Figure 4.7.


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Figure 4.7. MALDI-TOF mass spectrum of bioconjugate 16-GFP spiked with unmodified green fluorescent protein. Peaks corresponding to the calculated masses 26 807 and 27 239 Da, respectively, were detected. Spectrum was baseline correctedand noise reduced.

4.3 GFP Donor-Acceptor Studies

Electron transfer reactions of redox proteins including the electron transport chain for

photosynthesis and respiration are critical to the biological function of organisms.

Electron transfer reactions have been primarily focused on redox proteins including

cytochrome c28, however, it has been reported recently that a possible role of GFP is a

potential light-induced electron donor in the presence of suitable acceptors.15

GFP undergoes photoconversion into a red fluorescent state under anaerobic

conditions29 where the oxygen concentration is below 1% without external agents.30 On

the other hand, light-induced red photoconversion of GFP in the presence of electron

acceptors from a green to red fluorescent state under aerobic conditions has been

reported by Bogdanov et. al.15, known as oxidative redding. The proposed red


4-15

photoconversion mechanism is a one-photon and step-wise two-electron oxidation

process as shown in Scheme 4.1, however, the exact nature of the red chromophore is

unknown. In order for oxidative redding to occur, a tyrosine residue is required in the

protein tripeptide chromophore.

Scheme 4.4. Proposed photoinduced aerobic red photoconversion mechanism. An excited green chromophore (Chrg) transfers one electron to an acceptor molecule (A) resulting in a short-lived intermediate (Chr +). If the intermediate can transfer a second electron in its lifetime, a red chromophore (Chrr) is formed; otherwise, a permanent bleached chromophore (Chrbl) is formed.15

Quinone and viologen based electron acceptors are suitable as potential acceptor

molecules of GFP as quinone can be reduced via a two-electron process31 as shown in

Scheme 4.5a and viologen can be reduced to form a mono-reduced species which can be

detected spectroscopically as shown in Scheme 4.5b.32

Scheme 4.5. Reversible conversion of oxidised/reduced states of electron acceptors. (a) p-benzoquinone. (b) viologen.


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4.3.1 Steady-state spectroscopy studies

4.3.1.1 Viologen 16 acceptor studies

To monitor whether GFP (amFP497) could be a potential light-induced electron donor,

initial studies were performed on the viologen-GFP bioconjugate, 16-GFP. Electron

transfer studies were monitored using UV-Vis spectroscopy by following the

characteristic mono-reduced viologen species with a characteristic absorption band at

600 nm.32 Photoexcitation was performed at room temperature on the bioconjugate

16-GFP ( 13 M) in a phosphate buffer, pH 7.0 (20 mM) in the presence of EDTA

(present to stabilise the mono-reduced viologen species).33 Due to the instability of the

mono-reduced viologen species in the presence of oxygen, samples were degassed

(120 mbar) in a specialised small volume cuvette at 0 oC for 30 min and overlayed with

nitrogen. Photoexcitation was performed using a high powered xenon lamp (450 W)

fitted with a 2 mm iris to restrict the amount of excitation light and a UV filter equipped

for 30 min at a distance of 10 cm. The irradiated sample was monitored using UV-Vis

spectroscopy and followed the absorbance from 250 to 800 nm as shown in Figure 4.8a.

The characteristic reduced viologen species was not observed indicating no electron

transfer processes taking place and is consistent with time-resolved fluorescence studies

discussed in the next section (Chapter 4.3.2). In order to eliminate the possibility that

the formation of the mono-reduced species is hindered as a result of bioconjugation, the

degassed bioconjugate 16-GFP was chemically reduced using a strong reductant,

sodium dithionite and the characteristic 600 nm absorption band could be observed as

shown in Figure 4.8b. Similarly, in order to discern whether the lack of electron transfer

was a result of donor-acceptor distance ( 26 Å estimated from chromophore imidazole

ring to central bipyridinium ring via CYS119)10 or viologen is an unsuitable electron

acceptor, a non-covalent mixture of GFP (20 M) and viologen 16 (1 mM) was


4-17

photoexcited using a custom built LED source (3 × 490 nm, 30 W) for 3 h and no shifts

in absorbance or fluorescence emission bands were observed indicating no

photoconversion.

Figure 4.8. Photoexcitation studies of degassed viologen-GFP bioconjugate (16-GFP). (a) Irradiation for 0 min (black) and 30 min (red) with Xenon arc lamp (450 W) showing no change in absorbance spectra corresponding to photoreduction. (b) Chemical reduction of bioconjugate with sodium dithionite (black) showing mono-reduced species with characteristic absorption band at 600 nm; re-oxidised (red) with vigorous shaking showing reversibility of viologen oxidation states.

4.3.1.2 p-benzoquinone acceptor studies

To determine whether the lack of light-induced electron transfer was a result of the GFP

used in this study or viologen is a poor electron acceptor for GFP, p-benzoquinone (BQ)

was employed as an alternative acceptor molecule.16,34 As a result of the lack of

photoinduced electron transfer for viologen-GFP (16-GFP), non-covalent studies using

quinones were performed to determine if a donor-acceptor bioconjugate system is

viable. A non-covalent mixture of GFP (20 M) with fifty-fold excess of

p-benzoquinone (1 mM) in a phosphate buffer, pH 7.0 (20 mM) was irradiated using the

LED source for 3 h at a distance of 2 cm and monitored by UV-Vis and fluorescence

spectroscopy. After irradiation, the appearance of a new absorbance band was observed

at ca. 540 nm as shown in Figure 4.9a. No shift in the fluorescence emission spectra

(ca. 1.6 M) was observed as shown in Figure 4.9b, however, electron/energy transfer


4-18

was confirmed by fluorescence lifetime measurements which is discussed in the next

section.

Figure 4.9. Light-induced photoconversion of a non-covalent mixture containing GFP (20 M) and p-benzoquinone (BQ, 1 mM) in a 20 mM phosphate buffer, pH 7.0. (a) UV-Vis spectra (20 M) showing formation of new absorption band at 540 nm after 3 hirradiation (red) due to electron transfer compared to GFP (black). (b) Fluorescence emission spectra ( 1.6 M, ex = 476 nm) showing no red shift after 3 h irradiation (red) compared to GFP (black).

4.3.1.3 Anthraquinone acceptor studies

Steady state measurements of a non-covalent mixture of p-benzoquinone and GFP

demonstrated light-induced electron transfer showing quinone family acceptors are

viable for covalent donor-acceptor systems. However, due to their inherent long-term

instability23 and lack of functionalisation possibilities, anthraquinone was also screened.

A non-covalent mixture of GFP (20 M) and fifty-fold excess of

anthraquinone-2-carboxylic acid (AQ) in phosphate buffer (20 mM), pH 7.0 was

irradiated using the LED source for 3 h at a distance of 2 cm and photoconversion was

monitored by UV-Vis and fluorescence spectroscopy. After irradiation, bleaching of the

anionic chromophore absorbance band (476 nm) occurred as shown in Figure 4.10a.

Based on Figure 4.10b, analysis of the diluted mixture (ca. 1.6 M) by fluorescence

spectroscopy, a 15 nm stoke shift of the fluorescence emission band from 497 to


4-19

512 nm after irradiation was observed, indicating red photoconversion due to

photo-induced electron transfer.

Figure 4.10. Light-induced photoconversion of a non-covalent mixture containing GFP (20 M) and anthraquinone-2-carboxylic acid (AQ, 1 mM) in a 20 mM phosphate buffer, pH 7.0. (a) UV-Vis spectra (20 M) showing bleaching of the anionic chromophore absorption band at 476 nm after 3 h irradiation (red) due to electron transfer compared to GFP (black). (b) Fluorescence emission spectra ( 1.6 M, ex = 476 nm) showing red shifting of the fluorescence emission band from 497 to 512 nm after 3 h irradiation (red) compared to GFP (black).

It should be noted that the photophysical properties of GFP are different after irradiation

between p-benzoquinone and anthraquinone-2-carboxylic acid, either showing the

appearance of a new absorbance band or red shift of the fluorescence emission band,

respectively. It is proposed that this may be due to the different redox potentials

between the quinone species of p-benzoquinone and anthraquinone-2-carboxylic acid

(estimated using 9,10-anthraquinone) with redox potentials of +0.283 and -0.28 V (vs

SHE), respectively, affecting the chromophore differently.35

It was determined that anthraquinone-2-carboxylic acid is a viable electron

acceptor for light-induced electron transfer with GFP based on non-covalent studies.

Therefore, further photoexcitation experiments were performed on the non-specific

functionalised donor-acceptor bioconjugate, 12-GFP. The bioconjugate 12-GFP (18 M)

in 20 mM phosphate buffer, pH 7.0 was irradiated with the LED source for 3 h at a

distance of 2 cm. Irradiation showed conversion of the chromophore from the anionic to


4-20

neutral chromophore and photobleaching, however, no stoke shift of the fluorescence

emission band was observed indicating red photoconversion of 12-GFP did not occur as

shown in Figure 4.11. The lack of red photoconversion may be due to the

donor-acceptor distance being greater than distance required for electron transfer

(assuming maximum tunnelling distance of 20 Å via superexchange).36 The exact

residues functionalised (21 possible lysines available)1 is unknown due to the

non-specific nature of amine modification. In order to confirm whether energy/electron

transfer processes are taking place, time-resolved fluorescence spectroscopy were

performed on both the non-covalent mixture of GFP and anthraquinone-2-carboxylic

acid as well as the bioconjugate 12-GFP.

Figure 4.11. Photoexcitation of 12-GFP (18 M) in a 20 mM phosphate buffer, pH 7.0.(a) UV-Vis spectra (18 M) showing bleaching of the anionic chromophore absorption band at 476 nm and partial shifting to the neutral state after 3 h irradiation (red) compared to GFP (black). (b) Fluorescence emission spectra ( 1.6 M, ex = 476 nm) showing no shift of the fluorescence emission band (497 nm) after 3 h irradiation (red) compared to GFP (black).

4.3.2 Fluorescence lifetime studies

Fluorescence lifetime studies of GFP and electron transfer measurements were

performed in collaboration with A/Prof. Timothy W. Schmidt, Dr. Raphaël G. C. R.

Clady and Mr. Murad Tayebjee at the University of Sydney, Australia. Time-resolved

fluorescence spectroscopy was performed on GFP, a non-covalent mixture of


4-21

p-benzoquinone/anthraquinone-2-carboxylic acid and GFP, bioconjugates 12-GFP and

16-GFP.

Lifetime measurements of all samples were performed at a concentration of

1.6 M in a 5 mM phosphate buffer, pH 7.0 containing 5 mM

ethylenediaminetetraacetic acid at room temperature. Prior to measurements, samples

were irradiated with the polarised laser source at an excitation wavelength of 476 nm for

10 min to allow stabilisation of the fluorescence emission intensity. Additionally, it

should be noted that fluorescence emission was probed at a wavelength of either 515 or

540 nm as detection at the GFP emission maximum (497 nm) resulted in saturation of

the detector. Fluorescence lifetime measurements were performed in a specialised low

volume quartz cuvette with a 2-3 mm pathlength to minimise background fluorescence

due to delayed fluorescence of sample across the entire pathlength of the cell.

Based on Figure 4.12, it was found that GFP had a fluorescence lifetime of

1.65±0.06 ns. This is not dissimilar to wild type jellyfish Aequorea GFP, with a 2.8 ns

fluorescence lifetime due to photochromicity between the protonated and deprotonated

species.37 The fluorescence data was fitted assuming a mono-exponential decay (single

fluorescent species with no quencher) and estimated using the relative amplitudes:9b

/0)( teFtF (4.1)

where F(t) is the fluorescence intensity, F0 is the initial fluorescence intensity, t is time

and is the fluorescence lifetime. Additionally, fluorescence decay of bioconjugate

16-GFP showed a similar decay profile and modelled according to Equation 4.1. The

fluorescence lifetime of 16-GFP was estimated to be 1.66±0.05 ns, indicating no

electron or energy transfer processes occurring.

In contrast, quenching of the fluorescence decay of the non-covalent mixture of

GFP and p-benzoquinone was observed as shown in Figure 4.12, indicative of


4-22

electron/energy transfer processes.38 The fluorescence showed double exponential decay

behaviour with components having lifetimes of 0.76±0.04 and 0.14±0.01 ns, which

appears to correspond to two different types of static quenching which was modelled

according to Equation 4.2:9b,39

21 /0

/0 ')( tt eFeFtF (4.2)

where F(t) is the fluorescence intensity, F0 and F0’ is the initial fluorescence intensity, t

is time and is the fluorescence lifetime. Similarly, the initial fluorescence intensity

coefficients were found to be 0.47±0.03 and 0.95±0.05 for the two quenching modes,

respectively, indicating approximately 67% of the GFP population with the shorter

lifetime component is likely participating in electron/energy transfer processes.

Assuming that electron transfer takes place, from the lifetime value of the shorter

quenched lifetime, the rate constant of the forward electron transfer (ket) can be

calculated according to Equation 4.3:40

0

11etk (4.3)

where and 0 are the respective lifetimes in the presence and absence of the electron

acceptor. The calculated value was ket = 6.5±0.5 × 109 s-1. The corresponding ket for the

longer quenching lifetime would be 7.1±0.5 × 108 s-1.


4-23

Figure 4.12. Time-resolved fluorescence (ultrafast time-resolved photoluminescence) of 1.6 M green fluorescent protein ( = 1.65±0.06 ns, black), 16-GFP (red,= 1.66±0.05 ns) and a non-covalent mixture of green fluorescent protein and 1 mM

p-benzoquinone (blue, = 0.76±0.04 and 0.14±0.01 ns) in 5 mM phosphate buffer, 5 mMethylenediamine tetraacetic acid, pH 7.0. Excitation with 476 nm laser source and detected at 540 nm at room temperature.

Initial lifetime studies were based on ultrafast time-resolved photoluminescence

(UFTRPL). Subsequent lifetime measurements for both a non-covalent mixture of

anthraquinone-2-carboxylic acid and GFP as well as 12-GFP were performed using

time-correlated single photon counting (TCSPC) which sacrifices temporal resolution in

favour of a greater range and sensitivity compared to UFTRPL. Quenching of

fluorescence decay of anthraquinone samples were observed as shown in Figure 4.13.


4-24

Figure 4.13. Time-resolved fluorescence (time-correlated single photon counting) of a non-covalent mixture of 1.6 M green fluorescent protein and 1 mManthraquinone-2-carboxylic acid ( = 2.04±0.02 and 0.23±0.04 ns, black) and bioconjugate 12-GFP (red, = 1.87±0.02 and 0.28±0.04 ns) in 5 mM phosphate buffer, 5 mM ethylenediamine tetraacetic acid, pH 7.0. Excitation with 476 nm laser source and detected at 515 nm at room temperature.

The non-covalent mixture of GFP and anthraquinone-2-carboxylic acid was modelled as

a double exponential decay using Equation 4.2. Hence, the fluorescence lifetimes of the

non-covalent mixture was estimated to be =2.04±0.02 and 0.23±0.04 ns,

corresponding to native GFP fluorescence and quenching, possibly due to electron

transfer, respectively. The electron transfer rate was determined to be

ket = 3.7±0.7 × 109 s-1 using Equation 4.3. The larger value of the GFP fluorescence

lifetime component of =2.04±0.02 ns compared to the GFP only control

(1.65±0.06 ns) is attributed to the limitation of TCSPC, where the temporal resolution is


4-25

200 ps. Based on the initial fluorescence intensity coefficients, approximately 76%

of the GFP population participated in electron/energy transfer processes.

Similarly, quenching of the bioconjugate 12-GFP fluorescence decay was

observed in Figure 4.13 and modelled according to Equation 4.2 which gave

fluorescence lifetimes of =1.87±0.02 and 0.28±0.04 ns (ket = 2.9±0.4 × 109 s-1),

corresponding to GFP fluorescence and electron transfer, respectively. Based on these

time-resolved fluorescence spectroscopy measurements, electron transfer of the

bioconjugate was observed in contrast to steady-state fluorescence measurements which

did not exhibit red photoconversion. This may be attributed to the high power of the

laser source inducing intramolecular electron transfer of the bioconjugate 12-GFP.

Based on the initial fluorescence intensity coefficients, approximately 62% of the

12-GFP population participated in electron/energy transfer processes. From lifetime

studies, the possible electron transfer pathway is summarised in Scheme 4.6

Scheme 4.6. Schematic representation of the photo-induced electron transfer of green fluorescent protein (GFP, amFP497) at room temperature. (a) Non-covalent mixture of GFP and p-benzoquinone (BQ) or anthraquinone-2-carboxylic acid (AQ) with GFP fluorescent lifetime of = 1.65 ns and electron transfer rates of 6.5 109 and 3.7 109 s-1, respectively. (b) Bioconjugate 12-GFP with electron transfer rate of 2.9 109 s-1.

4.4 Conclusion

In this Chapter, the synthesis of covalent donor-acceptor systems based on green

fluorescent protein (GFP) as a light-induced electron donor were prepared for electron


4-26

transfer studies. For this purpose, viologen-GFP (16-GFP) and anthraquinone-GFP

(12-GFP) bioconjugates were prepared via cysteine-maleimide coupling and amine

modification, respectively. Based on steady-state UV-Vis and fluorescence

spectroscopy studies, evidence of electron transfer was observed by red

photoconversion of the chromophore with p-benzoquinone and anthraquinone acceptor

molecules. Time-resolved fluorescence spectroscopy measurements showed a GFP

fluorescence lifetime of 1.65±0.06 ns. Furthermore, quenching of GFP lifetimes to

0.14±0.01 and 0.23±0.04 ns was observed with non-covalent mixtures of GFP and

p-benzoquinone and anthraquinone-2-carboxylic acid, respectively, corresponding to a

rate constant of forward electron transfer of ket = 6.5±0.5 × 109 and 3.7±0.7 × 109 s-1.

The investigation of photoinduced electron transfer rates between GFP and electron

acceptors have never been reported. However, the donor-acceptor viologen

bioconjugate 16-GFP showed no evidence of photoinduced electron transfer as viologen

acts as a poor electron acceptor in 16-GFP. On the other hand, fluorescence lifetime

studies of the anthraquinone bioconjugate 12-GFP showed possible electron transfer

with a quenched GFP fluorescence lifetime of 0.28±0.04 ns and an electron transfer rate

of ket = 2.9±0.4 × 109 s-1.

4.5 References

(1) Angelo, C.; Denzel, A.; Vogt, A.; Matz, M. V.; Oswald, F.; Salih, A.; Nienhaus, G. U.; Wiedenmann, J. Mar. Ecol. Prog. Ser. 2008, 364, 97.

(2) (a) Tsien, R. Y. Annu. Rev. Biochem. 1998, 67, 509. (b) Lippincott-Schwartz, J.; Patterson, G. H. Science 2003, 300, 87. (c) Alieva, N. O.; Konzen, K. A.; Field, S. F.; Meleshkevitch, E. A.; Hunt, M. E.; Beltran-Ramirez, V.; Miller, D. J.; Wiedenmann, J.; Salih, A.; Matz, M. V. PLoS One 2008, 3, e2680.

(3) Goldstein, J. C.; Munoz-Pinedo, C.; Ricci, J. E.; Adams, S. R.; Kelekar, A.; Schuler, M.; Tsien, R. Y.; Green, D. R. Cell Death Differ. 2005, 12, 453.

(4) Chalfie, M.; Tu, Y.; Euskirchen, G.; Ward, W.; Prasher, D. Science 1994, 263,802.

(5) (a) Wilson, C. G. M.; Magliery, T. J.; Regan, L. Nat. Meth. 2004, 1, 255. (b) Wiedenmann, J.; Oswald, F.; Nienhaus, G. U. IUBMB Life 2009, 61, 1029.


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(6) Shimomura, O.; Johnson, F. H.; Saiga, Y. J. Cell. Comp. Physiol. 1962, 59, 223.(7) Heim, R.; Cubitt, A. B.; Tsien, R. Y. Nature 1995, 373, 663.(8) Yang, F.; Moss, L. G.; Phillips, G. N. Nat. Biotech. 1996, 14, 1246.(9) (a) van Thor, J. J.; Sage, J. T. Photochem. Photobiol. Sci. 2006, 5, 597. (b)

Cotlet, M.; Hofkens, J.; Maus, M.; Gensch, T.; Van der Auweraer, M.; Michiels, J.; Dirix, G.; Van Guyse, M.; Vanderleyden, J.; Visser, A. J. W. G.; De Schryver, F. C. J. Phys. Chem. B 2001, 105, 4999.

(10) Henderson, J. N.; Remington, S. J. Proc. Natl. Acad. Sci. U. S. A. 2005, 102,12712.

(11) Ormö, M.; Cubitt, A. B.; Kallio, K.; Gross, L. A.; Tsien, R. Y.; Remington, S. J. Science 1996, 273, 1392.

(12) Shu, X.; Shaner, N. C.; Yarbrough, C. A.; Tsien, R. Y.; Remington, S. J. Biochemistry 2006, 45, 9639.

(13) Ward, W. W. In Green Fluorescent Protein: Properties, Applications, and Protocols; 2nd ed.; Chalfie, M., Kain, S. R., Eds.; John Wiley & Sons, Inc., 2006; Vol. 47.

(14) Chattoraj, M.; King, B. A.; Bublitz, G. U.; Boxer, S. G. Proc. Natl. Acad. Soc. U. S. A. 1996, 93, 8362.

(15) Bogdanov, A. M.; Mishin, A. S.; Yampolsky, I. V.; Belousov, V. V.; Chudakov, D. M.; Subach, F. V.; Verkhusha, V. V.; Lukyanov, S.; Lukyanov, K. A. Nat. Chem. Biol. 2009, 5, 459.

(16) Sauvage, J. P.; Collin, J. P.; Chambron, J. C.; Guillerez, S.; Coudret, C.; Balzani, V.; Barigelletti, F.; De Cola, L.; Flamigni, L. Chem. Rev. 1994, 94, 993.

(17) Hodgson, D. R. W.; Sanderson, J. M. Chem. Soc. Rev. 2004, 33, 422.(18) (a) Thordarson, P.; Droumaguet, B. L.; Velonia, K. Appl. Microbiol. Biotechnol.

2006, 73, 243. (b) Veronese, F. M.; Pasut, G. Drug Discov. Today 2005, 10,1451.

(19) Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. Angew. Chem. Int. Ed. 2002, 41, 2596.

(20) (a) van Kasteren, S. I.; Kramer, H. B.; Jensen, H. H.; Campbell, S. J.; Kirkpatrick, J.; Oldham, N. J.; Anthony, D. C.; Davis, B. G. Nature 2007, 446,1105. (b) Wang, Q.; Chan, T. R.; Hilgraf, R.; Fokin, V. V.; Sharpless, K. B.; Finn, M. G. J. Am. Chem. Soc. 2003, 125, 3192.

(21) (a) Hvasanov, D.; Goldstein, D. C.; Thordarson, P. In Molecular Solar Fuels;The Royal Society of Chemistry, 2012; p 426. (b) Bertozzi, C. R. Acc. Chem. Res. 2011, 44, 651.

(22) Dirks, A. J.; van Berkel, S. S.; Hatzakis, N. S.; Opsteen, J. A.; van Delft, F. L.; Cornelissen, J. J. L. M.; Rowan, A. E.; van Hest, J. C. M.; Rutjes, F. P. J. T.; Nolte, R. J. M. Chem. Commun. 2005, 4172.

(23) Holmes, T. J.; John, V.; Vennerstrom, J.; Choi, K. E. J. Org. Chem. 1984, 49,4736.

(24) Goldstein, D. C. PhD Thesis, The University of New South Wales, 2011.(25) Sarkar, B.; Wigfield, Y. J. Biol. Chem. 1967, 242, 5572.(26) Himo, F.; Lovell, T.; Hilgraf, R.; Rostovtsev, V. V.; Noodleman, L.; Sharpless,

K. B.; Fokin, V. V. J. Am. Chem. Soc. 2004, 127, 210.(27) Opsteen, J. A.; Brinkhuis, R. P.; Teeuwen, R. L. M.; Lowik, D.; van Hest, J. C.

M. Chem. Commun. 2007, 3136.(28) Gray, H. B.; Winkler, J. R. Annu. Rev. Biochem. 1996, 65, 537.(29) Sawin, K. E.; Nurse, P. Curr. Biol. 1997, 7, R606.


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(30) Takahashi, E.; Takano, T.; Nomura, Y.; Okano, S.; Nakajima, O.; Sato, M. Am. J. Physiol. Cell Physiol. 2006, 291, C781.

(31) Bailey, S. I.; Ritchie, I. M. Electrochim. Acta 1985, 30, 3.(32) Prasad, D. R.; Mandal, K.; Hoffman, M. Z. Coord. Chem. Rev. 1985, 64, 175.(33) (a) Abdul-Ghani, A. J.; Abdul-Kareem, S. J. Photochem. Photobiol., A 1990, 51,

391. (b) Mandal, K.; Hoffman, M. Z. J. Phys. Chem. 1984, 88, 185.(34) Hay, S.; Wallace, B. B.; Smith, T. A.; Ghiggino, K. P.; Wydrzynski, T. Proc.

Natl. Acad. Sci. U. S. A. 2004, 101, 17675.(35) Clark, W. M. Oxidation-reduction potentials of organic systems; Robert E.

Krieger Publishing Company, 1972.(36) (a) Gray, H. B.; Winkler, J. R. Q. Rev. Biophys. 2003, 36, 341. (b) Cordes, M.;

Giese, B. Chem. Soc. Rev. 2009, 38, 892.(37) Striker, G.; Subramaniam, V.; Seidel, C. A. M.; Volkmer, A. J. Phys. Chem. B.

1999, 103, 8612.(38) Peterson, J. R.; Smith, T. A.; Thordarson, P. Chem. Commun. 2007, 1899.(39) (a) Andersson, A.; Danielsson, J.; Gräslund, A.; Mäler, L. Eur. Biophys. J. 2007,

36, 621. (b) Valeur, B. Molecular Fluorescence: Principles and Applications;WILEY-VCH Verlag: Wienheim, Germany, 2002.

(40) Nelissen, H. F. M.; Kercher, M.; De Cola, L.; Feiters, M. C.; Nolte, R. J. M. Chem. Eur. J. 2002, 8, 5407.

Chapter 5

Supramolecular Aggregates for Protein Encapsulation

Chapter 5 Supramolecular Aggregates for Protein Encapsulation

5-2

5 Supramolecular Aggregates for Protein Encapsulation

In this Chapter, the formation of vesicular compartments based on phospholipids and a

diblock copolymer for enzyme encapsulation and light-induced proton pumping studies

is described. Liposomes were prepared using either egg L- -phosphatidylcholine or

synthetic L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine based on variations of the

film hydration method allowing internal encapsulation of fluorescently labelled iso-1

cytochrome c.

Induced formation of polymersomes based on the polyelectrolyte diblock

copolymer, polystyrene140-b-poly(acrylic acid)48 (PS140-b-PAA48) in the presence of

positively charged proteins/peptides allowed concomitant encapsulation of highly

charged hydrophilic proteins with high encapsulation efficiencies.

Vesicles were purified by size exclusion chromatography or membrane dialysis

to remove non-encapsulated material and were characterised by dynamic light

scattering, transmission electron microscopy and confocal laser-scanning microscopy. A

brief review of the vesicle formation methods of liposomes and polymersomes is

outlined in this chapter.

5.1 Liposomes

The term liposome (also termed lipid vesicles) was first introduced by Bangham et al.1

in 1965 which was used to describe aqueous dispersions of multilamellar vesicle (MLV)

systems produced by physical agitation of an aqueous medium in the presence of a dry

lipid film. In modern literature, it is a generic term used to describe polymolecular

aggregates formed in aqueous solution on the dispersion of certain bilayer forming

amphiphilic lipids which may be large or small and may be unilamellar or multilamellar

in nature.2


5-3

Liposomes are spherical shells consisting of one or more (concentric) lamellae

composed of the amphiphiles under osmotically balanced conditions as shown in Figure

5.1.2b The lamellae are curved and self-closed molecular bilayers in which the

hydrophobic portion of the amphiphiles forms the hydrophobic interior of the bilayer

and the polar head group is in contact with the aqueous phase.3 The interior of the lipid

vesicle consists of an aqueous core with an approximate composition of the bulk phase.

The properties of vesicles including mean diameter, lamellarity and physical stability

are generally dependent on the method of vesicle preparation rather than the structure of

natural phospholipid amphiphile used.4

Figure 5.1. Schematic representation of a liposome (lipid vesicle).2b

Liposomes can be prepared based on four primary routes including: film hydration5,

emulsion6, micelle forming detergents7 and organic solvent injection.8 The most

common approach for forming liposomes for enzyme encapsulation is the film

hydration method due to the use of biologically friendly conditions requiring only an

enzyme containing buffered aqueous solution.9 Additionally, variations of the film

hydration method allows customisation of vesicle properties including lamellelarity and

mean size via dispersion (large multilamellar vesicles (LMV), diameters typically


5-4

1 m)10, sonication (small unilamellar vesicles (SUV), diameters typically <50 nm)11,

extrusion (large unilamellar vesicles (LUV), diameters typical of pore size used)12 or

electroformation (LMV, diameters typically 30 m).13 In general, vesicles for enzyme

encapsulation are typically prepared by hydration of a dry lipid film using a buffered

aqueous solution containing the (bio)molecule to be encapsulated resulting in LMV.

The mixture is then subjected to freeze-thaw or lyophilisation steps to increase

encapsulation efficiency and finally sized to form a homogenous mixture by either

sonication or extrusion. Subsequently, the liposome mixture is passed through a size

exclusion column or dialysed to remove non-encapsulated material.4b,c It should be

noted that mechanical treatments during preparation have to be carried out at 5-10 oC or

more above the main lamellar chain-melting phase transition temperature (Tm) where

the saturated hydrophobic chains change from a crystalline state (trans conformation) to

a fluid state (gauche conformation).14

Commonly, liposomes in literature are employed as drug delivery agents,15 gene

vectors16, nanoreactors17 and artificial cells.18 In this project, the ultimate goal is the

development of a photosynthetic-respiratory artificial organelle which requires

compartmentisation via an enclosed membrane which is discussed further in Chapter 7.

Liposomes are ideal for this purpose as LUV ( 100 nm) closely resemble biological

membranes based on surface curvature and membrane fluidity.2b,7

5.1.1 Liposome formation and characterisation

Liposomes based on the phospholipid egg L- -phosphatidylcholine (egg PC) derived

from egg yolk were selected for enzyme encapsulation purposes. Egg PC as shown in

Figure 5.2 was chosen as it is one of the most highly studied amphiphiles in lipid vesicle


5-5

literature2b, commercially available, present in eukaryotic cells19 and possesses a low Tm

(-5.8±6.5 oC)20 leading to a fluid membrane for potential enzyme encapsulation studies.

Figure 5.2. Chemical structure of egg L- -phosphatidylcholine (egg PC) or synthetically derived 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC).

In this project, large unilamellar vesicles (LUV) of ca. 100 nm in diameter are required

as potential compartments for enzyme encapsulation of light-harvesting bioconjugates

discussed in Chapter 2 and the reconstitution of transmembrane cytochrome c oxidase

for charge transfer and proton translocation studies. In order to achieve this, liposomes

from egg PC were prepared by formation of a thin film of dry lipid from a stock

solution in chloroform as shown in Scheme 5.1. The organic solvent was removed under

a stream on nitrogen and residual chloroform was removed under high vacuum for at

least 4 h as presence of trace organic solvent can affect vesicle morphology.3 The lipids

were hydrated in a phosphate buffer (pH 7.0) over 1 h with regular agitation.

Subsequently, the liposome preparation was extruded by passing the sample through a

100 nm polycarbonate membrane to size the vesicles into a homogenous mixture as it

has been reported that extrusion of vesicles through <200 nm pore membranes produce

unilamellar vesicles.12


5-6

Scheme 5.1. Schematic summary of the preparation of liposomes via the film hydration method.

The extruded liposomes were characterised by dynamic light scattering (DLS) to

determine the average hydrodynamic diameter (Dh) of the particles as it is an optimal

technique to determine particle size in dispersions and colloids.21 Particles (liposomes)

cause scattering of the light in all directions when the light passes through the

dispersion. With small particles (compared to the wavelength of the light), the intensity

of scattered light is uniform in all directions due to Rayleigh scattering. In contrast, with

larger particles (above 250 nm diameter), the scattering intensity is angle dependent

(Mie scattering). DLS allows detection of time-dependent fluctuations in the scattered

intensity due to Brownian motion of the particles causing constructive and destructive

interference of light scattered by neighbouring particles using a coherent

monochromatic (laser) source. The diffusion coefficient can be determined based on

analysis of the time dependence of the intensity fluctuation. As a result, the

hydrodynamic diameter of the particles can be elucidated via the Stokes-Einstein

equation, knowing the viscosity of the medium.22

DLS experiments carried out on 1 mg/mL egg PC extruded (100 nm) liposomes

showed a monodisperse sample with an average hydrodynamic radius of 105±12 nm as

shown in Figure 5.3.


5-7

Figure 5.3. Dynamic light scattering (DLS) experiments on 100 nm extruded egg phosphatidylcholine (egg PC) liposomes (1 mg/mL) in 20 mM sodium dihydrogen phosphate, pH 7.0. The average diameter of the aggregates is 105±12 nm.

Freeze-thaw and lyophilisation treatment prior to extrusion has been shown to increase

encapsulation efficiencies.2b In order to confirm that the treatment does not adversely

affect mean size, DLS experiments were also performed on the extruded freeze-thaw

and lyophilised samples as shown in Table 5.1. It was found that lyophilisation prior to

extrusion did not significantly affect mean diameter of the liposomes, however,

freeze-thaw cycles led to small average diameters (75±7 nm). This may be a result of

fracture or rupture of the vesicles leading to a change in size distribution.


5-8

Table 5.1. Mean size of different treatment methods of egg phosphatidylcholine liposomes.Treatment Mean hydrodynamic diameter (Dh) /nma

Extrusion 105±12Freeze-thaw/Extrusion 75±7Lyophilisation/Extrusion 114±8

a Experiments carried out on 1 mg/mL liposomes in 20 mM sodium dihydrogen phosphate, pH 7.0 at 25 oC. Errors are standard deviation.

Additionally, liposomes were characterised by transmission electron microscopy (TEM)

to confirm structural morphology and lamellarity. Due to the poor electron scattering

properties of phospholipids for TEM imaging, samples were negatively stained using

2% phosphotungstic acid. Negative stains are composed of salt solutions of strongly

electron scattering heavy metal compounds which are added to the drying specimen

which forms a mold of the specimen.23 TEM studies showed vesicular morphologies for

extruded liposomes as shown in Figure 5.4a. However, multilamellar vesicles were

observed even though it has been reported in literature that extrusion produces

unilamellar vesicles.12 To ascertain whether further mechanical treatment of the

liposomes produces unilamellar vesicles, TEM micrographs of freeze-thawed and

lyophilised specimens prior to extrusion were imaged; Figure 5.4b and c, respectively,

which showed multilamellar structural features.


5-9

Figure 5.4. TEM micrographs of egg phosphatidyl (egg PC) liposomes (0.5 mg/mL). (a) 100 nm pore extrusion. (b) Freeze-thawed and extruded. (c) Lyophilised and extruded. Specimens negatively stained with 2% phosphotungstic acid. Scale bars: 50 nm. Arrows indicate multilamellar structures.

It has been reported that naturally isolated egg PC contains negatively charged

impurities.24 Instead, synthetically isolated L-palmitoyl-2-oleoyl-sn-glycero-3-

phosphocholine (POPC) which is chemically identical to the natural product was used

for liposome formation to eliminate the possibility that impurities were causing

multilamellar features. To determine whether the multilamellar features are a result of

the use of 2% phosphotungstic acid, an alternative negative stain, 2% uranyl acetate was

used for POPC liposomes as shown in Figure 5.5a and b, respectively. Based on TEM

micrographs of Figure 5.5, identical multilamellar features were observed compared to

2% phosphotungstic acid stains of egg PC liposomes (Figure 5.4).


5-10

Figure 5.5. TEM micrographs of negatively stained L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) liposomes (0.5 mg/mL). (a) 2% phosphotungstic acid. (b) 2% uranyl acetate. Scale bars: 100 nm. Arrows indicate multilamellar structures.

To eliminate the use of negative stains and drying of sample during TEM preparation,

cryogenic TEM (cryo-TEM) was employed in collaboration with A/Prof. Filip Braet at

the Australian Centre for Microscopy & Microanalysis (The University of Sydney).

Cryo-TEM allows the imaging of suspended organic material in aqueous solution.25

Thin films (ca. 100 nm) of suspended sample on a TEM grid is plunged into liquid

ethane at -183 oC using an automated robot (vitrobot) which allows the nanostructures

to become instantaneously embedded in an electron transparent film of vitrified

amorphous ice.26 This allows samples to be analysed in their near native hydrated

state.27 As shown in Figure 5.6, it was found that 70% of the extruded (100 nm) POPC

liposomes were unilamellar and the morphology were generally typically ‘filled-cups’.

This demonstrates that staining of liposomes and the drying process was responsible for

multilamellar features resulting in electron micrograph artifacts.


5-11

Figure 5.6. Cryo-TEM micrograph of L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) liposomes. Typically ‘filled-cup’ morphologies with 70% unilamellarity. Scale bars: 50 nm. Arrows indicate unilamellar vesicle.

5.1.2 Enzyme encapsulation

In order to evaluate liposomes as potential compartments for light-induced electron

transfer studies of light-activated bioconjugates (discussed further in Chapter 6),

enzyme encapsulation studies of iso-1 cytochrome c as a model enzyme were

performed. Initially, to determine if cytochrome c could be successfully encapsulated in

the interior volume of POPC liposomes, fluorescently labelled iso-1 cytochrome c

(cyt c-Oregon Green 488) was encapsulated and monitored optically via confocal

laser-scanning microscopy (CLSM).

Encapsulation was achieved by adding cyt c-Oregon Green 488 (5 M) in 20 mM

sodium dihydrogen phosphate buffer, pH 7.0, to a dry lipid film resulting in a lipid

concentration of 15 mg/mL. Subsequently, the dispersion was lyophilised to increase

encapsulation efficiency (method with highest encapsulation efficiencies)2b, rehydrated

and extruded. It should be noted that extrusion was performed using 400 nm

polycarbonate membranes rather than 100 nm pores to allow visualisation of aggregates

as 100 nm particles are below the Abbe diffraction limit ( 250 nm, assuming numerical


5-12

aperture NA of 1.4).28 Non-encapsulated enzymes were removed by size exclusion

chromatography (Superdex 75) and imaged by CLSM as shown in Figure 5.7,

demonstrating successful encapsulation.

Figure 5.7. Transmission light (a) and confocal laser-scanning micrograph (b)-excitation 488 nm-of enzyme encapsulated fluorescently labelled cytochrome c(cyt c-Oregon Green 488) in L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) liposomes. Scale bars: 50 m.

Interestingly, it was found that purification of liposomes by size exclusion

chromatography (approximately three purification cycles) resulted in gradual blockage

of the column and required cleaning using an aqueous 70% v/v ethanol solution as

shown in Figure 5.8.


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Figure 5.8. Size exclusion chromatogram showing separation of fluorescently labelled cytochrome c-containing vesicles (peak A) from non-entrapped enzyme (peak B) using Superdex 75 (GE Healthcare) in 20 mM sodium dihydrogen phosphate buffer, pH 7.0 at 0.5 mL/min. Red trace indicates blockage of column preventing elution of liposomes. Black trace indicates elution of liposomes after 70% ethanol (v/v) wash. Elution was monitored at optical density 280 nm.

Evaluation of POPC as a potential compartment for enzyme encapsulation was screened

by quantifying the encapsulation efficiency of unmodified iso-1 cytochrome c (cyt c).

The encapsulation efficiency (EE(%)) was quantified based on Equation 5.1 and is

defined as the percentage amount of enzyme entrapped in the vesicles in relation to the

total amount of enzyme present during the vesicle formation and entrapment

procedure.2b,3 The EE(%) is dependent on a number of factors including method of

preparation, concentration of amphiphile used, chemical nature of amphiphile and

concentration and nature of (bio)molecule to be encapsulated.

100(%)usedenzymeofamounttotal

vesicleslipidtheinentrappedenzymeofamountEE (5.1)


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Encapsulation was achieved by adding cytochrome c (100 M) in 20 mM sodium

dihydrogen phosphate buffer, pH 7.0, to a dry lipid film resulting in a lipid

concentration of 15 mg/mL. The dispersion was lyophilised, rehydrated and extruded

through a 100 nm membrane. Subsequently, the enzyme containing liposomes were

purified using size exclusion chromatography (Superdex 75) in 20 mM sodium

dihydrogen phosphate buffer, pH 7.0, as shown in Figure 5.9. To ensure maximum

recovery of encapsulated enzyme, the column was first pre-cleaned with 70% v/v

ethanol solution before loading liposomes onto column.

Figure 5.9. Size exclusion chromatogram showing separation of iso-1cytochrome c-containing vesicles (peak A) from non-entrapped enzyme (peak B) using Superdex 75 (GE Healthcare) in 20 mM sodium dihydrogen phosphate buffer, pH 7.0 at 0.5 mL/min.


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The EE(%) was determined by collecting the purified liposomal fraction (peak A) and

estimating the concentration of encapsulated cyt c by UV-Vis spectroscopy using the

heme absorption band ( 410 = 97.6 mM-1cm-1).29 Due to scattering of the UV-Vis

baseline caused by liposome aggregates, the vesicles were disrupted by addition of

surfactant Triton X-100 (2%, v/v) in a ratio of 1:1 (v/v)30 and a low EE(%) of 1.5±0.1%

was determined. The maximum theoretical EE(%) under identical conditions and

assuming a random statistical distribution with an average POPC lipid head group area

of 69 31 was 7.0% (see Appendix D). A random statistical distribution was assumed as

there are no charge interactions between lipid and cytochrome c due to the zwitterionic

nature of POPC. Interestingly, from DLS studies, it was found that the average diameter

of the enzyme encapsulated vesicles was greater than enzyme free POPC liposomes

with diameters of 96±1 nm and 100±3 nm, respectively, as shown in Figure 5.10. This

is most likely due to accommodation of enzyme within the interior volume of the

compartment.


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Figure 5.10. Dynamic light scattering (DLS) experiments on lyophilised-100 nm extruded L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) liposomes (1 mg/mL) in 20 mM sodium dihydrogen phosphate, pH 7.0. The average diameter of the aggregates is 96±1 nm and 100±3 nm for enzyme free (black) and enzyme encapsulated (red) vesicles, respectively.

5.2 Polymersomes

One of the major disadvantages of liposomes is their poor thermodynamic and

mechanical properties resulting in morphology changes, degradation and leakage of

internal contents mainly due lipid peroxidation or hydrolysis when stored in aqueous

solution.32 In contrast, polymersomes overcome this inherent instability due to their

higher molecular weight, lower critical aggregation concentration, slower chain

mobility and polymer entanglement.33 Polymersomes are generally based on diblock or

triblock copolymer amphiphiles.34 Polymersomes have been used for the encapsulation

of drugs35, biomolecules as nanoreactors17, protein therapeutic applications for

biomedicine36 or artificial organelles.37


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Polymersomes are part of a continuum of supramolecular structures consisting

of micelles, rods, lamellae and vesicles.38 Multiple factors can be tailored to influence

resulting structures including temperature, solvent, block length, ratio of the blocks, pH

and ionic strength which affects the critical packing parameter (p = v/aolc), where v is

the volume of the hydrocarbon chains, ao is the optimal area of the hydrophilic (corona)

group and lc is the critical chain length of the hydrophobic (core) group as shown in

Figure 5.11.39 In this project, polymersome aggregate studies used the well-known

diblock copolymer polystyrene-b-poly(acrylic acid) (PS-b-PAA).40 It has been reported

that the major factor influencing PS-b-PAA morphologies was the length of the

hydrophilic block.41 Diblock copolymers such as PS-b-PAA self-assemble when they

are mixed with a solvent selective for one of the blocks, resulting in aggregation and

phase separation.42

Figure 5.11. Different morphologies predicted by the packing parameter (p). It is predicted that micelles form when p<1/2, vesicles when 1/2<p<1 and inverted structures are expected when p>1.17

Polymersomes are traditionally prepared using the ‘thermodynamic trapping’ method

with the block copolymer dissolved in an organic common solvent and water then

slowly added to induce aggregate morphologies, which is a non-biologically friendly

technique.43 Alternatively, they can be prepared using a biologically friendly ‘kinetic


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trapping’ method44, which involves injecting the dissolved block copolymer in organic

solution directly into the aqueous buffer. In this Chapter, both methodologies are

studied using commercially available PS140-b-PAA48 with a polydispersity index (PDI)

of 1.10 to determine which preparation method is ideal and robust for enzyme

encapsulation.

5.2.1 Aggregate formation using the ‘thermodynamic trapping’ method

Polymersome formation using PS140-b-PAA48, as shown in Figure 5.12, was attempted

using the ‘thermodynamic trapping’ method, also known as the slow addition method.

The role of solvent is crucial to the free energy of aggregation and polymer-solvent

interactions determine coil dimensions for each block of the diblock copolymer, which

affects the resulting aggregate dimensions and morphologies.39a Tetrahydrofuran (THF)

and dioxane were used as solvents for aggregation studies as they are a common solvent

for both blocks of the copolymer. Additionally, the solubility parameter of the solvents

is similar to the polymer.45

Figure 5.12. Chemical structure of polystyrene140-b-poly(acrylic acid)48(PS140-b-PAA48) with a polydispersity index (PDI) of 1.10.

Polymeric aggregates were prepared by adding water dropwise over a 3 h period

into a dissolved polymer solution (10 mg/mL) in THF or dioxane until a turbid solution

formed, indicating formation of polymeric aggregates. Finally, the turbid solution was

dialysed against water extensively to remove the organic solvent. The aggregates were

characterised by TEM microscopy without the use of negative stains due to the electron

scattering properties of the PS block.46 It should be noted that polymer aggregate TEM


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studies required a lower accelerating voltage (80 kV) compared to liposomes as higher

energy electron beams caused deformation and damage to the specimen. Only micelle

and large compound micelle structures were observed as shown in Figure 5.13. Due to

the unsuccessful formation of polymersome structures via the ‘thermodynamic trapping’

method and the biologically unfriendly conditions required, further aggregate studies

using this method was not pursued. As a result, the ‘kinetic trapping’ method was

investigated for enzyme encapsulation which is discussed in the next section (Chapter

5.2.2) and THF was exclusively used in further polymersome aggregate studies due to

the possible carcinogenic effects of dioxane.47

Figure 5.13. TEM micrographs of polysytrene140-b-poly(acrylic acid)48(PS140-b-PAA48) aggregates. (a) Tetrahydrofuran (THF). (b) Dioxane. Scale bars: 200 nm.

5.2.2 Polymersome formation using the ‘kinetic trapping’ method

It was found that polymersome formation could be induced from negatively charged

PS140-b-PAA48 using proteins and peptides with a complementary positive charge to

drive the formation of vesicles over micelles in this system under ‘kinetic trapping’

conditions.† This method of membrane encapsulation for biomolecules overcomes the

issues of traditional polymersome encapsulation, as discussed in the previous section

† Parts of this work have been published: Hvasanov, D.; Wiedenmann, J.; Braet, F.; Thordarson, P. Chem. Commun. 2011, 47, 6314.


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(Chapter 5.2.1), including denaturation of enzymes by harsh preparation conditions and

offers functionalities that polyelectrolyte capsules lack.48 For example, large internal

volumes could encapsulate a second functional (biological) system within the cavity for

cascade reactions as organelle mimics.34b The proteins and peptides employed in this

study induced polymersome formation and concurrently efficiently encapsulated in this

polymer vesicle system.

Aggregates were prepared by injection of a dissolved PS140-b-PAA48 in THF

into an enzyme containing phosphate buffer with the syringe submerged in solution at a

THF:water ratio of 1:6 (v/v). The solution was equilibriated for 24 h and dialysed

(50 kDa molecular weight cut-off) against water extensively to remove

non-encapsulated enzyme and THF. Due to the robust nature of polymer aggregates,

purification of enzyme-containing polymersomes via size exclusion chromatography is

not possible.

In this work, polymeric vesicles are defined as morphologies displaying distinct

contrast within individual particles indicating the presence of bilayer and cavity

formation, whereas micelles show uniform contrast. The addition of 5 M of a

positively charged biomolecule such as iso-1 cytochrome c to 8 M of PS140-b-PAA48

is very effective in promoting polymersome formation. Evidence of the dependence of

polymersome formation on positively charged proteins or peptides has been confirmed

by TEM microscopy studies as shown in Figure 5.14.49


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Figure 5.14. TEM micrographs of polysytrene140-b-poly(acrylic acid)48(PS140-b-PAA48) aggregates in the presence of 5 M additives in PBS (25 oC). (a) PBS control (no enzyme). (b) Iso-1 cytochrome c. (c) Green fluorescent protein. (d) Poly-L-lysine. (e) Myoglobin. (f) Bovine serum albumin. (g) Calmodulin. Scale bars: 200 nm.

The polymer aggregates were further characterised by cryo-TEM as shown in Figure

5.15 to conclusively verify aggregate morphologies. The predominant observed

morphology were micelles in the presence of buffer control (Figure 5.15a), consistent

with classical TEM analysis (Figure 5.14). Additionally, the electron micrographs

confirm the formation of vesicles when formed with positively charged additives such

as iso-1 cytochrome c (cyt c) (Figure 5.15c), green fluorescent protein (GFP) (Figure

5.15d) and poly-L-lysine (Figure 5.15e).


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Figure 5.15. Cryo-TEM micrographs of polysytrene140-b-poly(acrylic acid)48(PS140-b-PAA48) aggregates in the presence of 5 M additives in PBS (25 oC). (a) PBS control (no enzyme). (b) Calmodulin. (c) Iso-1 cytochrome c. (d) Green fluorescent protein. (e) Poly-L-lysine. Scale bars: 50 nm.

Interestingly, using negatively charged biomolecules such as bovine serum albumin

(BSA) and calmodulin (Figure 5.15b) in PBS resulted in micelle structures while with

the neutral myoglobin (Mb) inducer, an approximate 1:1 mixture of polymersomes and

micelles is obtained (Table 5.2).


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Table 5.2. Effect of polymersome formation with different biomolecules at physiological pH.Proteina M.W. /kDa pI Charge Morphologyb

None (H2O) N/A N/A N/A MicellesNone (PBS) N/A N/A +/- Micellesc

Cyt c50 12.7 10.6 + VesiclesCyt cd (30 min) 12.7 10.6 + VesiclesCyt cd (24 h) 12.7 10.6 + MicellesGFP49 26.8e 8.3f + VesiclesPoly-L-lysine 1-5 N/A + VesiclesMb51 17.6 7.3 Neutral Micelles/Vesiclesg

BSA52 66 4.7 - MicellesCalmodulin53 16.7 3.9-4.3 - Micelles

a Conditions: enzyme in buffered PBS, pH 7.2. b Determined by TEM (see also Figure 5.14). c Asmall amount of vesicles observed. d Post-addition to mixture. e GFP can form non-covalent dimers and tetramers. f pI for GFP estimated by theoretical calculations of peptide sequence. g

Mb displayed 1:1 ratio of micelles:vesicles.

The combined addition of salts and positively charged biomolecules (cyt c), induces

electrostatic shielding along the partially ionised corona block which decreases the coil

dimension and reduces steric hindrance.39a,54 This allows more polymer chains to

aggregate, leading to increased core-chain stretching, resulting in structural changes

from micelles to rods to vesicles due to unfavourable thermodynamics associated with

excessive stretching. Interestingly, post-addition of cytochrome c to a polymer micelle

mixture after 30 minutes produces a mixed population of polymersomes and amorphous

aggregates compared to micelles and delayed post-addition after 24 hours has no effect

on resulting micelle morphologies (Table 5.2 and Figure 5.16).


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Figure 5.16. TEM micrographs of polystyrene140-b-poly(acrylic acid)48(PS140-b-PAA48) aggregates in the presence of 5 M cytochrome c after post-addition to polymer micelle mixture in PBS (25 oC). (a) 30 min. (b) 24 h. Scale bars: 200 nm.

Only micelles are observed with PS140-b-PAA48 in the presence of BSA and

calmodulin. Further, there is no indication of BSA or calmodulin interacting with the

micelles in contrast to positively charged biomolecules. This suggests for negatively

charged proteins, there is electrostatic repulsion between the corona chains in

PS140-b-PAA48. The net result is that the aggregation of polymer chains and eventual

formation of polymersomes due to increased core-chain stretching is inhibited, leading

to micelle morphologies.

Given the variation in molecular weight and 3-dimensional structure of the

biomolecules screened, the primary factor causing vesicle formation are electrostatic

interactions between the biological and synthetic polymers rather than templation of the

surface curvature of the proteins. This is consistent with the myoglobin (Mb) results,

whereby, the aggregates form an approximate 1:1 ratio between micelles and

polymersomes. This is a result of the near neutrality of Mb with only a few positively

charged residues at pH 7.2 (pI = 7.3)51 even though it is fairly similar in size and shape

to cytochrome c (pI = 10.6).50 The induced polymersome formation can be generally

summarised as shown in Scheme 5.2.


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Scheme 5.2. Schematic showing effect of charged biomolecule additives on polysytrene140-b-poly(acrylic acid)48 (PS140-b-PAA48) aggregate morphologies. Negatively charged biomolecules lead to micelles and positively charged additives induce polymersome structures.

5.2.2.1 Optimisation of polymersome formation

In order to determine optimum conditions for polymersome formation, such as minimal

structural defects and larger encapsulation volumes in a sample population, different salt

buffer conditions and temperatures were selected as shown in Table 5.3. It can be seen

that the optimum parameters required for polymersome preparation was room

temperature incubation in the presence of phosphate buffered saline (PBS). It is

interesting to observe that higher incubation temperatures and lower salt concentration

results in poorly formed polymersomes with smaller vesicle diameters. Lower salt

concentration buffers (20 mM NaH2PO4, pH 7.0) reduces the degree of electrostatic

repulsion causing smaller vesicles to form compared to PBS (pH 7.2) with sizes

162±61 nm and 376±259 nm at room temperature, respectively. In addition, the increase

in buffer temperature increases the rate of evaporation of the organic co-solvent

plasticiser (THF). As a result, the vesicles are ‘frozen’ over a shorter time scale

compared to ambient temperature due to decreasing core-chain mobilities leading to

smaller vesicles being formed.46 It should be noted that the size determination of

polymersomes were analysed using TEM microscopy. DLS was unsuitable for average


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diameter determination of polymersomes as the aggregates were highly polydisperse,

resulting in poor data quality.

Table 5.3. Summary of salt buffer and temperature effects on polymersome formation.Enzymea Buffer T /oC Size /nm EE(%)b

Cyt c 20 mM NaH2PO4, pH 7.0 25 162±61 55±1Cyt c PBS, pH 7.2 25 376±259 66±7Cyt c 20 mM NaH2PO4, pH 7.0 40 96±25 -Cyt c PBS, pH 7.2 40 267±193 -GFP PBS, pH 7.2 25 308±136 35±1

a Enzymes prepared at 5 M concentration. b Determined for optimally formed polymersomes. EE(%) estimated using fluorescently labelled cyt c. Errors are standard deviation.

5.2.2.2 Concentration dependence of positively charged biomolecules

In order to probe the mechanism of polymersome formation, the percentage ratio of

polymersome-to-micelle aggregates were determined relative to the ratios of the

positive charge composition F+:55

]charges[]charges[charges][1 FF (5.2)

assuming 70% partial ionisation of the poly(acrylic acid) block ( 8 M) at pH 7.2.56

Additionally, yeast iso-1 cytochrome c has +14 charges (14 lysine residues)57 and green

fluorescent protein (Acropora millepora) has +17 charges (17 lysine residues).58 From

Figure 5.17, it can be seen that there is rapid binding leading to significant

polymersome formation with up to 60% formation with less than F+ 0.05 ( 1 M) for

both cytochrome c and green fluorescent protein due to the strong charge interactions

between corona and biomolecule.59 This is followed by saturation after addition of

greater than F+ 0.15 for both additives.§ At the saturation point, this is equivalent to an

approximate molar ratio of 2:1 polymer chain-to-enzyme.

§ Each data point is performed as independent experiments.


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Figure 5.17. Influence of different enzyme inducers on relative polymersome/micelle (p/m) formation as a function of positive charge composition (F+) for cytochrome c ( )and green fluorescent protein ( ). Error bars are standard deviation.

Additionally, box plot analysis of the effect of charge ratio F+ on the size of

polymersomes for cytochrome c (Figure 5.18a) and green fluorescent protein (Figure

5.18b) shows that the polymersome size is independent of F+.


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Figure 5.18. Influence of different enzyme inducers on dependence of polymersome size. (a) Cytochrome c. (b) Green fluorescent protein. Boxes extend to 25th and 75th and whiskers to 5th and 95th percentile, respectively. Red horizontal line marks the median.

5.2.2.3 Model of polymersome formation

Based on these observations, it is proposed that the positively charged cytochrome c and

green fluorescent protein are bound electrostatically to the coronal PAA block as shown

in Figure 5.19, causing aggregation of polymer chains and its vesicle structure. The 2:1

ion-pair supramolecular complex induces contraction of the effective PAA block length,

due to neutralisation of the PAA coronal chain with the multiply charged

cytochrome c57 and green fluorescent protein58 leading to micelle to vesicle

morphological transitions.39a This ion-pair interaction is reminiscent of gemini

surfactants, shown to form vesicles for long spacer groups between individual

amphiphiles consistent with the model.60


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Figure 5.19. Proposed model for polymersome formation (PS140-b-PAA48) as a 2:1 ion-pair supramolecular complex. (a) Cytochrome c. (b) Green fluorescent protein.

Owing to the highly polydisperse nature of the polymersomes, diameters as large as

3 m have been observed as shown in Figure 5.20, allowing optical visualisation of

coronal encapsulation via fluorescence.

Figure 5.20. Representative histogram of PS140-b-PAA48 polymersomes for 5 Mcytochrome c in PBS at 25 oC.


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The proposed model is consistent with confocal laser-scanning microscopy (CLSM)

images which show fluorescently labelled cyt c (cyt c-Oregon Green 488) and GFP

encapsulated in the membrane as shown in Figure 5.21.

Figure 5.21. Transmission light (a) and (c) and confocal laser scanning microscopy images (b) and (d) – excitation 488 nm – of enzymes encapsulated in the membrane of PS140-b-PAA48 polymersomes for fluorescently labelled cytochrome c (a) and (b) and green fluorescent protein (c) and (d). Scale bars: 50 m and inset 5 m. Contrast enhanced.

5.2.2.4 Biological stability

The membrane encapsulation of proteins and biomolecules into the polymersomes via

the injection method requires the presence of co-solvent, THF. In order to ensure that

the encapsulated enzymes are not denatured and remain biologically active for

encapsulation of light-harvesting bioconjugates and photo-induced charge transfer

studies further discussed in Chapter 6 and 7, biological stability studies were performed.


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The photophysical properties of the proteins studied including green fluorescent

protein and cytochrome c were monitored via fluorescence or UV-Vis spectroscopy,

respectively, to ensure that the emission band or absorption bands were not quenched or

shifted, indicating correct folding of the protein tertiary structure after polymersome

formation. It was found that the encapsulation of GFP maintained its native folded state

as evident by its preserved fluorescence as shown in Figure 5.21d and Figure 5.22.

Figure 5.22. Fluorescence emission profiles of green fluorescent protein (GFP) excited at 476 nm with emission maxima 497 nm. Solution GFP (solid red line), encapsulated GFP (dashed black line).

The stability of cytochrome c conformation was monitored using the Soret band at

410 nm indicating that the tertiary structure was preserved as shown in Figure 5.23.61

Significant blue shifting (>10 nm) of the Soret band indicating protein denaturation was

not observed.


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Figure 5.23. UV-Vis spectra of soret band (410 nm) of cytochrome c (cyt c). Solution cyt c (solid red line) and encapsulated cyt c (dashed black line). Sloping baseline for encapsulated cyt c due to scattering by polymersome aggregates.

The catalytic activity of cytochrome c has been widely reported with specific attention

on the oxidation of 2,2’-azino-bis[ethyl-benzothiazoline-(6)-sulfonic acid] (ABTS).62

Samples were prepared to 0.5 M solution (bulk) cytochrome c or encapsulated

cytochrome c, 200 M ABTS, 20 mM phosphate buffer at pH 7.0 and initiated with the

addition of hydrogen peroxide to a final concentration of 10 mM. Cytochrome c was

catalytically active after encapsulation by monitoring oxidation of ABTS at 415 nm as

shown in Figure 5.24.61 Interestingly, the encapsulated cytochrome c had an initial

increase in catalytic activity compared to native enzyme in solution due to sequestering

of substrate towards the enzyme.63


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Figure 5.24. Catalytic activity of cytochrome c using the ABTS assay. (a) Chemical structure of 2,2’-azino-bis[ethyl-benzothiazoline-(6)-sulfonic acid] (ABTS). (b) Schematic of catalytic oxidation of ABTS using cytochrome c (cyt c). (c) Catalytic activity of solution cyt c (solid red line) and encapsulated cyt c (dashed black line) by monitoring oxidation of ABTS at 415 nm.

5.2.2.5 Enzyme encapsulation efficiencies

It is usually difficult to achieve high encapsulation efficiencies with polymersomes, due

to thicker membranes (d 8-21 nm) compared to liposomes (d 3-5 nm).36 Highly

polydisperse membrane thicknesses of 120±117 nm for PS140-b-PAA48 were observed

and thicker than previously reported for PS-b-PAA systems.38 This may be due to the


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positively charged biomolecules acting as electrostatic bridges resulting in multilamellar

vesicles. A bilayer of PS140-b-PAA48 in the fully extended state is 92 nm.46

Furthermore, the bridging ability of these inducers is supported by observations of

polymersome aggregation and cluster formation in electron (Figure 5.14) and confocal

microscopy (Figure 5.21).

The method of encapsulation of biomolecules within the coronal block by

induction of positively charged additives overcomes the inherent disadvantage of

conventional polymersome encapsulation methods due to their thicker membranes.

Encapsulation efficiencies as high as 55±1% and 66±7% in 20 mM NaH2PO4 and PBS,

respectively, for cytochrome c and 35±1% for green fluorescent protein were observed

for optimised polymersomes (Table 5.3). It should be noted that the EE(%) of enzymes

in polymersomes was performed using steady-steady fluorescence of fluorescently

labelled cytochrome c (cyt c-Oregon Green 488) and native fluorescence of green

fluorescent protein to estimate loading (see Appendix D). This is due to the fact that

unlike liposomes, polymersomes are robust and treatment with a surfactant such as

Triton X-100 does not disassemble the aggregates which contributes to scattering of the

UV-Vis spectra. Hence, fluorescence emission provides a more accurate EE(%)

estimate.

5.3 Conclusion

In this Chapter, the formation of vesicles as compartments for enzyme encapsulation

and potential light-induced charge transfer studies were achieved. Lipid vesicles were

formed using the film hydration method and sized via extrusion to produce

monodisperse unilamellar vesicles of ca. 100 nm in diameter. Confirmation of

monodispersity and average diameter were achieved using dynamic light scattering


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techniques. Additionally, unilamellar vesicles were characterised using traditional TEM

microscopy and cryo-TEM techniques and it was found that structural characterisation

of lamellar features using conventional TEM microscopy introduced artifacts as a result

of the necessity of negative stains and drying of specimen. The possibility of liposomes

based on egg PC and POPC as potential compartments were evaluated by the

encapsulation of model protein cytochrome c, which is used as an electron transfer

metalloprotein for photo-active bioconjugates. Encapsulation was confirmed using

confocal laser-scanning microscopy (CLSM) of fluorescently labelled cytochrome c.

However, low encapsulation efficiencies of 1.5±0.1% precluded their use for further

bioconjugate encapsulation studies.

Alternatively, polymersomes were formed as potential compartments. A novel

method of polymersome preparation by the injection method which induces

polymersome aggregates with the polyelectrolyte-based diblock copolymer,

PS140-b-PAA48, in a facile and biologically friendly manner was demonstrated.

Concomitantly, encapsulation of positively charged biomolecules occurs within the

coronal block. Additionally, this induced formation is versatile and can be applied to

various positively charged biomolecules with high encapsulation efficiencies allowing

for potential applications as nanoreactors64 (further discussed in Chapter 6 and 7) or

protein therapeutic vessels including use of polycationic copolymers as non-viral

vectors for DNA/RNA gene therapy65 and simplified preparation compared to

polyelectrolyte microcapsules.66 Similar to liposomes, polymersomes were

characterised by conventional and cryo-TEM microscopy as well as CLSM. The

biological activities of encapsulated enzymes were demonstrated to remain active using

spectroscopy studies and the ABTS assay. Due to the high encapsulation efficiencies of

55±1% and 66±7% in 20 mM NaH2PO4 and PBS, respectively, for cytochrome c and


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35±1% for green fluorescent protein, polymersomes were selected as the most suitable

compartments for enzyme encapsulation.

5.4 References

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1986, 858, 161. (b) Walde, P.; Ichikawa, S. Biomol. Eng. 2001, 18, 143.(3) Ostro, M. J. Liposomes; Marcel Dekker, Inc.: New York, 1983.(4) (a) Haluska, C. K.; Riske, K. A.; Marchi-Artzner, V.; Lehn, J.-M.; Lipowsky,

R.; Dimova, R. Proc. Natl. Acad. Soc. U. S. A. 2006, 103, 15841. (b) Blocher, M.; Walde, P.; Dunn, I. J. Biotechnol. Bioeng. 1999, 62, 36. (c) Gaspar, M. M.; Perez-Soler, R.; Cruz, M. E. M. Cancer Chemother. Pharmacol. 1996, 38, 373.

(5) Walde, P.; Marzetta, B. Biotechnol. Bioeng. 1998, 57, 216.(6) Szoka, F.; Papahadjopoulos, D. Proc. Natl. Acad. Soc. U. S. A. 1978, 75, 4194.(7) Enoch, H. G.; Strittmatter, P. Proc. Natl. Acad. Soc. U. S. A. 1979, 76, 145.(8) Schenning, A. P. H. J.; Escuder, B.; van Nunen, J. L. M.; de Bruin, B.; Löwik,

D. W. P. M.; Rowan, A. E.; van der Gaast, S. J.; Feiters, M. C.; Nolte, R. J. M. J. Org. Chem. 2000, 66, 1538.

(9) Das, N.; Gupta, S.; Mazumdar, S. Biochem. Biophys. Res. Commun. 2001, 286,311.

(10) Lichtenberg, D.; Markello, T. J. Pharm. Sci. 1984, 73, 122.(11) Fuhrhop, J.-H.; Mathieu, J. Angew. Chem. Int. Ed. 1984, 23, 100.(12) Mui, B.; Chow, L.; Hope, M. J. Methods Enzymol. 2003, 367, 3.(13) Angelova, M. I.; Dimitrov, D. S. Faraday Discuss. Chem. Soc. 1986, 81, 303.(14) Yeagle, P. The Structure of Biological Membranes; CRC Press, 2011.(15) Touitou, E.; Junginger, H. E.; Weiner, N. D.; Nagai, T.; Mezei, M. J. Pharm.

Sci. 1994, 83, 1189.(16) Ewert, K.; Slack Nelle, L.; Ahmad, A.; Evans Heather, M.; Lin Alison, J.;

Samuel Charles, E.; Safinya Cyrus, R. Curr. Med. Chem. 2004, 11, 133.(17) Vriezema, D. M.; Comellas Aragonès, M.; Elemans, J. A. A. W.; Cornelissen, J.

J. L. M.; Rowan, A. E.; Nolte, R. J. M. Chem. Rev. 2005, 105, 1445.(18) (a) Steinberg-Yfrach, G.; Liddell, P. A.; Hung, S.-C.; Moore, A. L.; Gust, D.;

Moore, T. A. Nature 1997, 385, 239. (b) Steinberg-Yfrach, G.; Rigaud, J.-L.; Durantini, E. N.; Moore, A. L.; Gust, D.; Moore, T. A. Nature 1998, 392, 479. (c) Kurihara, K.; Tamura, M.; Shohda, K.-i.; Toyota, T.; Suzuki, K.; Sugawara, T. Nat. Chem. 2011, 3, 775.

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91.(21) Ottaviani, M. F.; Matteini, P.; Brustolon, M.; Turro, N. J.; Jockusch, S.;

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M. F.; Favuzza, P.; Bigazzi, M.; Turro, N. J.; Jockusch, S.; Tomalia, D. A. Langmuir 2000, 16, 7368.


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(24) Dimitrova, M. N.; Tsekov, R.; Matsumura, H.; Furusawa, K. J. Colloid Interface Sci. 2000, 226, 44.

(25) Taylor, K. A.; Glaeser, R. M. Science 1974, 186, 1036.(26) Dubochet, J.; Chang, J. J.; Freeman, R.; Lepault, J.; McDowall, A. W.

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University Press: New York, 2011.(29) Peterson, J. R.; Thordarson, P. Chiang Mai J. Sci. 2009, 26, 236.(30) Kato, T.; Higuchi, M.; Endo, R.; Maruyama, T.; Haginoya, K.; Shitomi, Y.;

Hayakawa, T.; Mitsui, T.; Sato, R.; Hori, H. Pestic. Biochem. Physiol. 2006, 84,1.

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Commun. 2000, 1433. (b) Vriezema, D. M.; Garcia, P. M. L.; Sancho Oltra, N.;Hatzakis, N. S.; Kuiper, S. M.; Nolte, R. J. M.; Rowan, A. E.; van Hest, J. C. M. Angew. Chem. Int. Ed. 2007, 46, 7378.

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Caruso, F. ACS Nano 2007, 1, 63.

Chapter 6

Photoinduced Electron Transfer Studies of Cytochrome c

Chapter 6 Photoinduced Electron Transfer Studies of Cytochrome c

6-2

6 Photoinduced Electron Transfer Studies of Cytochrome c

In this Chapter, photoinduced electron transfer studies between Ru(II)-bisterpyridine

complexes and iso-1 cytochrome c (cyt c) as non-covalent mixtures and asymmetric

bioconjugate 8-cyt c were explored. Electron transfer studies were carried out as bulk

solution bioconjugates and membrane encapsulated 8-cyt c. The electron transfer

processes resulting in reduction of the heme of cytochrome c is followed by steady-state

UV-Vis spectroscopy. Electron transfer to the heme was achieved by irradiation using a

465 nm LED light under anaerobic conditions at room temperature in the presence of

sacrificial electron donor, ethylenediaminetetraacetic acid (EDTA). Quantum

efficiencies for bulk and polystyrene140-b-poly(acrylic acid)48 (PS140-b-PAA48)

encapsulated 8-cyt c, were estimated to be 5.9±1.5 10-4% and 1.1±0.3 × 10-3%,

respectively.

In addition, light-induced electron transfer studies between a non-covalent

mixture of Ir(III)-bisterpyridine complex and horse heart cytochrome c were studied to

mimic nitrite reductase behaviour. Excitation and catalysis were performed using a

372 nm LED source under anaerobic conditions at room temperature. Nitrite reductase

behaviour was observed in the presence of 100 nm L—

palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) liposomes and detected using

the Griess assay.

6.1 Background

Electron transfer processes in a direct and controlled fashion are critical for all

functioning living organisms.1 The most notable electron transfer reactions in nature are

photosynthesis and respiration which is responsible for the generation of adenosine


6-3

triphosphate (ATP). ATP is used to drive biochemical reactions and to store energy by

converting carbon dioxide into carbohydrates.2

Cytochrome c is a naturally occurring metalloprotein with iron (cytochroe) heme

cofactor and is part of the electron transport chain in photosynthesis and mitochondria

(respiration) where quinones are its natural electron donors.1 Cytochrome c is heavily

studied for electron transfer mechanisms as it is a robust small protein with an

accessible redox centre, highly stable and soluble in aqueous solvents and is well

characterised with crystal structures.

To improve the understanding of electron transfer processes in complex

biological systems, electron transfer models have been developed and have been

experimentally studied using the model protein cytochrome c with notable contributions

by Gray3 and Millet4 over the last three decades. Electron transfer studies involving

cytochrome c has generally been achieved by modification of either wild type or mutant

species with ruthenium bipyridine complexes at different positions via functionalisation

of histidine3a,5, lysine4a,6 or cysteine residues.7 Electron transfer rates of the

bioconjugates can be elucidated using molecular modelling by combining crystal

structure data as shown in Figure 6.1. These studies have been used to measure electron

pathways in proteins, such as electron tunnelling electron transfer velocities are

dependent on the peptide matrix3d as well as distance dependence between donor and

acceptor components.8 Moreover, the electron transfer process of cytochrome c with its

biological redox partners cytochrome c oxidase9, cytochrome c peroxidase4b and

plastocyanin10 have been studied.


6-4

Figure 6.1. X-ray crystal structure of cytochrome c peroxidase:cytochrome c complex. Mutant yeast iso-1 cytochrome c was functionalised, where the histidine residue 39 has been substituted with cysteine and attached to a tris(bipyridyl)ruthenium(II) complex by molecular modelling.7

The Thordarson group have reported light-harvesting bioconjugates capable of electron

transfer based on Ru(II)-bisterpyridine iso-1 cytochrome c bioconjugates.11

Ru(II)-bisterpyridine complexes have shorter fluorescent lifetimes and lower quantum

yields compared to bipyridine complexes and therefore less desirable for room

temperature electron transfer studies.12 However, they are employed in this project due

to ease of preparation due to their achirality and ease of 4’-functionalisation.

6.2 Room Temperature Photoinduced Electron Transfer Studies

The room temperature light-activated electron transfer between ruthenium bipyridine

complexes (donor) and variants of cytochrome c (acceptor) using sacrificial electron


6-5

donors such as aniline13 and ethylenediaminetetraacetic acid (EDTA)4b,14 have been well

established in the literature. Peterson et al.11a in the Thordarson group have previously

demonstrated room temperature photoinduced electron transfer in Ru(II)-bisterpyridine

cytochrome c bioconjugates using a short and long chain spacer. Photoreduction was

monitored using steady state UV-Vis spectroscopy and low temperature time-resolved

fluorescence lifetime measurements. It was found that long chain spacers resulted in

optimum electron transfer as the use of short chain spacers caused inactivation of the

protein.

Based on the findings of Peterson et al.11a, photoreduction studies of the novel

long chain spacer bioconjugate 8-cyt c and non-covalent mixtures with reference

complex 6 and iso-1 cytochrome c as shown in Figure 6.2, discussed in Chapter 2 and 3

were used to determine the effect of membrane encapsulation for use as light-activated

photosynthesis-respiratory hybrids (further discussed in Chapter 7).

Figure 6.2. Bioconjugate 8-cyt c and reference compounds [Ru(tpy)2]2+ 6 and iso-1cytochrome c (cyt c) used for room temperature photoreduction studies.


6-6

Previous photoreduction studies by the Thordarson group were based on

literature procedures by Hamachi and co-workers, who measured room temperature

electron transfer in apomyoglobin reconstituted with ruthenium bisterpyridine complex

appended heme as the enzyme cofactor.15 Reported photoexcitation conditions were

performed at room temperature with a high pressure mercury lamp (equipped with

<450 nm filter) in the presence of sacrificial electron donor EDTA under argon

atmosphere. It was found that complete photoreduction was observed after 5 h in the

presence of 20 mM EDTA, pH 6.3 at 25 oC.

In contrast to Hamachi et al.15 or Peterson et al.11b which used either a high

pressure mercury lamp or xenon arc lamp that are broad spectrum light sources,

photoreduction studies in this project were based on well-defined narrow wavelength

LED sources (465 nm). In order to demonstrate light-activated electron transfer in

bioconjugate 8-cyt c, samples were prepared in a specialised small volume cuvette,

degassed and irradiated (2.5 cm) with a constant area at a bioconjugate concentration of

2.3±0.1 M and an equivalent 1:1 non-covalent mixture of complex 6 and iso-1 cyt c in

a 5 mM phosphate buffer, 5 mM EDTA at pH 7.0 in either bulk solution or encapsulated

in the PS140-b-PAA48 membrane. This experiment was also conducted under different

conditions to determine the effect on heme reduction rate, including the presence of

oxygen or the absence of sacrificial electron donor EDTA. The concentrations were

estimated by UV-Vis absorption spectroscopy with molar absorption coefficients: iso-1

cyt c/8-cyt c ( 410 = 97.6 mM-1cm-1)16 and reference complex 6 ( 476 = 17.7 mM-1cm-1).12

It should be noted that concentrations used for photoreduction studies were low to

prevent intermolecular electron transfer in bioconjugate samples.


6-7

The reduction rate of the heme group in cytochrome c bioconjugate 8-cyt c was

monitored using UV-Vis spectroscopy from 350 nm to 650 nm as shown in Figure 6.3

and the degree of reduction was calculated based on absorbance changes at 550 nm

according to Equation 6.1:

100%)Reduction(oxred

oxt

AAAA (6.1)

where At is the absorbance after reduction time t, Aox is the original absorbance of

oxidised cytochrome c or bioconjugate and Ared is the final absorbance after complete

reduction.

Figure 6.3. UV-Vis absorbance spectra showing an increase in 550 nm absorbance band over time corresponding to photoreduction of Ru(II) bioconjugate 8-cyt c(2.3±0.1 M) in 5 mM phosphate buffer, 5 mM ethylenediaminetetraacetic acid (EDTA), pH 7.0. Samples irradiated with 20±2.3 mW/cm2 of 465 nm light.


6-8

Bioconjugate 8-cyt c appeared to be fully reduced in 5 mM phosphate buffer, 5 mM

ethylenediaminetetraacetic acid (EDTA), pH 7.0 under degassed conditions in a period

of 50 minutes as shown in Figure 6.4.

Figure 6.4. Room temperature photoreduction of ruthenium(II)-bisterpyridine based iso-1 cytochrome c (cyt c) samples irradiated with 20±2.3 mW/cm2 of 465 nm light. All sample measurements were made at a concentration of 2.3±0.1 M in 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid (EDTA), pH 7.0. Ru(II)bioconjugate 8-cyt c ( ), 8-cyt c:polymersome ( ), 8-cyt c non-degassed ( ), 8-cyt c no EDTA ( ), non-covalent (1:1) Ru(II) 6:cyt c mixture ( ) and non-covalent (1:1) Ru(II)6:cyt c:polymersome mixture ( ).† Error bars indicate standard deviation.

To interpret changes in the 550 nm absorbance band in terms of quantum yield ( ) for

photoreduction, the rate of photon emission of the 465 nm light source was determined

to be 1.40±0.02 × 1016 photons/s at a distance of 2.5 cm between sample and light

source. In the context of these studies, quantum yield is expressed as the ratio of initial

† Different photoreduction conditions were scaled relative to bulk 8-cyt c experiment as a ratio of molar percentage of reduced protein.


6-9

rate of heme reduction vs. absorbed photons. The number of photons absorbed could be

determined by correcting for the optical density of the Ru(II) complex. A quantum

efficiency ( ) of bioconjugate 8-cyt c was estimated to be 5.9±1.5 10-4% (see

Appendix F). It is most likely that the low quantum efficiency is attributed to the short

lifetimes of Ru(II)-bisterpyridine complex at room temperature and the relatively long

distance between the donor and acceptor.11b,12

The Ru(II) bioconjugate 8-cyt c was analysed via molecular modelling to

estimate the long range electron transfer distance from Ru(II) complex to heme as

shown in Figure 6.5. The donor-acceptor distance was estimated by summation of the

linear bond distance between the heme group and cysteine (CYS102) in the crystal

structure of iso-1 cytochrome c (Fe-S: 11 Å) and the maximum distance between the

ruthenium to thiol (CYS102) in a protein attached complex 8 (Ru-S: 21 Å), resulting in

an estimated maximum distance between ruthenium and heme centre of 32 Å.

Figure 6.5. A model of 8-cyt c generated from the X-ray crystal structure of yeast iso-1cytochrome c (cyt c) attached to the X-ray structure of ruthenium(II)-bisterpyridine complex 8 via the CYS102 residue. Maximum distance between Ru-Fe estimated as

32 Å. Cytochrome c structure derived from the protein data bank file ‘1YCC’.17


6-10

To determine the effect of electron transfer after encapsulation within PS140-b-PAA48

polymersome membranes, the polymer aggregates were characterised by transmission

electron microscopy (TEM) prior to room temperature photoreduction measurements to

ensure that bioconjugate 8-cyt c induced polymersome formation as discussed in

Chapter 5. As shown in Figure 6.6, polymersome aggregates were observed with an

average diameter of 290±132 nm.

Figure 6.6. TEM micrograph of polystyrene140-b-poly(acrylic acid)48 (PS140-b-PAA48)polymersome aggregates in the presence of 14 M Ru(II) bioconjugate 8-cyt c in PBS (25 oC). Average polymersome diameter of 290±132 nm. Scale bar: 200 nm.

As shown in Figure 6.4, encapsulation of Ru(II) 8-cyt c resulted in an increase in initial

rate of reduction of heme and fully reduced within 50 minutes. Encapsulation was

estimated to be double the of bulk solution 8-cyt c photoreduction with a of

1.1±0.3 × 10-3%. The increase in could be explained using semi-classical theory

(Marcus-Hush theory of electron transfer) which describes electron tunnelling in

proteins as shown in Equation 6.1:3d,18


6-11

TkGH

Tkhk

B

o

ADB

ET 4)(exp)4(

22

2

3

(6.1)

where the electron transfer rate (kET) from a donor (D) to an acceptor (A) at fixed

separation and orientation depends on the reaction driving force (- Go), a nuclear

reorganisation parameter ( ) and the electronic-coupling strength representing the

probability that an electron tunnels through the potential barrier between A and D

(HAD). Also, kB is the Boltzmann constant, T is temperature and h is Planck’s constant.

It has been reported that embedding reactants in a membrane (low dielectric medium)

dramatically reduces reorganisational energies ( ), hence, the encapsulation of 8-cyt c

in the polymersome membrane leads to an increase in electron transfer rate (kET).3d

Another possibility leading to increased may be due to the flexible linker used in

complex 8 for attachment to cytochrome c allowing the complex to lie flat on the

protein surface when embedded in the polyelectrolyte membrane as shown in Figure

6.5. This decrease in distance between the donor and acceptor increases the coupling

strength HAD by reducing the potential barrier.1

To determine the effect of covalent linkage of Ru(II)-bisterpyridine donor to

cytochrome c, non-covalent studies using a 1:1 mixture of reference complex 6 and

cytochrome c in bulk solution or encapsulated in membrane were performed in the

presence of EDTA and degassed. Complex 6 was chosen as a control complex

compared to complex 8 as the lack of maleimide functional group prevents reaction

with protein. It was observed in Figure 6.4 that a dramatic decrease in photoreduction

yield resulted in non-covalent mixtures. This indicates that covalent attachment to

protein is essential to ensure proximity between donor and acceptor to increase

photoreduction efficiency. However, the behaviour of increasing was observed after

encapsulation, consistent with covalent bioconjugate 8-cyt c studies. The reduction


6-12

efficiencies and quantum efficiencies of bioconjugate 8-cyt c and non-covalent mixtures

in bulk solution and membrane encapsulation is summarised in Table 6.1.

Table 6.1. Estimated rates of heme reduction, reduction efficiency and quantum efficiency ( ) for the photoinduced reduction of Ru(II)-cytochrome c systems.Samplea Heme reduction

(electrons/s)bEfficiency(%) (electrons/photons)c

(%) (electrons/absorbed photons)d

8-cyt ce 7.9±1.8 109 5.6±0.1 10-5 5.9±1.5 10-4

8-cyt cf 1.5±1.8 1010 1.1±0.3 10-4 1.1±0.3 10-3

6::cyt ce,g 4.6±0.6 109 3.3±0.6 10-5 3.4±0.6 10-4

6::cyt cf,g 5.1±1.3 109 3.6±1.0 10-5 3.8±1.1 10-4

a Sample concentration of 2.3±0.1 M in 5 mM sodium dihydrogen phosphate, 5 mMethylenediaminetetraacetic acid (EDTA), pH 7.0 at 25 oC with 80 L volume. Irradiated with 20±2.3 mW/cm2 of 465 nm light. b Initial rate of heme reduction was estimated using amount of protein reduced in the first 1860 s interval. c Efficiency(%) was determined by dividing initial rate of heme reduction by incident photons (1.40±0.2 1016 photons/s). Incident photons calculated from power ouput over an irradiation area of 1.0 0.3 cm for a 465 nm photon. d Quantum efficiency was determined by correcting for the optical density of the solutions (see Appendix F). e Bulk solution measurement. f Encapsulated measurement in PS140-b-PAA48membrane. g Non-covalent mixture. Errors are standard deviation.

Photoexcitation of Ru(II)-bisterpyridine to the triplet metal-to-ligand charge-transfer

(3MLCT) state is short lived and non-luminescent at room temperature with an

excitated-state lifetime estimated to be 250 ps.19 To investigate if the presence of

oxygen significantly quenches the excited chromophore Ru(II)*, photoreduction studies

were performed without degassing. A quenching of photoreduction by 50% was

observed in non-degassed experiments as shown in Figure 6.4. Additionally, the

presence of sacrificial electron donor, EDTA, is essential as bioconjugate 8-cyt c in

phosphate buffer shows negligible photoreduction (Figure 6.4).

6.2.1 Biological activity using cytochrome c oxidase assay

The biological activity of bioconjugate 8-cyt c in bulk solution and encapsulated in

PS140-b-PAA48 membranes were measured by cytochrome c oxidase after

photoreduction. The cytochrome c oxidase assay is used to demonstrate that the proteins


6-13

remain biologically active after bioconjugation and encapsulation in the presence of

organic co-solvents, acetonitrile and tetrahydrofuran, respectively. Additionally,

unmodified iso-1 cytochrome c was oxidised as a reference. The oxidation of reduced

cytochrome c by cytochrome c oxidase can be followed by UV-Vis absorbance

following the characteristic decrease in the 550 nm absorbance band as shown in Figure

6.7. Biological activity was measured using horse heart cytochrome c oxidase. The

samples were prepared at 2.3 M reduced cytochrome c or bioconjugate 8-cyt c (bulk or

encapsulated) in 5 mM phosphate buffer, 5 mM EDTA, pH 7.0 and oxidation initiated by

addition of a catalytic amount of cytochrome c oxidase (0.25 M).

Figure 6.7. Biological activity of iso-1 cytochrome c ( ), Ru(II) bioconjugate 8-cyt cand ( ) and 8-cyt c:polymersome ( ). Samples are at concentration of 2.3±0.1 M in 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid (EDTA), pH 7.0 and oxidation achieved using a catalytic amount of horse heart cytochrome coxidase (0.25 M). Error bars indicate standard deviation.


6-14

As shown in Figure 6.7, bioconjugates 8-cyt c in bulk solution and encapsulated in

membranes demonstrate comparable biological activity to that of native iso-1

cytochrome c indicating attachment of complex 8 to the CYS102 residue does not

influence cytochrome c oxidase binding.4d Furthermore, the ability of cytochrome c

oxidase to oxidise bioconjugate 8-cyt c after encapsulation also supports the

polymersome model discussed previously in Chapter 5, where bioconjugates are

encapsulated in the coronal block of the PS-b-PAA diblock copolymer.20

6.3 Nitrite Reductase Mimics

Nitric oxide (NO) is a biologically relevant cellular signalling molecule responsible for

mediating biological responses such as hypoxic vasodilation, regulation of gene and

protein expression and cytoprotection after ischemia-reperfusion.21 Nitrite anions (NO2-

) comprise the largest vascular storage pool of nitric oxide under physiological

conditions and regulates nitric oxide levels via reduction reactions with

heme-containing proteins.22 It was originally proposed that nitrite was biologically

inert23, however, it has been demonstrated recently that nitrite is an intrinsic vasodilator

and signalling molecule at physiological concentrations in vivo.24 Similar to nitric oxide,

nitrite has been shown to inhibit cytotoxicity and apoptosis after ischemia-reperfusion

injury of the heart, liver and brain.25


6-15

Two distinct types of nitrite reductases are capable of catalysing the reduction of

nitrite anions into nitric oxide, heme cd1 containing chromophores and copper based

enzymes.26 The reaction catalysed by both types of enzymes can be represented by the

simple Equation 6.2:

OHNO2HeNO 22 (6.2)

It should be noted however, that other nitrite reductases reduce nitrite to ammonia such

as cytochrome c nitrite reductase (Escherichia coli) which catalyses the six electron

reduction of nitrite to ammonia.27

One of the most common enzymes responsible for nitric oxide synthesis in

biological systems is nitric oxide synthase (EC 1.14.13.39). Nitric oxide synthase is a

complex enzymatic system which acts on molecular oxygen, arginine and reduced

nicotinamide adenine dinucleotide phosphate (NADPH) to produce nitric oxide,

citrulline and NADP+ as shown in Figure 6.8.28 Five additional cofactors including

flavin mononucleotide (FMN), flavin adenine dinucleotide (FAD), heme, calmodulin

and tetrahydrobiopterin as well as two divalent cations (calcium and heme iron) are

required for catalytic function.

It has been demonstrated that heme containing proteins are capable of additional

roles as nitrite reductase mimics including hemoglobin24, myoglobin29, xanthine

oxidoreductase30, cytochrome c21 and cytochrome c oxidase31 under anoxic conditions.

In this project, nitrite reductase activity of cytochrome c was induced by

light-activation.


6-16

Figure 6.8. Schematic of nitric oxide synthase (EC 1.14.13.39). (a) Representation showing nitric oxide synthesis by acting on molecular oxygen, arginine and reduced nicotinamide adenine dinucleotide phosphate (NADPH) to produce nitric oxide, citrulline and NADP+.32 (b) Solid ribbon representation of human inducible nitric oxide synthase with heme group indicated as stick representations. The structure was derived from the protein data bank file ‘1NSI’.33

6.3.1 Photoinduced nitrite reductase activity of cytochrome c

Basu et al.21 have shown that chemically reduced cytochrome c is capable of reducing

nitrite anions under anaerobic conditions. In order for heme proteins to react with

nitrite, a pentacoordinate state is required. Cytochrome c is a six-coordinate heme iron

which hinder reactions which nitrite, however, in the presence of anionic phospholipids

the weakening or breaking of the iron-methionine bond can occur resulting in a


6-17

pentacoordinate state to activate nitrite reductase activity.34 In addition, the

iron-methionine bond in cytochrome c has been proposed to play a role in apoptosis.35

In this study, an iridium(III)-bistepyridine complex 18 was used to photo-reduce

cytochrome c in the presence of POPC liposomes using a UV LED light source

(372 nm) as shown in Scheme 6.1.

Scheme 6.1. Photoinduced nitrite reductase activity of horse heart cytochrome c (cyt c)using complex 18 irradiated with UV light (372 nm) to reduce cyt c under anaerobic conditions in the presence of L—palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) liposomes which induces pentacoordination of heme state. Solid ribbon representation of horse heart cytochrome c with heme group, MET80 and HIS80 indicated as stick representations. The structure was derived from the protein data bank file ‘1HRC’.36

Photoreduction studies inducing nitrite reductase activity were performed using a 1:1

non-covalent mixture between Ir(III) complex 18 and cytochrome c as work in the

Thordarson group have previously shown that photoreduction efficiency is similar to the

bioconjugate equivalent unlike its ruthenium(II) counterpart.37 Due to experimental

limitations preventing large scale synthesis of Ru(II) bioconjugate 8-cyt c, photoinduced


6-18

nitrite reductase activity studies could not be performed. Additionally, horse heart

cytochrome c was used in this study rather than yeast iso-1 cytochrome c due to its low

cost and does not require further purification.

Nitrite reductase measurements were conducted using conditions adapted from

Basu and coworkers.21 Samples were prepared in plastic eppendorf vials contained in a

vacuum flask and irradiated (2.0 cm) using a 1:1 non-covalent mixture of horse heart

cyt c and complex 18 at a concentration of 400 M, 10 M sodium nitrite, 3 mg/mL

POPC liposomes (4 mM) in a 25 mM phosphate buffer, 5 mM EDTA at pH 5.4 under

anoxic conditions. The concentration of horse heart cytochrome c was estimated by

UV-Vis absorption spectroscopy with molar absorption coefficient

410 = 106.1 mM-1cm-1.38 After irradiation for 3 h followed by incubation for a further

14 h, the final nitrite concentration was determined via the Griess assay as shown in

Scheme 6.2 which converts nitrite into a deep purple azo compound and monitored at

540 nm using UV-Vis absorbance to estimate nitric oxide conversion by difference (see

Appendix C).39

Scheme 6.2. Detection of nitrite via the Griess assay by converting nitrite into a deep purple azo compound and concentration measured spectroscopically at an absorbance of 540 nm.39


6-19

It should be noted that in order to prevent spectroscopic interference by cytochrome c,

the solution was centrifuged using a 3 000 molecular weight cut-off concentrator to

remove protein and filtrate was analysed using the Griess assay. Additionally, studies

were performed at pH 5.4 as Equation 6.2 predicts faster nitrite reduction under acidic

conditions.

Irradiation with 372 nm light of the non-covalent mixture of Ir(III) 18 and

cytochrome c in the presence of POPC liposomes at an initial concentration of 10 M

sodium nitrite under anoxic conditions resulted in photo-activated nitrite reductase

activity leading to an 82% conversion of nitrite anions into nitric oxide (1.8±0.2 M

nitrite remaining) as shown in Figure 6.9. It should be noted that although the liposomes

induce a pentacoordinate heme, unconverted nitrite was present when a forty-fold

excess of cytochrome c relative to nitrite was used due to auto-inhibition of

cytochrome c caused by binding of nitric oxide preventing further reactions with nitrite

and there is always substantial hexacoordinate cytochrome c remaining.21

To eliminate the possibility of interference during the centrifugation step such as

nitrite-membrane binding, a nitrite only control was studied and no effect on nitrite

concentration was observed after centrifugation (NO2- = 10.5±1.0 M). Additionally,

chemically reduced cytochrome c (no liposomes) exhibited neglible nitrite reductase

behaviour as no nitrite was converted during the time-scale of the study within

experimental error, NO2- = 9.3±1.3 M. Nitrite reductase activity was observed only

after reduced cytochrome c was incubated in the presence of POPC liposomes with

50% NO conversion (4.9±1.6 M nitrite remaining). Reduced cytochrome c in the

presence of liposomes is essential to induce nitrite reductase activity as the

cytochrome c (oxidised):liposome mixture exhibited no observable nitrite conversion

Pre-reduced chemically with sodium dithionite, desalted and dialysed against water for 2 h.


6-20

(NO2- = 11.5±1.7 M) as shown in Figure 6.9. This also indicates that zwitterionic

phospholipid liposomes can induce pentacoordination in addition to anionic

phospholipids.34

Figure 6.9. Room temperature photoinduced nitrite reductase activity of horse heart cytochrome c (cyt c) under an anaerobic environment using a 1:1 non-covalent mixture of cyt c and Ir(III) complex 18 (400 M), 3 mg/mL 100 nm POPC liposomes (4 mM), sodium nitrite (10 M) and control conditions. Irradiated with a 0.04±0.02 mW/cm2 UV light source (372 nm) at a distance of 2.0 cm for 3 h and incubated for a further 13 hand 45 min. All sample measurements were made in 25 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid (EDTA), pH 5.4. Error bars arestandard deviation. Pre-reduced chemically with sodium dithionite, desalted and dialysed for 2 h.


6-21

To confirm that nitrite reductase activity was photoinduced, a cytochrome c

(oxidised):liposome mixture was irradiated with 372 nm in the absence of Ir(III) 18 and

no observable nitrite reduction was detected (NO2- = 9.5±0.3 M). Furthermore, a dark

control sample composed of Ir(III) 18:cytochrome c (oxidised):liposomes was

investigated, however, approximately 57% NO conversion was observed (4.3±0.8 M

nitrite remaining) as shown in Figure 6.9. This is attributed to sample exposure to

ambient light during sample preparation resulting in photoreduction of cytochrome c. It

has been shown in the Thordarson group that Ir(III)-bisterpyridine complexes are

extremely effective in heme photoreduction even in aerobic conditions in contrast to

Ru(II) complexes allowing cytochrome c to be fully reduced in under 10 min.37 Also,

the long-lived photoreduction EDTA by-products (amine radicals) are capable of

continued protein reduction once transferred to a dark environment.


In this Chapter, the photophysical properties of a Ru(II)-bistepyridine cytochrome c

bioconjugate 8-cyt c was investigated. Using steady-state UV-Vis absorption

spectroscopy studies, the photoreduction of 8-cyt c could be followed by monitoring the

increase in the 550 nm absorption band corresponding to heme reduction. It was found

that Ru(II) 8-cyt c could be fully reduced in 50 min via irradiation with a 465 nm LED

light source. The quantum efficiency ( ) of bioconjugate 8-cyt c in bulk solution was

estimated to be 5.9±1.5 10-4%. In order to determine the effect of membrane

encapsulation for use as a component in a semi-synthetic electron transport chain,

bioconjugate 8-cyt c was encapsulated in diblock copolymer

polystyrene140-b-poly(acrylic acid)48 (PS140-b-PAA48) as discussed in Chapter 5.

Polymersome formation was confirmed by transmission electron microscopy with an


6-22

average diameter of 290±132 nm. The encapsulated conjugate 8-cyt c, exhibited a

two-fold enhancement of the rate of heme reduction with an estimated of

1.1±0.3 × 10-3%. Molecular modelling of 8-cyt c showed that electron transfer is

occurring over a maximum distance between the ruthenium and heme centre of 32 Å.

Expanding on photoreduction of cytochrome c, electron transfer studies between

a non-covalent mixture of Ir(III)-bisterpyridine complex 18 and horse heart

cytochrome c using a 372 nm UV light source was investigated to induce nitrite

reductase mimicry behaviour. It was shown that in the presence of zwitterionic

phospholipid (POPC) liposomes (100 nm), a pentacoordinate heme state could be

induced allowing reactions with nitrite anions to form nitric oxide. The photo-activated

nitrite reductase activity of cytochrome c exhibited an 82% conversion of nitrite anions

into nitric oxide after irradiation under anoxic conditions in an acidic phosphate

buffered environment (pH 5.4). Due to experimental limitations, the exact irradiation

power could not be determined due scattering and absorption by the vacuum flask and

nitrite levels could only measured at a single time point. For future work, a custom

experimental set-up could be developed to allow controlled sample irradiation under an

anaerobic environment and an amperometric nitric oxide probe could be incorporated to

allow kinetic measurements of nitric oxide formation.

6.5 References

(1) Cordes, M.; Giese, B. Chem. Soc. Rev. 2009, 38, 892.(2) Purves, W. K.; Sadava, D.; Orians, G. H.; Heller, C. H. Life: The Science of

Biology; 7th ed.; Sinauer Associates and W. H. Freeman: U.S.A., 2003.(3) (a) Winkler, J. R.; Nocera, D. G.; Yocom, K. M.; Bordignon, E.; Gray, H. B. J.

Am. Chem. Soc. 1982, 104, 5798. (b) Meade, T. J.; Gray, H. B.; Winkler, J. R. J. Am. Chem. Soc. 1989, 111, 4353. (c) Mayo, S.; Ellis, W.; Crutchley, R.; Gray, H. Science 1986, 233, 948. (d) Gray, H. B.; Winkler, J. R. Annu. Rev. Biochem.1996, 65, 537.

(4) (a) Pan, L. P.; Durham, B.; Wolinska, J.; Millett, F. Biochemistry 1988, 27,7180. (b) Geren, L.; Hahm, S.; Durham, B.; Millett, F. Biochemistry 1991, 30,


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9450. (c) Millett, F.; Durham, B. Methods Enzymol. 2009, 456, 95. (d) Geren, L.; Durham, B.; Millett, F. Methods Enzymol. 2009, 456, 507.

(5) Yang, X.-J.; Drepper, F.; Wu, B.; Sun, W.-H.; Haehnel, W.; Janiak, C. Dalton Trans. 2005, 256.

(6) Hahm, S.; Durham, B.; Millett, F. Biochemistry 1992, 31, 3472.(7) Wang, K.; Mei, H.; Geren, L.; Miller, M. A.; Saunders, A.; Wang, X.; Waldner,

J. L.; Pielak, G. J.; Durham, B.; Millett, F. Biochemistry 1996, 35, 15107.(8) Gray, H. B.; Winkler, J. R. Q. Rev. Biophys. 2003, 36, 341.(9) Geren, L. M.; Beasley, J. R.; Fine, B. R.; Saunders, A. J.; Hibdon, S.; Pielak, G.

J.; Durham, B.; Millett, F. J. Biol. Chem. 1995, 270, 2466.(10) Pan, L. P.; Frame, M.; Durham, B.; Davis, D.; Millett, F. Biochemistry 1990, 29,

3231.(11) (a) Peterson, J. R.; Smith, T. A.; Thordarson, P. Chem. Commun. 2007, 1899. (b)

Peterson, J. R.; Smith, T. A.; Thordarson, P. Org. Biomol. Chem. 2010, 8, 151.(12) Sauvage, J. P.; Collin, J. P.; Chambron, J. C.; Guillerez, S.; Coudret, C.; Balzani,

V.; Barigelletti, F.; De Cola, L.; Flamigni, L. Chem. Rev. 1994, 94, 993.(13) Nilsson, T. Proc. Natl. Acad. Soc. U. S. A. 1992, 89, 6497.(14) Mandal, K.; Hoffman, M. Z. J. Phys. Chem. 1984, 88, 185.(15) Hamachi, I.; Matsugi, T.; Tanaka, S.; Shinkai, S. Bull. Chem. Soc. Jpn. 1996, 69,

1657.(16) Peterson, J. R.; Thordarson, P. Chiang Mai J. Sci. 2009, 26, 236.(17) Louie, G. V.; Brayer, G. D. J. Mol. Biol. 1990, 214, 527.(18) Marcus, R. A.; Sutin, N. Biochim. Biophys. Acta 1985, 811, 265.(19) Winkler, J. R.; Netzel, T. L.; Creutz, C.; Sutin, N. J. Am. Chem. Soc. 1987, 109,

2381.(20) Hvasanov, D.; Wiedenmann, J.; Braet, F.; Thordarson, P. Chem. Commun. 2011,

47, 6314.(21) Basu, S.; Azarova, N. A.; Font, M. D.; King, S. B.; Hogg, N.; Gladwin, M. T.;

Shiva, S.; Kim-Shapiro, D. B. J. Biol. Chem. 2008, 283, 32590.(22) Dejam, A.; Hunter, C. J.; Tremonti, C.; Pluta, R. M.; Hon, Y. Y.; Grimes, G.;

Partovi, K.; Pelletier, M. M.; Oldfield, E. H.; Cannon, R. O.; Schechter, A. N.; Gladwin, M. T. Circulation 2007, 116, 1821.

(23) Lauer, T.; Preik, M.; Rassaf, T.; Strauer, B. E.; Deussen, A.; Feelisch, M.; Kelm, M. Proc. Natl. Acad. Soc. U. S. A. 2001, 98, 12814.

(24) Cosby, K.; Partovi, K. S.; Crawford, J. H.; Patel, R. P.; Reiter, C. D.; Martyr, S.; Yang, B. K.; Waclawiw, M. A.; Zalos, G.; Xu, X.; Huang, K. T.; Shields, H.; Kim-Shapiro, D. B.; Schechter, A. N.; Cannon, R. O.; Gladwin, M. T. Nat. Med.2003, 9, 1498.

(25) (a) Webb, A.; Bond, R.; McLean, P.; Uppal, R.; Benjamin, N.; Ahluwalia, A. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 13683. (b) Gladwin, M. T. Nat. Chem. Biol. 2005, 1, 245. (c) Duranski, M. R.; Greer, J. J. M.; Dejam, A.; Jaganmohan, S.; Hogg, N.; Langston, W.; Patel, R. P.; Yet, S.-F.; Wang, X.; Kevil, C. G.; Gladwin, M. T.; Lefer, D. J. J. Clin. Invest. 2005, 115, 1232. (d) Jung, K.-H.; Chu, K.; Ko, S.-Y.; Lee, S.-T.; Sinn, D.-I.; Park, D.-K.; Kim, J.-M.; Song, E.-C.; Kim, M.; Roh, J.-K. Stroke 2006, 37, 2744.

(26) Averill, B. A. Chem. Rev. 1996, 96, 2951.(27) Clarke, T. A.; Mills, P. C.; Poock, S. R.; Butt, J. N.; Cheesman, M. R.; Cole, J.

A.; Hinton, J. C. D.; Hemmings, A. M.; Kemp, G.; Söderberg, C. A. G.; Spiro, S.; Van Wonderen, J.; Richardson, D. J. Methods Enzymol. 2008, 437, 63.

(28) Liu, Q.; Gross, S. S. Methods Enzymol. 1996, 268, 311.


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(29) Shiva, S.; Huang, Z.; Grubina, R.; Sun, J.; Ringwood, L. A.; MacArthur, P. H.; Xu, X.; Murphy, E.; Darley-Usmar, V. M.; Gladwin, M. T. Circ. Res. 2007, 100,654.

(30) Li, H.; Samouilov, A.; Liu, X.; Zweier, J. L. J. Biol. Chem. 2004, 279, 16939.(31) Castello, P. R.; Woo, D. K.; Ball, K.; Wojcik, J.; Liu, L.; Poyton, R. O. Proc.

Natl. Acad. Soc. U. S. A. 2008, 105, 8203.(32) Cayman Chemical Company: Ann Arbor, MI, 2009.(33) Li, H.; Raman, C. S.; Glaser, C. B.; Blasko, E.; Young, T. A.; Parkinson, J. F.;

Whitlow, M.; Poulos, T. L. J. Biol. Chem. 1999, 274, 21276.(34) Tuominen, E. K. J.; Wallace, C. J. A.; Kinnunen, P. K. J. J. Biol. Chem. 2002,

277, 8822.(35) Kagan, V. E.; Tyurin, V. A.; Jiang, J.; Tyurina, Y. Y.; Ritov, V. B.; Amoscato,

A. A.; Osipov, A. N.; Belikova, N. A.; Kapralov, A. A.; Kini, V.; Vlasova, I. I.; Zhao, Q.; Zou, M.; Di, P.; Svistunenko, D. A.; Kurnikov, I. V.; Borisenko, G. G. Nat. Chem. Biol. 2005, 1, 223.

(36) Bushnell, G. W.; Louie, G. V.; Brayer, G. D. J. Mol. Biol. 1990, 214, 585.(37) Goldstein, D. C. PhD Thesis, The University of New South Wales, 2011.(38) Margoliash, E.; Frohwirt, N. Biochem. J. 1959, 71, 570.(39) Titheradge, M. A. In Methods in Molecular Biology: Nitric Oxide Protocols;

Titheradge, M. A., Ed.; Humana Press Inc.: Totowa, NJ, 1998; Vol. 100.

Chapter 7

Self-Assembled Light-Driven Proton Pumping Studies

Chapter 7 Self-Assembled Light-Driven Proton Pumping Studies

7-2

7 Self-Assembled Light-Driven Proton Pumping Studies

In this Chapter, the construction of a self-assembled artificial hybrid

photosynthetic-respiratory electron transport chain for light-induced proton pumping is

described. The synthetic hybrid photosynthetic-respiratory chain is based on

ruthenium(II)-bistepyridine linked to cytochrome c (8-cyt c) coupled with the terminal

electron acceptor of the mitochondrial electron-transport chain, cytochrome c oxidase

(CcOx) and membrane encapsulated in diblock copolymer polystyrene140-b-poly(acrylic

acid)48 based on the work discussed in previous chapters of this Thesis.

Upon irradiation at an initial pH of 7.2, a proton pumping rate of 3.3 ×

103 H+/s across the polymer bilayer generating a gradient up to pH 0.2 was observed

by steady-state fluorescence spectroscopy studies using fluorescent pH dye, 8-hydroxy-

1,3,6-pyrenetrisulfonate (HPTS). Reconstituted cytochrome c oxidase exhibited a

greater than 50% native mitochondrial orientation in the polymersome membrane. This

chapter demonstrates that the reconstituted photoactive hybrid electron-proton chain

generates a proton gradient which can store chemical energy and is a step towards the

development of a model artificial protocell.

7.1 Background

A high order of organisation of assembly of discrete components to allow direct and

controlled electron transfer processes are critical for all functioning living organisms.1

Electron transfer across the photosynthetic electron transport chain is an ideal example

of this phenomenon which is involved in the production of adenosine triphosphate

(ATP) to drive biological processes.2

Photosynthesis in plants is one of the most important reactions in maintaining

life by providing the source of oxygen and biomaterial on Earth. It converts light energy


7-3

into stored chemical energy by driving the reaction between carbon dioxide and water to

produce sugar and the release of oxygen.3 In plants, photosynthesis occurs in the

organelle chloroplast, which comprises the electron transport chain in the thylakoid

membrane. Upon absorption of light energy (photons), charge separation events are

initiated by reaction centres allowing rapid electron transfer through cofactor chains

which stabilises the charge separation and prevents recombination. Light induced

charge separation generates a strong oxidant (water oxidising complex) in photosystem

II (PSII) which extracts electrons from water producing oxygen and subsequent electron

transfer to photosystem I (PSI) generates a strong reducing agent (oxidised chlorophyll

P700) catalysing nicotinamide adenine dinucleotide phosphate (NADP+) reduction as

shown in Figure 7.1.4 The electron flow between PSII and PSI through the electron

transport chain is coupled to proton translocation in a redox loop mechanism to produce

a proton gradient which generates ATP by ATP synthase.2,4

Figure 7.1. Light-induced photosynthetic electron transport chain catalysing oxidation of water generating oxygen and the subsequent proton gradient produces adenosine triphosphate (ATP) to drive biological processes.3


7-4

The respiratory electron transport chain of the mitochondria is analogous to

photosynthesis where electrons are translocated through protein complexes that span

cell membranes consisting of chains of organic and inorganic cofactors that act as

stepping stones for electron transfer, which is coupled to proton translocation. The

proton gradient is used to drive ATP production by ATP synthase.5

In the process of respiration, the proton gradient is contributed by the terminal

oxidase of the respiratory chain of mitochondria, cytochrome c oxidase (complex IV,

EC 1.9.3.1). Since the discovery by Wikström in 1977 that cytochrome c oxidase redox

activity is coupled to membrane translocation of protons, it has become one of the most

well characterised and studied integral membrane proteins.6 Cytochrome c oxidase

catalyses the reduction of oxygen to form water consuming four electrons (from

cytochrome c) and protons while concomitantly translocating four protons across a

membrane as shown in Figure 7.2.7 It is a 200 kDa Y-shaped multisubunit enzyme

where the two arms of the Y span the inner membrane and the stalk extends into the

cytoplasmic side allowing binding to cytochrome c.8 In this project, work is primarily

focused on cytochrome c oxidase, however, it should be noted that complex I (NADH

dehydrogenase) and complex III (cytochrome bc1 complex) also contribute to the

generation of a proton gradient across the inner mitochondrial membrane during

respiration.2


7-5

Figure 7.2. Solid ribbon representation of bovine heart cytochrome c oxidase (complex IV, EC 1.9.3.1). The structure was derived from the protein data bank file ‘2OCC’.9

Natural electron transport systems (photosynthetic or respiratory) are highly

sophisticated and require control of components in space, energy and time while being

organised in a supramolecular system.1 Moreover, the storage of chemical energy in a

compartmentalised space is also one of the key requisites for any model artificial

protocell; the other key attributes being the ability to self-replicate the vesicle

compartment and genetic information.10 Scientists have attempted to develop artificial

photosynthetic systems to produce chemical energy as an electrochemical proton

gradient across a membrane in a confined space using self-assembly.

One approach to construct a photosynthetic hybrid is to use a light-harvesting

photosensitiser linked to a donor and acceptor capable of generating a redox potential

gradient across a membrane. This gradient can then translocate protons across a


7-6

phospholipid membrane vesicle (liposome) using a quinone relay as first reported by

Steinberg-Yfach et al.11 Additionally, Bhosale et al.12 have developed an elegant mimic

using aryl diimides which undergo - stacking, and upon photoexcitation achieves

directional electron transfer in liposomes coupled with proton transfer due to quinone

reduction which acts as an electron acceptor. In both these cases, the proton gradient is

generated passively by the virtue of quinone-redox chemistry that either shuttles protons

into or depletes protons from the interior of the vesicle. Finally, another approach

reported to produce a photosynthetic mimic was to encapsulate an active light-driven

transmembrane proton pump, bacteriorhodopsin in a polymer vesicle (polymersome) to

generate a proton gradient.13 Here, the proton pumping action is based on the natural

ability of bacteriorhodopsin to unidirectionally gate protons through an ion channel

upon light-induced transient conformational changes in the protein chromophore.

Attempts have been made by Meier et al.14 as well as Rowan and van Hest et

al.15 to form proteo-polymersomes by reconstitution of channel proteins or catalytic

enzymes to form a catalytic cascade pathway, respectively. However, the reconstitution

of enzymes in polymersome membranes to form a semi-synthetic electron transport

chain has never been attempted.

7.2 Photosynthetic-Respiratory Hybrid System

The construction of a chloroplast mimic that links membrane-bound photoinduced

electron transfer across an enzyme cascade with concomitant proton translocation to

generate an electrochemical potential ( ) based on synthetic polymersomes is

described. In order to develop the chloroplast hybrid using a synthetic polymer

membrane, ruthenium(II)-bisterpyridine complex 8 was selected as the light-harvesting

component and linked to iso-1 cytochrome c (8-cyt c) which functions as an electron


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transport redox protein as shown in Figure 7.3. Ru(II) 8-cyt c was prepared by

covalently linking the synthetic maleimide functionalised ruthenium(II)-bisterpyridine

complex 8 with CYS102 of cytochrome c via Michael addition as discussed in Chapter

3.16

Figure 7.3. Bioconjugate 8-cyt c used as the light-harvesting electron transport redox protein.

To complete the photosynthesis-electron transport chain hybrid, mitochondrial

cytochrome c oxidase was selected as the natural acceptor which mediates electron

transfer across the enzyme cascade and simultaneously vectorially translocates protons

upon reduction.17 The primitive chloroplast is a modified synthetic hybrid of the

electron transport chain and is capable of converting photon energy into a by direct

electron transport from the light-harvesting centre of the hybrid 8-cyt c to the active

proton pump cytochrome c oxidase as shown in Figure 7.4.


7-8

Figure 7.4. Schematic representation of light-harvesting enzyme cascade generating a proton gradient upon irradiation. Encapsulation of 8-cyt c within hydrophilic poly(acrylic acid) region of diblock copolymer membrane PS140-b-PAA48 coupled electrostatically with transmembrane cytochrome c oxidase in the hydrophobic polystyrene block. Irradiation in the presence of sacrificial electron donor ethylenediaminetetraacetic acid (EDTA) leads to generation of pH which is detected by fluorescent pH dye, 8-hydroxy-1,3,6-pyrenetrisulfonate (HPTS). Proton translocation from the interior negative (N-) side to the positive (P-) side illustrated.

The resulting semi-synthetic enzyme cascade was membrane encapsulated in

polymersomes based on the polystyrene140-b-poly(acrylic acid)48 (PS140-b-PAA48)

diblock copolymer. Polymeric vesicles were prepared in this study using the ‘kinetic

trapping’ method previously discussed in Chapter 5 by injecting a tetrahydrofuran

solution of dissolved PS140-b-PAA48 into a solution of bioconjugate 8-cyt c and

cytochrome c oxidase. It was found that this preparation method is highly robust against

changes in salt concentration and pH and that the predominant factor controlling


7-9

formation of polymersomes based on PS140-b-PAA48 is the addition of positively

charged proteins which induce vesicle formation (Chapter 5).18 This achieves

concomitant encapsulation of both enzymes within the membrane. Additionally, Ru(II)

8-cyt c and cytochrome c oxidase were selectively encapsulated in separate membrane

domains, with the positively charged 8-cyt c entrapped in the hydrophilic poly(acrylic

acid) block and lipophilic cytochrome c oxidase in the hydrophobic polystyrene block.

7.2.1 Polymersome morphologies and membrane reconstitution

To eliminate the possibility that the induced pH gradient was a result of morphological

changes of polyelectrolyte PS140-b-PAA48 during irradiation, transmission electron

microscopy studies (TEM) were performed on enzyme cascade (8-cyt c:CcOx)

polymersome membrane reconstituted samples upon irradiation with 465 nm light over

56 min. TEM studies demonstrated that there were no significant changes in

polymersome morphologies and diameters are identical within experimental

uncertainty, indicating that the pH gradient is a result of proton translocation (inducing

a pH gradient at an initial pH of 7.2) and excluding the possibility that the pH gradient

is a result of changing morphologies of polymer aggregates as shown in Figure 7.5a-d.

These polymersome morphologies have been previously confirmed by cryo-TEM in

Chapter 518 and vesicles are defined as morphologies displaying a distinct contrast

within individual particles indicating the presence of bilayer and cavity formation.


7-10

Figure 7.5. Structural characterisation of hybrid proteo-polymersomes (8-cyt c:CcOx:HPTS) by transmission electron microscopy. (a) Low magnification electron micrograph of proteo-polymersomes at time 0 min and initial pH 7.2. Scale bar: 400 nm. (b) Electron micrograph of proteo-polymersomes at time 0 min and initial pH 7.2. (367±185 nm). Scale bar: 200 nm. (c) Electron micrograph after irradiation for 24 min using 465 nm light. (273±83 nm). Scale bar: 200 nm. (d) Electron micrograph after irradiation for 56 min using 465 nm light to a final light induced pH gradient of

pH 7.4. (291±74 nm). Scale bar: 200 nm. All sample measurements were made in a 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid buffer at an initial pH 7.2. Errors are standard deviation.

In order to confirm reconstitution of cytochrome c oxidase within the hydrophobic

membrane of PS140-b-PAA48, confocal laser scanning microscopy was performed on

reconstituted fluorescently labelled cytochrome c oxidase excited using a 488 nm laser

as shown in Figure 7.6a-b.


7-11

Figure 7.6. Structural characterisation of hybrid proteo-polymersomes (8-cyt c:CcOx:HPTS) by confocal laser scanning microscopy of fluorescently labelled cytochrome c oxidase encapsulated in PS140-b-PAA48 polymersome membrane.(a) Transmission light micrograph of fluorescently labelled cytochrome c oxidaseencapsulated proteo-polymersomes. (b) Confocal laser scanning micrograph excited at 488 nm (contrast enhanced). Scale bars: 10 m and inset 1 m. All sample measurements were made in a 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid buffer at an initial pH 7.2.

Polymersomes containing cytochrome c:cytochrome c oxidase-Dye could be visualised

optically via fluorescence due to aggregate structures with diameters up to 1 m as

shown in Figure 7.7. This indicates that cytochrome c oxidase is successfully

reconstituted in the polymer membrane rather than remaining as a homogenous mixture

in the bulk or internal polymersome aqueous phase due to the localised fluorescence in

the polymersome aggregates.18


7-12

Figure 7.7. Representative histogram of PS140-b-PAA48 polymersome diameters for 7.5 M cytochrome c:0.75 M cytochrome c oxidase in PBS at 25 oC.

7.2.2 Photoinduced pH gradient

Water soluble fluorescent pH dye, 8-hydroxy-1,3,6-pyrenetrisulfonate11-13,19 (HPTS,

Figure 7.8a) was entrapped in the internal volume of polymersomes to monitor pH

correlating to proton translocation from the negative (N-) to the postive (P-) side of the

membrane. Figure 7.8b shows the pH dependent changes in the fluorescence properties.


7-13

Figure 7.8. pH dependent photophysical properties of 8-hydroxy-1,3,6-pyrenetrisulfonate (HPTS). (a) Chemical structure of HPTS. (b) pH titration curve of HPTS fluorescence intensity emission ( ex = 460 nm, em = 510and encapsulated pyranine ( ) in PS140-b-PAA48 in 20 mM sodium dihydrogen phosphate. Bulk pKa = 7.56 and encapsulated pKa = 7.62. Inset: excitation spectra ( em = 510 nm) of internally encapsulated HPTS in dialysed polymersomes and incubated at pH 5.1 (black solid line) and pH 9.2 (black dashed line). Error bars are standard deviation.

Internal pH changes in hybrid proteo-polymersomes were determined based on the

fluorescence intensity of enclosed HPTS as well as the ratiometric ratio of the two

peaks 405 and 460 nm as shown in Figure 7.9. Irradiation with a 465 nm light source


7-14

results in initiation of electron transport across the 8-cyt c:CcOx electron transport

chain, leading to an overall proton translocation in the presence of sacrificial electron

donor ethylenediaminetetraacetic acid (EDTA).

Figure 7.9. Light-induced transmembrane proton translocation in hybrid proteo-polymersomes. Fluorescence excitation (left, em 510 nm) and emission spectra (right, ex = 460 nm; em = 510 nm) of encapsulated HPTS after irradiation at an initial pH 6.5. The polymersomes were irradiated for 0 ( ), 12 ( ), 24 ( ), 36 ( )and 56 ( ) min with 20.0±2.3 mW/cm2 of 465 nm light. Measurement was made in a 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid buffer, pH 6.5.

Determination of internal pH could be achieved by monitoring fluorescence intensity

( ex = 460 nm; em = 510 nm) and calibrating to a pH titration curve (Figure 7.8b).19

The proteo-polymersome ensemble (Figure 7.4) shows an increase in fluorescence

intensity after irradiation with actinic light at 465 nm over a 1 h period. An increase in

fluorescence intensity corresponds to basification of the internal aqueous compartment


7-15

indicating overall vectorial proton translocation from the N- to P-side. Proton pumping

rate at an initial pH of 7.2 was estimated to be 3.3±1.0 × 103 H+/s (see Appendix F).

Figure 7.10. Light-induced transmembrane proton translocation in hybrid proteo-polymersomes. Relative fluorescence intensity of internal HPTS as a function of irradiation time at an initial pH of 7.2. 8-cyt c:CcOx:polymersome

[H+] = 1.6 × 10-8 M 8-cyt c:CcOx bulk [H+] = 3.8 × 10-9 M ( ), 8-cyt c:polymersome [H+] = 6.8 × 10-9 M ( ), internal EDTA [H+] = 4.3 × 10-9 M( ), cyt c:CcOx:polymersome [H+] = 4.4 × 10-10 M ( ), dark control

[H+] = 1.3 × 10-9 M ( ), polymer micelle [H+] = -1.8 × 10-9 M ( ) and poly-L-lysine induced:polymersome [H+] = -1.1 × 10-9 M (×). Error bars indicate standard deviation. All sample measurements were made in a 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid buffer at a desired pH.

Additionally, in order to exclude artefacts, the excitation spectra as shown in Figure 7.9

displays a ratiometric change of the two maxima peaks at 405 and 460 nm ensuring that

the fluorescence intensity increase is a result of pH change. Based on Figure 7.11, it can

be observed that there is a decrease in the ratiometric intensity of the two peaks

indicating basification of the interior at an initial pH of 6.5 in which the proton pumping


7-16

activity is greater (see Chapter 7.2.4). This correlates to the fluorescence intensity

increase in the emission spectra indicating basification of the interior compartment.

Interestingly, the excitation spectra showed unusual behaviour with concomitant

increase in the excitation peaks as shown in Figure 7.9. It is proposed that this

behaviour is attributed to scattering caused by EDTA by-products which is formed by

photoreduction of oxygen with the Ru(II)-bisterpyridine complex (see Appendix E). It

was found that due to scattering and the small overall pH changes in the hybrid

proteo-polymersome system, the fluorescence intensity emission method was more

reliable and sensitive to pH changes and the proton pumping rates were determined

using this approach which has been previously reported for monitoring internal pH

changes in cells.19-20

Figure 7.11. Intensity ratio of the two peaks (405 and 460 nm) of the excitation spectra of internal HPTS after irradiation at an initial pH 6.5. Error bars are standard deviation. Measurements were made in a 5 mM sodium dihydrogen phosphate, 5 mMethylenediaminetetraacetic acid buffer, pH 6.5.


7-17

To demonstrate that internal pH was a result of proton translocation within the

polymersome, pumping studies consisting of the ensemble components in a bulk

solution were performed without encapsulation. Fluorescence studies indicated a minor

increase in internal pH. However, it can be seen that the change in intensity is

significantly smaller than the encapsulated 8-cyt c:CcOx:polymersome light-harvesting

hybrid. It is proposed that this background change in internal pH may be due to the

electron transfer between sacrificial electron donor EDTA and 8-cyt c producing

reaction intermediates consuming internal protons. Another possibility may be a result

of light-induced or structurally enzyme modified loss of peroxidase-like activity which

leads to radical generation and propagation mechanisms.16,21 Additionally, this

background pH change is similarly observed for the 8-cyt c:polymersome control (i.e.,

no CcOx). A pH of 0.2 units can be observed for the 8-cyt c:CcOx:polymersome

construct, whereas, the control samples generally exhibit a pH <0.07 units.

The block copolymer membranes of the polyelectrolyte system PS140-b-PAA48

are considerably thicker (98±35 nm) than phospholipid bilayers ( 5 nm).22 This

corresponds to a bilayer of PS140-b-PAA48 in the fully extended state. Cytochrome c

oxidase (CcOx) spans 12 nm9 and can be reconstituted to span the polymer membrane

due to the high flexibility and the conformational freedom of the polymer molecules,

allowing the block copolymer to adapt to the dimensions or conformation of the

transmembrane protein without loss of free energy while remaining functional. This

effect has been reported by Meier et al.14a for reconstitution of small channel proteins

(bacterial porin OmpF) within a triblock copolymer. In addition, van Hest et al.23 have

shown that in mixed solvent mixtures of water and tetrahydrofuran, where

tetrahydrofuran is a good solvent and plasticiser of polystyrene (PS), the flexibility of

hydrophobic block (PS) is preserved despite the high glass transition temperature (Tg).


7-18

This property of PS allows the rearrangement of the diblock copolymer to conform to

the dimensions of CcOx as the self-assembly process of polymersomes occurs in the

presence of the transmembrane protein in a co-solvent mixture of water and

tetrahydrofuran over at least 12 h. This is supported by previous work as discussed in

Chapter 5 which showed that PS140-b-PAA48 polymersomes could be induced from

micelle aggregates after addition cytochrome c within 30 min in a THF:PBS (1:6 v/v)

co-solvent mixture at room temperature.18

The above shows that the measured fluorescence intensity changes of

encapsulated HPTS were light-induced effects. Both the dark incubation control and

cyt c:CcOx:polymersome which lacks the light-harvesting chromophore as shown in

Figure 7.10 resulted in neglible intensity change. It should be noted that EDTA

by-products are highly effective at reducing cytochrome c as discussed in Chapter 6. To

minimise background effects caused by EDTA and its by-products, EDTA solution was

incubated in a dark environment for at least 24 h and the hybrid proteo-polymersome

system was incubated in the dark for 1 h prior to dark control measurements. Moreover,

the dark control sample was measured only at time 0 min and 56 min as the excitation

wavelength for HPTS fluorescence can cause photoexcitation of the

Ru(II)-bisterpyridine complex.

Additionally, to demonstrate that the polyelectrolyte PS-b-PAA does not

contribute to the basification of the interior compartment, a polymer only control

(micelle) and a biologically inert positively charged polypeptide (poly-L-lysine) control

was used to induce polymersome formation encapsulating HPTS were chosen to

eliminate the possibility that poly(acrylic acid) was acting as a proton transporter upon

irradiation. Interestingly, over the period of illumination, slight acidification of the

interior was observed in contrast to basification exhibited by the enzyme ensemble


7-19

system. This slight acidification in the absence of the proton pumping assembly may be

attributed to the weak acid nature of poly(acrylic acid).24 This shows that our system

undergoes light-induced vectorial proton translocation.

7.2.3 Orientation of reconstituted cytochrome c oxidase

Determination of the average orientation of reconstituted cytochrome c oxidase can be

deduced based on the relative increase or decrease of HPTS emission intensity. HPTS

undergoes an increase in fluorescence luminescence from acidic to a basic environment.

Fluorescence measurements as shown in Figure 7.9 and Figure 7.11 indicate an increase

in HPTS emission after irradiation with actinic light indicating basification of the

interior volume of the polymersome. This indicates favoured orientation of the average

reconstituted cytochrome c oxidase in PS140-b-PAA48 in the native mitochondrial

configuration (Figure 7.4).2,5 It should be noted that the orientation of the reconstituted

cytochrome c oxidase indicates a bias in the average orientation (>50% in the native

mitochondrial configuration) and not exclusively uniform orientation in the

cytochrome c oxidase population. This indicates that proton pumping from the exterior

into the interior compartment (acidification) can occur simultaneously and the

fluorescence measurements indicate an average proton translocation from the N- to P-

side (basification).

The bias of the average orientation (>50% native orientation) during

reconstitution may be due to the polar region of cytochrome c oxidase on the native

cytoplasmic side of the mitochondrial membrane being significantly larger than the

polar region on the matrix side.25 This may cause the orientation to be biased during the

self-assembly process, whereby the larger radii of curvature of the outer polymersome

(PAA block) accommodates this polar surface. The observation of favoured native


7-20

oriented cytochrome c oxidase was further confirmed based on the internally entrapped

ethylenediaminetetraacetic acid (EDTA) control as shown in Figure 7.10 (no addition of

EDTA to external bulk phase) indicating a reduced pH.

7.2.4 Dependency of proton translocation rates on pH

It was found that the proton pumping rates are sensitive to pH environment as shown in

Figure 7.12. Polymersomes were equilibrated in a phosphate buffer (5 mM) at fixed pH

such that the internal (pHin) and external pH (pHout) were equal. Under an acidified

polymersome environment at pH 6.5, a proton pumping rate of 1.3±0.4 × 104 H+/s was

observed. In contrast, a basified environment at pH 7.9 showed significant inhibition of

proton translocation rate to 8.3±2.5 × 102 H+/s. These results demonstrate that even a

slight change of pH from physiological pH 7.2 to acidic pH 6.5 can lead to a 4-fold

enhancement. This is despite the apparent higher proton “back-pressure” in the more

acidic environment (pH 6.5).26 The rate dependency on pH environment is attributed to

the intrinsic pH-dependent kinetics of cytochrome c oxidase. These results are

consistent with previously reported findings showing that the proton pumping rate is

dependent on pHin which determines the kinetics of internal electron transfer within

cytochrome c oxidase linking proton translocation.27


7-21

Figure 7.12. Dependency of proton translocation rates on pH. Proton pumping rates under equilibrated physiological (pH 7.2), acidic (pH 6.5) and basic (pH 7.9) conditions. Error bars indicate standard deviation. All sample measurements were made in a 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic acid buffer at the desired pH.

7.2.5 Proton pumping quantum efficiencies ( )

To interpret changes in fluorescence intensity in terms of quantum yield ( ) for proton

translocation, the rate of photon emission of the actinic light source was determined to

be 1.40±0.02 × 1016 photons/s at a distance of 2.5 cm between sample and light source.

In the context of these studies, quantum yield is expressed as the ratio of proton

translocation vs. absorbed photons. The number of photons absorbed could be

determined by correcting for the optical density of the Ru(II) complex antenna (see

Appendix F). A for the synthetic photosynthetic-respiratory hybrid system was

estimated to be 1.1±0.3 × 10-9% at pH 7.2. This poor may be a result of poor


7-22

electronic communication between Ru(II) 8-cyt c and cytochrome c oxidase in the

membrane system as the rate limiting step.

Polymersomes offer the advantage of greater stability for weeks to months

compared to liposomes of hours to days due to lower chain mobilities which ‘freeze’ the

structure.28 However, in this hybrid system, this proves to be disadvantageous by

simultaneously reducing the rates of diffusion of both reconstituted cytochrome c

oxidase and 8-cyt c. This reduces the probability of 8-cyt c:CcOx binding. The rate of

replenishing electrons to cytochrome c oxidase across the electron transport chain is

therefore dramatically reduced and dependent on the rate of reduction of 8-cyt c at the

single enzyme level. These findings were also supported as the of intramolecular

electron transfer for 8-cyt c was 1.1±0.3 × 10-3% as discussed in Chapter 6 in the

polymersome membrane environment which is 1 × 106 fold more efficient than proton

pumping . Consequently, the apparent turnover number of cytochrome c oxidase

proton translocation is also significantly inhibited and determined to be

6.2±1.9 × 10-11 H+/CcOx/s as the rate limiting step is directly correlated to the for

proton pumping. In comparison, the turnover number of cytochrome c oxidase in rat

liver mitochondria is 160 H+/CcOx/s.29


In this Chapter, a synthetic hybrid photosynthetic-respiratory system that generates a

proton potential upon photoexcitation has been constructed via self-assembly.

Reconstitution of the hybrid enzyme cascade consisting of bioconjugate 8-cyt c,

cytochrome c oxidase in the polymersome membrane of polyelectrolyte PS140-b-PAA48

was characterised by transmission electron microscopy and confocal laser scanning

microscopy. Irradiation with actinic light converts photon energy into an


7-23

electrochemical potential via electron transfer. Charge transfer occurs across an

artificial electron transport chain within a polymersome membrane coupled with overall

vectorial proton translocation and resulting pH changes were monitored by steady-state

fluorescence spectroscopy using internally entrapped HPTS. From these investigations,

it was also found that cytochrome c oxidase appears to have the same orientation

(average >50% native orientation) in the polymersome membrane encountered in the

mitochondria. This biased orientation is attributed to the alignment of cytochrome c

oxidase with the curvature of the polymersomes. The photosynthetic-respiratory system

demonstrates the construction of an artificial hybrid organelle capable of storing

chemical energy in the form of a proton potential across a membrane with a proton

pumping rate of 3.3 × 103 H+/s at physiological pH 7.2 ( pH 0.2 units) correlating

to a of 1.1±0.3 × 10-9%. Proton translocation rates were investigated using

pH-dependent studies showing that a four-fold increase in proton pumping rate is

observed under acidic conditions of pH 6.5. This synthetic photoactive electron

transport chain capable of storing chemical energy as a proton gradient brings vesicle

and polymersome chemists towards satisfying one of the three requirements10a for an

artificial model protocell, which include (1) catalyst (storing of chemical energy), (2)

self-reproducing vesicles10b and (3) self-reproducing informational substance (genetic

information).10b

For future work, analogous Ir(III)-bisterpyridine based bioconjugates can be used

in contrast to Ru(II) bioconjugates to improve proton pumping rates and due to their

improved photoreduction abilities. Additionally, it is possible to utilise this system and

harness the potential to drive biomimetic processes such as generation of ATP11b,13 and

ATP-linked processes by potentially reconstituting transmembrane ATP synthase.

Alternatively, reagents can be encapsulated within the interior cavity of the


7-24

proteo-polymersome to induce pH sensitive reactions which can be activated upon

photoinduced pH changes.

7.4 References

(1) Sauvage, J. P.; Collin, J. P.; Chambron, J. C.; Guillerez, S.; Coudret, C.; Balzani, V.; Barigelletti, F.; De Cola, L.; Flamigni, L. Chem. Rev. 1994, 94, 993.

(2) Cordes, M.; Giese, B. Chem. Soc. Rev. 2009, 38, 892.(3) Purves, W. K.; Sadava, D.; Orians, G. H.; Heller, C. H. Life: The Science of

Biology; 7th ed.; Sinauer Associates and W. H. Freeman: U.S.A., 2003.(4) Nugent, J. H. A. Eur. J. Biochem. 1996, 237, 519.(5) Rich, P. R. Biochem. Soc. Trans. 2003, 31, 1095.(6) Wikstrom, M. K. F. Nature 1977, 266, 271.(7) (a) Bloch, D.; Belevich, I.; Jasaitis, A.; Ribacka, C.; Puustinen, A.; Verkhovsky,

M. I.; Wikström, M. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 529. (b) Jasaitis, A.; Verkhovsky, M. I.; Morgan, J. E.; Verkhovskaya, M. L.; Wikström, M. Biochemistry 1999, 38, 2697.

(8) (a) Deatherage, J. F.; Henderson, R.; Capaldi, R. A. J. Mol. Biol. 1982, 158, 487. (b) Love, B.; Chan, S. H. P.; Stotz, E. J. Biol. Chem. 1970, 245, 6664.

(9) Yoshikawa, S.; Shinzawa-Itoh, K.; Nakashima, R.; Yaono, R.; Yamashita, E.; Inoue, N.; Yao, M.; Fei, M. J.; Libeu, C. P.; Mizushima, T.; Yamaguchi, H.; Tomizaki, T.; Tsukihara, T. Science 1998, 280, 1723.

(10) (a) Szostak, J. W.; Bartel, D. P.; Luisi, P. L. Nature 2001, 409, 387. (b) Kurihara, K.; Tamura, M.; Shohda, K.-i.; Toyota, T.; Suzuki, K.; Sugawara, T. Nat. Chem. 2011, 3, 775.

(11) (a) Steinberg-Yfrach, G.; Liddell, P. A.; Hung, S.-C.; Moore, A. L.; Gust, D.; Moore, T. A. Nature 1997, 385, 239. (b) Steinberg-Yfrach, G.; Rigaud, J.-L.; Durantini, E. N.; Moore, A. L.; Gust, D.; Moore, T. A. Nature 1998, 392, 479.

(12) Bhosale, S.; Sisson, A. L.; Talukdar, P.; Fürstenberg, A.; Banerji, N.; Vauthey, E.; Bollot, G.; Mareda, J.; Röger, C.; Würthner, F.; Sakai, N.; Matile, S. Science2006, 313, 84.

(13) Choi, H.-J.; Montemagno, C. D. Nano Lett. 2005, 5, 2538.(14) (a) Nardin, C.; Thoeni, S.; Widmer, J.; Winterhalter, M.; Meier, W. Chem.

Commun. 2000, 1433. (b) Palivan, C. G.; Fischer-Onaca, O.; Delcea, M.; Itel, F.; Meier, W. Chem. Soc. Rev. 2012, 41, 2800.

(15) Vriezema, D. M.; Garcia, P. M. L.; Sancho Oltra, N.; Hatzakis, N. S.; Kuiper, S. M.; Nolte, R. J. M.; Rowan, A. E.; van Hest, J. C. M. Angew. Chem. Int. Ed.2007, 46, 7378.

(16) Peterson, J. R.; Smith, T. A.; Thordarson, P. Chem. Commun. 2007, 1899.(17) Ferguson-Miller, S.; Babcock, G. T. Chem. Rev. 1996, 96, 2889.(18) Hvasanov, D.; Wiedenmann, J.; Braet, F.; Thordarson, P. Chem. Commun. 2011,

47, 6314.(19) Damiano, E.; Bassilana, M.; Rigaud, J. L.; Leblanc, G. FEBS Lett. 1984, 166,

120.(20) Agostiano, A.; Mavelli, F.; Milano, F.; Giotta, L.; Trotta, M.; Nagy, L.; Maroti,

P. Bioelectrochemistry 2004, 63, 125.


7-25

(21) (a) Valderrama, B.; García-Arellano, H.; Giansanti, S.; Baratto, M. C.; Pogni, R.; Vazquez-Duhalt, R. FASEB J. 2006, 20, 1233. (b) Michael J, D. Biochem. Biophys. Res. Commun. 2003, 305, 761.

(22) (a) Walde, P.; Ichikawa, S. Biomol. Eng. 2001, 18, 143. (b) Mui, B.; Chow, L.; Hope, M. J. Methods Enzymol. 2003, 367, 3.

(23) Meeuwissen, S. A.; Kim, K. T.; Chen, Y.; Pochan, D. J.; van Hest, J. C. M. Angew. Chem. Int. Ed. 2011, 50, 7070.

(24) Choi, J.; Rubner, M. F. Macromolecules 2005, 38, 116.(25) Madden, T. D.; Hope, M. J.; Cullis, P. R. Biochemistry 1984, 23, 1413.(26) Westerhoff, H. V.; Scholte, B. J.; Hellingwerf, K. J. Biochim. Biophys. Acta,

Bioenerg. 1979, 547, 544.(27) Faxén, K.; Brzezinski, P. Biochim. Biophys. Acta, Bioenerg. 2007, 1767, 381.(28) Discher, D. E.; Eisenberg, A. Science 2002, 297, 967.(29) Hochli, L.; Hackenbrock, C. R. Biochemistry 1978, 17, 3712.

Chapter 8

Experimental

Chapter 8 Experimental

8-2

8 Experimental

8.1 Chemicals, Equipment and General Methods

8.1.1 Chemicals, solvents and materials

Chemicals were purchased from Sigma Aldrich with the exceptions of ammonium

acetate, sodium dihydrogen phosphate, sodium hydroxide, sodium bicarbonate,

anhydrous sodium sulphate, citric acid, iodine, pyridine and hydrazine monohydrate

were purchased from Ajax Finechem Pty. Ltd., 4-nitrobenzaldehyde (Hopkins and

Williams Ltd.) and ammonium hexafluorophosphate (Acros Organics), ruthenium(III)

trichloride hydrate (Precious Metals Online), Oregon Green® 488 carboxylic acid,

succinimidyl ester *5-isomer* and Oregon Green® 488 maleimide (Invitrogen).

Bis(4’-(4-hydroxymethylphenyl)-2,2’:6’:2”-terpyridine)iridium(III)tris(hexafluorophosp

hate) was kindly provided by Dr. Daniel C. Goldstein, N-propargyl maleimide by Dr.

Joshua R. Peterson and 4,4’-bipyridinium-N,N-di(maleimidopropyl) hexaflurophosphate

provided by Mr. Dithepon Pornsaksit. Silica was purchased from Davisil (40-63 μM).

Thin layer chromatography plates (Kieselgel 60 F-254 pre-coated sheets 0.25 mm) were

purchased from Merck. Neutral alumina oxide was purchased from Merck (Alumina

Oxide 90 active neutral, 20 – 230 mesh). Dichloromethane (CH2Cl2) and methanol

(CH3OH) were distilled before use. Dry solvents such as acetonitrile (CH3CN),

dichloromethane, diethyl ether (Et2O) and tetrahydrofuran (THF) were obtained from a

Pure Solv dry solvent system (Innovative Technology, Inc. model #PS-MD-7). Dry

methanol was distilled and stored over calcium chloride. Dry N,N-dimethylformamide

(DMF) was purchased from Sigma Aldrich and used directly from the bottle. Deuterated

solvents for NMR were obtained from Cambridge Isotope Laboratories. For

aggregation studies and preparation of all salt buffers, ultra pure water (R > 18 × 106 )

was used. Water and organic solvent was filtered through a 0.45 m cellulose


8-3

membrane filter (Minisart RC 25, Sartonius Stedim Biotech) prior to polymer

aggregation studies. Diblock copolymer, polystyrene140-b-poly(acrylic acid)48 (PDI =

1.10), was purchased from Encapson (The Netherlands, catalogue number 1036).

Chloroform was distilled and passed through neutral alumina oxide prior to lipid

dissolution. Lipids, extrusion kit (Avanti® Mini-Extruder) and accessories were

purchased from Avanti Polar Lipids. All other chemicals were used as received.

Confocal fluorescence laser-scanning microscopy petri dishes (FluoroDish) were

purchased from World Precision Instruments.

8.1.2 Enzymes and phosphate buffers

Yeast cytochrome c from Saccharomyces cerevisiae (catalogue number C2436), bovine

serum albumin (BSA, catalogue number A-0281), equine heart cytochrome c (catalogue

number C2506), equine skeletal muscle myoglobin (catalogue number M0630), bovine

testes calmodulin (catalogue number P1431) and poly-L-lysine hydrobromide (1000-

5000 Da, catalogue number P0879) were purchased from Sigma Aldrich. Cytochrome c

oxidase was purchased from NBS Biologicals (catalogue number 11197-B). Yeast iso-1

cytochrome c was purified following previously published procedures prior to

bioconjugation reactions.1 Acropora millepora green fluorescent protein (amGFP) was

expressed and purified as previously described and was provided by Dr. Jörg

Wiedenmann (University of Southampton, UK) and Dr. Chris Marquis (University of

New South Wales, Australia).2 For the preparation of salt buffers, ultrapure water (R >

18 × 106 , MilliQ Ultrapure Water System, Millipore) was used. Phosphate buffers

were prepared using concentrated stock solutions of sodium dihydrogen phosphate (1 M,

pH ~4), sodium chloride (5 M), ethylenediaminetetraacetic acid (EDTA, 200 mM) and

diluted with ultrapure water as necessary. Buffer pH values were adjusted with aqueous


8-4

sodium hydroxide (1 M) or hydrochloric acid (1 M) using a Scholar 425 pH meter

(Corning) and filtered using a 0.45 m (Millipore, 47 mm regenerated cellulose) prior to

use. Bioconjugation reactions were prepared as above except a stock solution of sodium

dihydrogen phosphate (97 mM), EDTA (97 mM), pH 7.0 was used to eliminate the need

for adjusting pH of small volume solutions.

8.1.3 Spectroscopy and melting points

A Varian Cary 50 Bio UV-Vis or Cary 5 UV-Vis-NIR spectrometer was used for UV-

Vis spectra measurements. Fluorescence spectra were recorded using a Varian Cary

Eclipse spectrometer with excitation and emission slits at 5 nm, excitation filter “auto”,

emission filter “open” and PMT voltage set to “medium”, unless otherwise stated. NMR

spectra (1H and 13C) were recorded in the designated solvents using a Bruker Avance

DPX (300 MHz) spectrophotometer. Multiplicities are assigned as singlet (s), doublet

(d), triplet (t), quartet (q), pentet (p), multiplet (m) and denoted as broad (br) where

appropriate. Chemical shifts are measured in parts per million (ppm), internally

referenced relative to tetramethylsilane (SiMe4, 1H and 13C = 0 ppm) or residual solvent

peaks (CD3CN: 1H = 1.94 and 13C = 1.32; DMSO-d6: 1H = 2.50, 13C = 39.52; CDCl3:

1H = 7.26 ppm, 13C = 77.16 ppm). IR spectra were recorded on a Shimadzu FTIR-

8400S, ThermoNicolet Avatar model 370 FT-IR or a Perkin Elmer Spotlight 400 FT-IR

spectrometer equipped with a microscope and attenuated total reflectance (ATR)

accessories with diamond crystal inset. Intensity abbreviations are as follows: weak (w),

medium (m), strong (s). Low resolution Electrospray Ionisation (ESI) mass spectra were

recorded on a Waters Micromass ZQ electrospray instrument or on a Shimadzu

Prominence UFLC system equipped with a Shimazdu 2010 EV LC-MS detector and a

Shimazdu FRC-10A fraction collector. High resolution ESI mass spectrometry was


8-5

performed on a Thermo Linear Quadropole Ion Trap Fourier Transform Ion Cyclotron

Resonance (LQT FT Ultra) mass spectrometer in electrospray mode with a 7 T

superconducting magnet or a Thermo Scientific LTQ Orbitrap XL hybrid Fourier

Transform Mass Spectrometer (FTMS) in static nanospray mode. MALDI-TOF mass

spectra were recorded on an Applied Biosystems Voyager DE STR MALDI reflectron

TOF MS (protein and bioconjugate measurements were made in linear mode). Melting

points were recorded on a Mel-temp II hot stage apparatus.

8.1.4 Protein and bioconjugate purification equipment

Protein and bioconjugate purification was performed using a GE Healthcare

ÄKTApurifier. Cation exchange chromatography (CEX) was performed using a strong

cation exchange column (TSKgel SP-5PW, Supelco). Immobilised metal affinity

chromatography (IMAC) was performed using either a Ni2+ charged HisTrap HP (GE

Healthcare, 1 mL) or an Acrosep Hypercel (Pall, 1 mL) column. Size exclusion

chromatography (SEC) was performed using either a Superdex 75 10/300 GL (GE

Healthcare) or Sephadex G25 desalting (Superfine, GE Healthcare) column.

Programmed elution gradients are expressed in column volumes (CV) to ease

comparison between columns of different size. Flow rates were based on manufacturer

recommendations for flow and pressure limits and were typically between 0.5 to

1 mL/min. Fractions were collected in 1 mL Eppendorf tubes using either a Frac 901 or

Frac 902 collector (GE Healthcare). Protein solutions were concentrated using 3 000

molecular weight cut-off (MWCO) centrifuge concentrators (Amicon Ultra-15, Amicon

Ultra-4 or Amicon Ultra-0.5, Millipore) and centrifugation at 4 000 rpm in a Sigma 2-6

for large volume samples (<50 mL) or 13 800 rpm in a Sigma 1-14 bench top centrifuge

(John Morris Scientific) for small volume samples (<1.5 mL). Samples were dialysed


8-6

into MilliQ water using Slide-A-Lyzer Mini Dialysis Units (3 500 MWCO, Pierce) or

Tube-O-Dialyzer Dialysis Units (4 000 MWCO, G Biosciences).

8.1.5 Preparation of samples for analysis by MALDI-TOF mass spectrometry

Protein and bioconjugate samples for MALDI were prepared by diluting 5 L of protein

with either 5 L of a saturated solution of sinapic acid and -cyano-4-hydroxycinnamic

acid (10:1 w/w ratio in acetonitrile/water/trifluoroacetic acid (70:30:0.03, v/v/v)) or a

solution of caffeic acid (10 mg/mL in acetonitrile/water/trifluoroacetic acid (80:20:0.1,

v/v/v)). Prepared samples were spotted by adding a 0.5 L droplet (× 2) onto a MALDI

target plate. At times, ZipTips (C4, 0.6 L bed volume, Millipore) were used for

desalting by priming the tip with acetonitrile/water/trifluoroacetic acid (80:20:0.1, v/v/v,

3 × 10 L), washing with acetonitrile/water/trifluoroacetic acid (2:98:0.1, v/v/v,

3 × 10 L), loading the column (5 × 10 L), washing with

acetonitrile/water/trifluoroacetic acid (2:98:0.1, v/v/v, 3 × 10 L) and eluting with

2.5 L of a caffeic acid solution (10 mg/mL in acetonitrile/water/trifluoroacetic acid

(80:20:0.1, v/v/v)) directly onto the MALDI target plate.

8.1.6 Transmission electron microscopy (TEM) studies

TEM micrographs were recorded on a JEOL 1400 (80 kV for polymersomes and

100 kV for liposomes) instrument and cryo-TEM micrographs were recorded on a

JEOL 2100 (200 kV) instrument. Conventional TEM samples were prepared by placing

20 L of sample onto a formvar-coated copper grid and the excess water was blotted

away after 2 min with a filter paper. For statistical analysis, a population of 100 micelles

or polymersomes were measured from TEM micrographs for determination of the

average and standard deviation of diameters. For analysis of polymersome-to-micelle


8-7

(p/m) ratio, a population of 100 aggregates from TEM micrographs were selected and

aggregate morphologies displaying distinct contrast within individual particles

(indicating the presence of bilayer and cavity formation) were classified as vesicles

whereas micelles were defined as particles showing uniform contrast. Ratio

measurements were repeated in triplicate. For cryo-TEM, 3 L of sample was directly

placed onto glow-discharged holey carbon grids (Quantifoil, Germany). Grids were

blotted once at a blotting angle of 2 mm for 2 s under 100% relative humidity at 25 oC

and subsequently plunged into liquid ethane using the automated vitrobot (F.E.I, The

Netherlands). Vitrified samples were stored in liquid nitrogen upon cryo transfer for

cryo-TEM investigation.3

8.1.7 Confocal laser-scanning microscopy studies

Optical and confocal laser microscopy experiments were carried out with an Olympus

Fluoview FV1000, fitted with monochromatic laser light sources for fluorescence

measurements. Images were acquired on a confocal laser scanning microscope with a

40x 0.9NA water-immersion objective. Excitation was at 488 nm using an Ar+ laser.

Detection was in the range 500-600 nm using internal PMTs (gains were set to 468 V).

The confocal pinhole (aperture) was set to auto and images with a 640×640 pixel

resolution were recorded at a scan rate of 40 s/pixel with a total acquisition time of

16 s.

8.1.8 amGFP (amFP497) (Acropora millepora) expression and isolation

The following procedure was performed in the laboratory of Dr. Jörg Wiedenmann at

the University of Southampton. The coding sequence of the green fluorescent protein

(GFP)-like protein from the reef coral Acropora millepora (amFP497)4 was introduced


8-8

in the plasmid pQE32 (Qiagen, Hilden, Germany), resulting in the addition of an N-

terminal 6×histidine tag to the recombinant protein. Bacteria (Escherichia coli, M15

pREP4) were transformed with the plasmid and grown at 37 ºC in 2YT medium to an

optical density of 0.6. Expression of the protein was induced with isopropyl- -D-

thiogalactopyranosid (IPTG) and the culture was incubated at 20 ºC on a shaker at

220 rpm for 12 h. Cultures were slightly agitated further for 7 days at 4 ºC to increase

the yield of soluble protein.5 Subsequently, cells were harvested and the recombinant

protein was purified by immobilised metal ion chromatography using TalonTM matrix

(Clontech; Palo Alto, USA), following the protocol of the manufacturer. Yields were

determined by UV-Vis spectroscopy ( 476 = 31.42 mM-1cm-1).

8.1.9 Gel electrophoresis

Gel electrophoresis was performed using Invitrogen Novex® NuPage® 12% Bis-Tris,

1 mm, 10-well gels, SeeBlue® Plus2 molecular weight marker, NuPage® LDS sample

buffer (4×), NuPage® sample reducing agent (10×), NuPage® MES SDS running

buffer, SimplyBlueTM safestain and the gels run using a Zoom Dual Power supply

(model ZP10002, Invitrogen). Samples for gel electrophoresis were prepared by dilution

in Novex® NuPage® LDS sample buffer (Invitrogen). Samples were reduced (to

eliminate disulfide dimers) by adding NuPage® sample reducing agent (active

ingredient dithiothreitol (DTT)). Samples were heated at 70 oC for 10 min to denature

the protein. Novex® NuPage® gels (12% Bis-Tris, 10-wells) were then loaded with

1-3 g of protein per well, run at a constant 200 V for 40 min and stained according to

the procedure included with SimplyBlueTM safestain.


8-9

8.2 X-ray Crystallography

8.2.1 Crystal growth of Ru(II)-bisterpyridine complexes

In general, Ru(II)-bisterpyridine complex (ca. 5 mg) was dissolved in minimal solvent

(DMF) resulting in a concentrated solution. Crystals were grown by slow diffusion of

anhydrous diethyl ether into a solution of complex in N,N-dimethylformamide at room

temperature or 4 oC for complex 7 and 8, respectively. A suitable single crystal was

selected under a polarising microscope (Leica M165Z) for single crystal X-ray

diffraction analysis.

8.2.2 X-ray structure determination

8.2.2.1 Complex 7

The X-ray diffraction measurement for complex 7 was performed on a Bruker kappa

APEX-II CCD diffractometer at 150 K by using graphite-monochromated Mo-K

radiation ( = 0.71075 Å). The crystal was mounted on the goniometer using cryo loops

for intensity measurements, coated with paraffin oil and immediately transferred to the

cold stream using Oxford Cryostream 700 system attachment. Upon obtaining an initial

refinement of unit cell parameters, the data collection strategy was calculated to achieve

a redundancy of at least 4 throughout the resolution range ( - 0.80 Å) at 10 s exposure

time per frame utilising the kappa offsets on the four circle goniometer geometry. Data

integration, reduction with multi-scan absorption correction method was carried out

using Bruker APEX2 Suite software.6 The structure was solved by Direct Methods

program SHELXS-97 and refined by full-matrix least-squares refinement program

SHELXL.7 All non-hydrogen atoms were refined anisotropically and hydrogen atoms

were included by using a riding model. Further crystal and refinement data given in

Appendix A.


8-10

8.2.2.2 Complex 8

The X-ray diffraction measurement for complex 8 was carried out at MX1 beamline at

the Australian Synchrotron Facility, Melbourne. The crystal was mounted on the

goniometer using cryo loop for intensity measurements, coated with paraffin oil and

immediately transferred to the cold stream using a Cryostream attachment. Data was

collected using Si<111> monochromated synchrotron X- = 0.71023 Å)

at 100(2) K and was corrected for Lorentz and polarization effects using the XDS

software.8 The structure was solved by Direct methods and the full-matrix least-squares

refinements was carried out using SHELXL.7 Further crystal and refinement data given

in Appendix A.

8.3 Synthesis of Terpyridine Chromophores

8.3.1 4-nitro-2’-azachalcone (1)9

To a solution of 4-nitrobenzaldehyde (2.08 g, 13.8 mmol) and aqueous sodium

hydroxide (1 M, 13.75 mL) in methanol (60 mL) was added 2-acetylpyridine (1.60 mL,

14.3 mmol). The solution was stirred for 1 h, filtered, and the collected precipitate

washed with cold methanol, dissolved in dichloromethane, washed with water, dried

over anhydrous sodium sulphate and the solvent removed in vacuo. The resulting

yellow solid was recrystallised from ethanol to give azachalcone 1 as yellow crystals

(1.01 g, 29%). 1H NMR (300 MHz, CDCl3) 8.76 (d, J = 4.9 Hz, 1H), 8.43 (d,

J = 16.2 Hz, 1H), 8.28 (d, J = 9.0 Hz, 2H), 8.21 (d, J = 7.9 Hz, 1H), 7.96 – 7.84 (m,

4H), 7.56 – 7.51 (m, 1H). MS (ESI) m/z: ([M + H]+) calcd. for C14H11N2O3, 255.08;

found, 254.96. These results are in agreement with those reported in the literature.9


8-11

8.3.2 1-(2-oxo-2-(2-pyridyl)ethyl)pyridinium iodide (2)10

To a stirred solution of iodine (7.78 g, 30.7 mmol) in dry pyridine (20 mL) under

nitrogen at 60 oC was added 2-acetylpyridine (3.68 g, 30.4 mmol). The resulting

mixture was stirred at 100 oC for 2 h, cooled, then the crystals filtered and washed with

chloroform and diethyl ether to yield pyridinium iodide 2 as black crystals (7.84 g,

79%). 1H NMR (300 MHz, DMSO-d6) 9.00 (d, J = 5.6 Hz, 2H), 8.87 (d, J = 6.0 Hz,

1H), 8.73 (t, J = 7.2 Hz, 1H), 8.27 (t, J = 7.2 Hz, 2H), 8.18 – 8.03 (m, 2H), 7.83 (td,

J = 5.84, 1.5 Hz, 1H), 6.50 (s, 2H). MS (ESI) m/z: ([M - I]+) calcd. for C12H11N2O,

199.08; found, 199.09. These results are in agreement with those reported in the

literature.10

8.3.3 4’-(4-nitrophenyl)-2,2’:6’,2’’-terpyridine (3)9

A solution of azachalcone 1 (0.870 g, 3.42 mmol), pyridinium iodide 2 (1.13 g,

3.47 mmol) and ammonium acetate (3.75 g, 48.6 mmol) in dry methanol (60 mL) was

refluxed for 20 h. The crystals were cooled, filtered and washed with cold methanol

(6 × 50 mL) to yield terpyridine 3 as a purple solid (0.91 g, 75%). 1H NMR (300 MHz,

CDCl3) 8.76 (s, 2H), 8.74 (d, J = 4.9 Hz, 2H), 8.69 (d, J = 7.9 Hz, 2H), 8.38 (d,

J = 8.7 Hz, 2H), 8.06 (d, J = 8.7 Hz, 2H), 7.91 (td, J = 7.7, 1.9 Hz, 2H), 7.42 – 7.35 (m,

2H). MS (ESI) m/z: ([M + H]+) calcd. for C21H15N4O2, 355.12; found, 354.95. These

results are in agreement with those reported in the literature.9

8.3.4 4’-(4-aminophenyl)-2,2’:6’,2’’-terpyridine (4)11

A solution of compound 3 (891.8 mg, 2.52 mmol) in absolute ethanol (60 mL) was

refluxed for 45 min in the presence of 10% Pd/charcoal catalyst (120.6 mg). Hydrazine

monohydrate (4.75 mL, 98.0 mmol) in absolute ethanol (60 mL) was then added drop-


8-12

wise to the reaction mixture and refluxed for 3 h. The solution was filtered over celite

and washed with dichloromethane (150 mL). The organic phase was washed with water

(4 × 100 mL), dried over anhydrous sodium sulphate and solvent removed in vacuo to

yield terpyridine 4 as yellow crystals (658.4 mg, 81%). 1H NMR (300 MHz, CDCl3)

8.73 (d, J = 4.5 Hz, 2H), 8.69 (s, 2H), 8.66 (d, J = 7.9 Hz, 2H), 7.87 (td, J = 7.7, 1.9 Hz,

2H), 7.79 (d, J = 8.7 Hz, 2H), 7.34 (t, J = 6.2 Hz, 2H), 6.80 (d, J = 8.7 Hz, 2H), 5.61 (s,

2H). MS (ESI) m/z: ([M + H]+) calcd. for C21H17N4, 325.14; found, 325.04. These

results are in agreement with those reported in the literature.11

8.3.5 [Ru(tpy)]Cl3 (5)12

A solution of terpyridine (418.9 mg, 1.795 mmol), and ruthenium(III) trichloride hydrate

(722.0 mg, 3.481 mmol) in absolute ethanol (50 mL) was refluxed for 23 h under

nitrogen. The product was collected by filtration and washed with absolute ethanol,

water and diethyl ether yielding [Ru(tpy)]Cl3 5 as a black solid (695 mg, 88%). MS

(ESI) m/z: ([M – 2Cl]+) calcd. for C15H11N3RuCl+, 369.97, found, 369.93. These results

are in agreement with those reported in the literature.12

8.3.6 [Ru(tpy)2](PF6)2 (6)12b

A solution of 2,2’;6’,2”-terpyridine (113.8 mg, 0.488 mmol) and ruthenium(III)

trichloride hydrate (49.6 mg, 0.239 mmol) in ethylene glycol (33 mL) was heated at

110 oC for 21 h. The solution was then diluted with water (150 mL), filtered through

celite and the product was precipitated with ammonium hexafluorophosphate. The solid

was collected by centrifugation, washed with water and recrystallised with

acetonitrile/diethyl ether and collected by filtration and washed with acetonitrile

yielding [Ru(tpy)2](PF6)2 6 as a red solid (103.1 mg, 50%). 1H NMR (300 MHz,


8-13

CD3CN) 8.73 (d, J = 8.1 Hz, 4H), 8.48 (dt, J = 8.1, 1.0 Hz, 4H), 8.40 (t, J = 8.3 Hz,

2H), 7.91 (td, J = 7.8, 1.5 Hz, 4H), 7.32 (dq, J = 5.6, 0.8 Hz, 4H), 7.19 – 7.11 (m, 4H).

MS (ESI) m/z: ([M – 2PF6]2+) calcd. for C30H22N6Ru2+, 284.05; found, 283.80. These

results are in agreement with those reported in the literature.12b

8.3.7 [Ru(tpy)(4’-(4-aminophenyl)-2,2’:6’,2”-tpy)](PF6)2 (7)

Method 1

A solution of terpyridine 4 (273.3 mg, 0.8425 mmol) and [Ru(tpy)]Cl3 5 (373.6 mg,

0.8447 mmol) in ethylene glycol (50 mL) was heated to 110 oC for 20 h under nitrogen.

Reaction mixture was diluted to 150 mL with water and filtered over celite. Filtrate was

precipitated using ammonium hexafluorophosphate, washed with water (3 × 50 mL) and

collected with acetonitrile. Purified over silica using a gradient from 90:9:1

CH3CN:H2O:KNO3 (saturated) to 20:3:1 CH3CN:H2O:KNO3 (saturated). Fractions

precipitated with ammonium hexafluorophosphate and washed with water. Product

further purified over alumina (neutral) using a gradient from acetonitrile to 90:9:1

CH3CN:H2O:KNO3 (saturated). Fractions pooled, precipitated with ammonium

hexafluorophosphate, washed with water (3 × 20 mL) and collected with acetonitrile

yielding complex 7 as a red solid (609.6 mg, 76%). mp >274 oC (decomposed); 1H

NMR (300 MHz, CD3CN) 8.90 (s, 2H), 8.72 (d, J = 7.9 Hz, 2H), 8.60 (d, J = 7.9 Hz,

2H), 8.48 (d, J = 7.9 Hz, 2H), 8.38 (t, J = 8.3 Hz, 1H), 8.00 (dt, J = 8.6, 2.4 Hz, 2H),

7.91 (tt, J = 7.9, 1.3 Hz, 4H), 7.44 (dd, J = 4.9, 0.8 Hz, 2H), 7.30 (dd, J = 4.5, 0.8 Hz,

2H), 7.21 – 7.09 (m, 4H), 6.95 (dt, J = 8.7, 2.4 Hz, 2H); 13C NMR (75 MHz, CD3CN)

159.1, 156.5, 156.0, 153.5, 153.3, 144.9, 138.9, 136.4, 129.8, 128.4, 128.2, 125.3,

124.6, 120.9, 115.8; IR (ATR) max/cm-1 3646 (w), 3593 (w), 3506 (w), 3414 (w), 3121

(w), 1990 (w), 1633 (m), 1597 (m), 1529 (w), 1430 (m); UV-Vis (CH3CN) max/nm


8-14

( /M-1cm-1) 490 (1.71 × 104), 364 (9.80 × 103), 308 (5.00 × 104), 283 (2.55 × 104), 272

(2.86 × 104), 231 (2.67 × 104); HRMS (ESI) m/z: ([M – PF6)]+) calcd. for

C36H27N7P1F6Ru+, 804.1023; found, 804.1006. MS (ESI) m/z: ([M – 2PF6]2+) calcd.

for C36H27N7Ru2+, 329.57; found, 329.49.

Method 2

A solution of terpyridine 4 (179.0 mg, 0.5518 mmol) and [Ru(tpy)]Cl3 5 (248.8 mg,

0.5645 mmol) in ethanol (100 mL) was refluxed for 20 h under nitrogen. Reaction

mixture was filtered over celite, concentrated in vacuo and diluted with water (100 mL).

Filtrate was precipitated using ammonium hexafluorophosphate, washed with water

(3 × 50 mL) and collected with acetonitrile. The product was recrystallised with

acetonitrile/diethyl ether yielding complex 7 as a red solid (138.7 mg, 28%).

Characterisation data was identical to the compound obtained from method 1 above.

8.3.8 [Ru(tpy)(maleimide-hexylcarboxamido-phenyl-tpy)](PF6)2 (8)

A solution of 6-maleimidocaproic acid (53.2 mg, 0.252 mmol), O-(7-azabenzotriazole-

1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU, 95.9 mg, 0.252 mmol)

and N,N-diisopropylethylamine (66.8 mg, 0.517 mmol) in dry dimethylformamide

(5 mL) was stirred at room temperature under nitrogen for 1 h. Subsequently, complex 7

(71.7 mg, 0.0756 mmol) in dry dimethylformamide (5 mL) was added to the mixture

and stirred for a further 26 h in the dark at room temperature under nitrogen.

Dichloromethane (50 mL) was added to the solution and the organic phase washed with

aqueous citric acid (10% w/v, 2 × 20 mL), water (3 × 10 mL) and dried over anhydrous

sodium sulphate. Dichloromethane was removed in vacuo and the concentrated red

dimethylformamide phase containing product was precipitated from dry diethyl ether


8-15

and the solid collected by filtration, washed with diethyl ether and collected with

acetonitrile. Product was purified over silica using a gradient from acetonitrile to

70:29:1 CH3CN:H2O:KNO3 (saturated). Fractions pooled, precipitated with ammonium

hexafluorophosphate, washed with water (3 × 20 mL) and collected with acetonitrile

yielding complex 8 as a red solid (27.9 mg, 41%). mp >246 oC (decomposed); 1H NMR

(300 MHz, CD3CN) 8.98 (s, 2H), 8.75 (d, J = 8.3 Hz, 3H), 8.63 (d, J = 7.9 Hz, 2H),

8.50 (d, J = 7.9 Hz, 2H), 8.40 (t, J = 8.3 Hz, 1H), 8.16 (dd, J = 8.7, 1.5 Hz, 2H), 8.00 –

7.85 (m, 6H), 7.43 (dd, J = 5.6, 0.8 Hz, 2H), 7.37 (dd, J = 6.4, 0.8 Hz, 2H), 7.20 – 7.10

(m, 3H), 6.75 (s, 2H), 3.49 (t, J = 7.0 Hz, 2H), 2.41 (t, J = 7.4 Hz, 2H), 1.80 – 1.56 (m,

5H), 1.45 – 1.33 (m, 2H); 13C NMR (75 MHz, CD3CN) 173.60, 172.51, 159.49,

159.41, 159.26, 156.56, 153.61, 153.48, 149.09, 142.43, 139.19, 136.87, 135.37,

132.53, 129.51, 128.61, 128.56, 125.64, 125.56, 124.85, 122.17, 121.17, 38.44, 37.76,

29.17, 27.21, 25.94; IR (ATR) max/cm-1 3647 (w), 3403 (w), 3112 (w), 2933 (w), 2857

(w), 1769 (w), 1699 (s), 1595 (m), 1523 (m), 1449 (m), 1407 (m); UV-Vis (CH3CN)

max/nm ( /M-1cm-1) 485 (2.60 × 104), 410 (4.88 × 103), 308 (7.85 × 104), 282

(4.32 × 104), 272 (4.65 × 104); HRMS (ESI) m/z: ([M – PF6])+ calcd. for

C46H38N8O3P1F6Ru+, 997.1618; found, 997.1741 and ([M – 2PF6])2+ calcd. for

C46H38N8O3Ru2+, 426.0988; found, 426.1050. MS (ESI) m/z: ([M – 2PF6])2+, 426.11.

Prior to bioconjugation, complex 8 was exchanged with chloride salt to increase

solubility and yield.

8.3.9 [Ru(4’-(4-aminophenyl)-2,2’:6’2’’-terpyridine)2](PF6)2 (9)13

A solution of compound 4’-(4-aminophenyl)-2,2’:6’,2’’-terpyridine 4 (141.2 mg,

0.4353 mmol) and ruthenium(III) trichloride hydrate (44.3 mg, 0.214 mmol) in ethylene

glycol (50 mL) was stirred at 110 oC for 21 h. The reaction mixture was diluted with


8-16

water (150 mL) and filtered over celite. The product was precipitated using ammonium

hexafluorophosphate and collected by centrifugation (4 min, 4 000 rpm) and washed

with water (3 × 40 mL). The product was recrystallised with acetonitrile/diethyl ether to

yield terpyridine 9 as red crystals (116.3 mg, 52%). 1H NMR (300 MHz, CD3CN)

8.90 (s, 4H), 8.60 (d, J = 7.9 Hz, 4H), 8.00 (d, J = 8.7 Hz, 4H), 7.91 (td, J = 7.8, 1.5 Hz,

4H), 7.41 (d, J = 4.9 Hz, 4H), 7.20 – 7.09 (m, 4H), 6.95 (d, J = 8.7 Hz, 4H), 4.77 (s,

4H). MS (ESI) m/z: ([M – 2PF6]2+) calcd. for C42H32N8Ru2+, 375.09; found, 374.72.

These results are in agreement with those reported in the literature.13

8.3.10 [Ru(4’-(4-maleimide-hexylcarboxyamido-phenyl)-2,2’:6’2’’-

terpyridine)2](PF6)2 (10)11b

A solution of 6-maleimidocaproic acid (73.8 mg, 0.349 mmol), O-(7-azabenzotriazole-

1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU, 136.1 mg,

0.3580 mmol) and N,N-diisopropylethylamine (96.5 mg, 0.746 mmol) in dry

dimethylformamide (5 mL) was stirred at room temperature for 1 h. Complex [Ru(4’-

(4-aminophenyl)-2,2’:6’2’’-terpyridine)2](PF6)2 9 (62.5 mg, 0.0601 mmol) in dry

dimethylformamide (5 mL) was added to the reaction mixture and stirred for an

additional 25 h in the dark. Dichloromethane (50 mL) was added and the organic phase

was washed with 10% w/v citric acid (2 × 10 mL), water (3 × 10 mL), dried over

anhydrous sodium sulfate, filtered and dichloromethane removed in vacuo. The

concentrated dimethylformamide phase was diluted and precipitated from dry diethyl

ether and solid collected by filtration, washed with diethyl ether and collected with

acetonitrile. The dried solid was purified on silica (acetonitrile/saturated aqueous

potassium nitrate/water, 20:1:3 (v/v/v)) followed by concentration of the product

containing fractions, collection of the precipitate, washing with water and collection


8-17

with acetonitrile afforded the complex 10 as a red solid (22.8 mg, 27%). 1H NMR

(300 MHz, CD3CN) 8.98 (s, 4H), 8.63 (d, J = 8.3 Hz, 6H), 8.16 (d, J = 8.7 Hz, 4H),

7.93 (td, J = 7.9, 1.5 Hz, 8H), 7.42 (d, J = 5.7 Hz, 4H), 7.17 (td, J = 6.7, 1.5 Hz, 4H),

6.75 (s, 4H), 3.49 (t, J = 7.0 Hz, 4H), 2.41 (t, J = 7.4 Hz, 4H), 1.78 – 1.68 (m, 4H),

1.68 – 1.57 (m, 4H), 1.46 – 1.31 (m, 4H). MS (ESI) m/z: ([M – 2PF6]2+) calcd. for

C62H54N10O6Ru2+, 568.16; found, 568.14. These results are in agreement with those

previously reported.11b

Prior to bioconjugation, complex 10 was exchanged with chloride salt to increase

solubility and yield.

8.3.11 Anion exchange of Ru(II)-bisterpyridine complexes 8 and 10

Hexafluorophosphate salts of complexes 8 and 10 (10 mg) were dissolved in acetonitrile

to which excess tetrabutylammonium chloride was added until precipitation was

complete. The mixture was centrifuged to a pellet and solution was decanted. The pellet

was resuspended in acetonitrile and recrystallised from diethyl ether followed by

centrifugation (3 ×) to give the chloride adducts of complexes 8 and 10 which were

subsequently used without further purification or characterisation for bioconjugation

synthesis.

8.4 Synthesis of Anthraquinone-Based Acceptors

8.4.1 1-amino-3-azidopropane (11)14

Sodium azide (1.002 g, 15.42 mmol) was added slowly to a solution of

1-amino-3-bromopropane hydrobromide (840.8 mg, 3.840 mmol) in water (50 mL) and

refluxed under nitrogen for 23 h. The solution was allowed to cool to 0 oC, extracted

with diethyl ether (3 × 50 mL), dried with anhydrous sodium sulphate and filtered.


8-18

Solvent was removed in vacuo to yield a clear yellow oil (166.2 mg, 43%). 1H NMR

(300 MHz, CDCl3) 3.39 (t, J = 6.6 Hz, 2H), 2.83 (t, J = 7.5 Hz, 2H), 1.75 (p, J = 6.5

Hz, 2H), 1.42 (br s, 2H). MS (ESI) m/z: ([M + H]+) calcd. for C3H9N4+, 101.08; found,

100.80. These results are in agreement with those previously reported.14

8.4.2 Anthraquinone-2-carboxylic acid N-hydroxysuccinimide ester (12)

Anthraquinone-2-carboxylic acid (604.7 mg, 2.398 mmol) and N-hydroxysuccinimide

(275.8 mg, 2.396 mmol) was dissolved in dry dichloromethane (150 mL). The solution

was cooled to 0 oC and N,N’-dicyclohexylcarbodiimide (687.9 mg, 3.330 mmol) was

added. The reaction mixture was stirred at room temperature under nitrogen in the dark

for 23 h. The resulting cloudy yellow mixture was filtered and solvent removed in

vacuo yielding the NHS-ester of anthraquinone-2-carboxylic acid 12 as a yellow solid

(1.013 g, crude). The crude was used for subsequent reactions without further

purification.

8.4.3 Anthraquinone-2-azidopropylamide (13)

The NHS-ester of anthraquinone-2-carboxylic acid 12 (1.013 g, crude) was suspended

into a cloudy solution in acetone/ethanol (75 mL/50 mL). To this stirring solution, an

aqueous solution of sodium bicarbonate (223.0 mg, 2.655 mmol) and

1-amino-3-azidopropane 11 (166.2 mg, 1.660 mmol) in water (25 mL) was added and

allowed to stir for 23 h at room temperature in the dark under nitrogen. Organic phase

was removed in vacuo and dichloromethane (250 mL) was added, washed with basic

water (3 × 200 mL, pH 14), followed by acidic water (3 × 200 mL, pH 2). Organic

phase was dried with anhydrous sodium sulphate and purified on silica

(dichloromethane) followed by concentration of the product containing fractions


8-19

yielding the anthraquinone 13 as a yellow solid (196.6 mg, 35%). mp >160 oC

(decomposed); 1H NMR (300 MHz, CDCl3) 8.58 (d, J = 2.1 Hz, 1H), 8.44 – 8.25 (m,

4H), 7.89 – 7.81 (m, 2H), 6.61 (br s, 1H), 3.64 (q, J = 6.4 Hz, 2H), 3.50 (t, J = 6.5 Hz,

2H), 1.97 (p, J = 6.5 Hz, 2H); 13C NMR (75 MHz, CDCl3) 182.68, 182.57, 165.87,

139.43, 135.40, 134.67, 134.60, 133.58, 133.26, 128.19, 127.61, 124.93, 49.74, 38.35,

28.87; IR (KBr) max/cm-1 3321 (m), 3288 (s), 3073 (w), 3031 (w), 2928 (m), 2850 (m),

2192 (w), 2113 (s), 1677 (s), 1632 (s), 1590 (s), 1571 (m), 1544 (s), 1472 (m), 1463

(m), 1449 (w), 1436 (w); HRMS (ESI) m/z: ([M + H])+ calcd. for C18H15N4O3+,

335.1144; found, 335.1134 and ([M + Na])+ calcd. for C18H14N4O3Na+, 357.0964;

found, 357.0953. MS (ESI) m/z: ([M - H])- calcd. for C18H13N4O3-, 333.09, found,

332.90.

8.4.4 Anthraquinone-2-bromopropylamide (14)

The NHS-ester of anthraquinone-2-carboxylic acid 12 (691.5 mg, crude) was suspended

into a cloudy solution in acetone/ethanol (75 mL/50 mL). To this stirring solution, an

aqueous solution of sodium bicarbonate (281.6 mg, 3.35 mmol) and

3-bromopropylamine hydrobromide (373.6 mg, 1.707 mmol) in water (25 mL) was

added and allowed to stir for 18 h at room temperature in the dark under nitrogen.

Organic phase was removed in vacuo and dichloromethane (200 mL) was added,

washed with basic water (3 × 100 mL, pH 14), followed by acidic water (3 × 100 mL,

pH 2). Organic phase was dried with anhydrous sodium sulphate and purified on silica

(dichloromethane) followed by concentration of the product containing fractions

yielding the anthraquinone 14 as a yellow solid (344.5 mg, 56%). mp >178 oC

(decomposed); 1H NMR (300 MHz, CDCl3) 8.58 (d, J = 1.5 Hz, 1H), 8.44 – 8.27 (m,

4H), 7.89 – 7.81 (m, 2H), 6.58 (t, J = 7.5 Hz, 1H), 3.70 (q, J = 6.4 Hz, 2H), 3.53 (t,


8-20

J = 6.3 Hz, 2H), 2.27 (p, J = 6.5 Hz, 2H); 13C NMR (75 MHz, CDCl3) 182.65, 182.53,

166.00, 139.38, 135.37, 134.66, 133.55, 133.27, 128.14, 127.59, 124.99, 49.34, 39.20,

34.10, 32.19, 30.92, 25.75, 25.08; IR (KBr) max/cm-1 3318 (m), 3072 (w), 2929 (m),

2851 (w), 1669 (s), 1635 (s), 1591 (s), 1554 (m), 1447 (w), 1413 (w); UV-Vis (CH2Cl2)

max/nm ( /M-1cm-1) 328 (3.38 × 103), 258 (2.96 × 104); HRMS (ESI) m/z: ([M + H])+

calcd. for C18H15NO3Br+, 372.0235; found, 372.0226 and ([M + Na])+ calcd. for

C18H14NO3BrNa+, 394.0055; found, 394.0047. MS (ESI) m/z: ([M + H])+, 373.80.

8.4.5 Attempted synthesis of anthraquinone-2-propylamido-triazole-maleimide

(15)

Anthraquinone-2-azidopropylamide 13 (59.8 mg, 0.179 mmol) and N-propargyl

maleimide (26.8 mg, 0.198 mmol) was dissolved in tetrahydrofuran (10 mL). Copper(II)

sulphate pentahydrate (67.0 mg, 0.268 mmol) and L-ascorbic acid (65.4 mg,

0.371 mmol) in water (2 mL) was slowly added to the reaction mixture. Solution was

stirred in the dark under nitrogen at room temperature for 22.5 h. Organic phase was

removed in vacuo, resuspended in dichloromethane (100 mL), washed with water

(3 × 100 mL), dried with anhydrous sodium sulphate, filtered and solvent removed in

vacuo to yield a yellow oil (34.8 mg). 1H NMR spectroscopy and mass spectrometry

analysis of the crude product did not indicate that the desired compound 15 was present.

8.5 Synthesis and Purification of Bioconjugates

8.5.1 Purification of iso-1 cytochrome c

Crude yeast cytochrome c from Saccharomyces cerevisiae (11.3 mg) was dissolved in

phosphate buffer (5.65 mL, 20 mM, pH 7.0), reduced with dithiothreitol (DTT, 40 L of

1 M stock), loaded onto a Supelco strong cation exchange column (TSKgel SP-5PW,


8-21

10 m resin beads, 7.5 cmh × 0.75 cmd). The protein was eluted using a sodium

chloride gradient from 328 mM to 450 mM in 14.5 mL at pH 7.0 and 1 mL/min. The

main peak (eluting from 15.8 to 18 mL) was collected and concentrated using a

Millipore 3 000 molecular weight cut-off (MWCO) centrifuge concentrator giving pure

iso-1 cytochrome c in 54% yield based on UV-Vis absorbance of the final product

( 410 = 97.6 mM-1cm-1)1,11b. MS (MALDI) m/z: 12 705 ([M]+ requires 12 706).

8.5.2 8-cyt c

A solution of complex (chloride counter ion) 8 (0.900 mg, 0.975 mol) in acetonitrile

(613 L) was added to a solution of 94 mM sodium dihydrogen phosphate, 94 mM

ethylenediaminetetraacetic acid, pH 7.0 (3.19 mL) in water (10.9 mL) at room

temperature. Purified, reduced iso-1 cytochrome c (1.90 mg, 0.150 mol) in water

(351 L) was then added and the resulting solution stirred in darkness at room

temperature for 20 h. The reaction mixture was then concentrated, dialysed into water

and purified by immobilised metal affinity chromatography (IMAC, HisTrapTM HP, GE

Healthcare or Acrosep HypercelTM, Pall) using a gradient from 0 to 125 mM imidazole

in 20 mM sodium dihydrogen phosphate, 0.5 M sodium chloride, pH 7.0 from 7.7 to

19.2 mL at 0.5 mL/min. The product fraction (eluting from 11.1 to 18.9 mL) was

pooled, concentrated and dialysed into water to yield bioconjugate 8-cyt c. (9.33 nmol,

6%). MS (MALDI) m/z: 13 555 ([M – 2Cl]+ requires 13 559).

8.5.3 cyt c-10-BSA dimer

A solution of complex 10 (chloride counter ion) (0.0410 mg, 0.0340 mol) in

acetonitrile (170 L) was added to a solution of 94 mM sodium dihydrogen phosphate,

94 mM ethylenediaminetetraacetic acid, pH 7.0 (723 L) in water (1.87 mL) at room


8-22


(636 L) was then added and the resulting solution stirred in darkness at room

temperature for 2 h. Bovine serum albumin (45.9 mg, 0.687 mol) was subsequently

added to the mixture and stirred in darkness at room temperature for an additional 21 h.

The reaction mixture was then concentrated, dialysed into water and purified by

immobilised metal affinity chromatography (IMAC, HisTrapTM HP, GE Healthcare)

using a gradient from 0 to 250 mM imidazole in 20 mM sodium dihydrogen phosphate,

0.5 M sodium chloride, pH 7.0 from in 9 mL at 0.5 mL/min. The product fraction

(eluting from 5.6 to 10.2 mL) was pooled, concentrated and dialysed into water. The

pooled fractions were further purified using a strong cation exchange column (CEX, SP-

5PW, Supelco) using a gradient from 320 to 450 mM sodium chloride in 20 mM sodium

dihydrogen phosphate, pH 7.0 in 14.4 mL at 1 mL/min. The product fraction (eluting

from 12.5 to 14 mL) was pooled, concentrated and dialysed into water to yield

bioconjugate cyt c-10-BSA (0.0340 mol, 30%). MS (MALDI) m/z: 80 609 ([M – 2Cl]+

requires 80 618).

8.5.4 cyt c-10-cyt c dimer



94 mM ethylenediaminetetraacetic acid, pH 7.0 (665 L) in water (805 L) at room


(1.5 mL) was then added and the resulting solution stirred in darkness at room



Healthcare) using a gradient from 0 to 250 mM imidazole in 20 mM sodium dihydrogen


8-23

phosphate, 0.5 M sodium chloride, pH 7.0 in 9 mL at 0.5 mL/min. The product fraction

(eluting from 6 to 10 mL) was pooled, concentrated and dialysed into water to yield

bioconjugate cyt c-10-cyt c (1.90 nmol, 1%). MS (MALDI) m/z: 26 551 ([M – 2Cl]+

requires 26 550).

8.5.5 cyt c-16-GFP dimer

A solution of 4,4’-bipyridinium-N,N-di(maleimidopropyl) hexaflurophosphate 16

(0.78 mg, 1.08 mol) in acetonitrile (344 L) was added to a solution of 94 mM sodium

dihydrogen phosphate, 94 mM ethylenediaminetetraacetic acid, pH 7.0 (1.46 mL) in

water (4.55 mL). Acropora millepora green fluorescent protein (amFP497)4 (1.79 mg,

0.0688 mol) was added at room temperature in darkness and stirred for 3 h. The

reaction mixture was then concentrated to ca. 1 mL and desalted by size exclusion

chromatography (SEC, HiTrapTM desalting, GE Healthcare) using 20 mM sodium

dihydrogen phosphate, pH 7.0 at 0.5 mL/min. The product fraction (eluting from 1.5 to

3.1 mL) was pooled and concentrated (527 L). Purified, reduced iso-1 cytochrome c

(3.68 mg, 0.290 mol) in water (264 L) and a solution of 94 mM sodium dihydrogen

phosphate, 94 mM ethylenediaminetetraacetic acid, pH 7.0 (213 L) was then added to

the desalted product fraction and the resulting solution stirred in darkness at room



Healthcare) using a gradient from 0 to 250 mM imidazole in 20 mM sodium dihydrogen

phosphate, 0.5 M sodium chloride, pH 7.0 in 6 mL at 0.5 mL/min. The product fraction

(eluting from 5.5 to 8.2 mL) was pooled, concentrated and dialysed into water to yield

bioconjugate cyt c-16-GFP (0.49 nmol, 0.1%). MS (MALDI) m/z: 40 245 ([M – 2PF6]+

requires 40 236).


8-24

8.5.6 BSA-10-BSA dimer



94 mM ethylenediaminetetraacetic acid, pH 7.0 (723 L) in water (2.50 mL) at room

temperature. Bovine serum albumin (90.9 mg, 1.36 mol) was then added and the

resulting solution stirred in darkness at room temperature for 23 h. The reaction mixture

was then concentrated, dialysed into water and purified by immobilised metal affinity

chromatography (IMAC, HisTrapTM HP, GE Healthcare) using a gradient from 0 to

125 mM imidazole in 20 mM sodium dihydrogen phosphate, 0.5 M sodium chloride,

pH 7.0 in 6 mL at 0.5 mL/min. The product fraction (eluting from 5.7 to 8.1 mL) was

pooled, concentrated and dialysed into water to yield bioconjugate BSA-10-BSA

(0.43 nmol, 1%). Conjugate characterised by reduced SDS-PAGE (NuPage®, 12% Bis-

Tris Gel, Invitrogen) gel electrophoresis.

8.5.7 16-GFP

A solution of 4,4’-bipyridinium-N,N-di(maleimidopropyl) hexaflurophosphate 16

(0.590 mg, 0.817 mol) in acetonitrile (344 L) was added to a solution of 94 mM

sodium dihydrogen phosphate, 94 mM ethylenediaminetetraacetic acid, pH 7.0

(1.46 mL) in water (4.26 mL). Acropora millepora green fluorescent protein

(amFP497)4 (1.79 mg, 0.0688 mol) was added at room temperature in darkness and

stirred for 19 h. Excess 2-mercaptoethanol (14 M, 1 L) was added to the reaction

mixture to quench unreacted maleimide and allowed to stir for 1 h in the dark at room

temperature. The reaction mixture was then concentrated to ca. 1 mL and purified by

immobilised metal affinity chromatography (IMAC, HisTrapTM HP, GE Healthcare)

using an imidazole gradient from 0 to 500 mM in 20 mM sodium dihydrogen phosphate,


8-25

0.5 M sodium chloride, pH 7.0 in 16 mL at 0.5 mL/min. The product fraction (eluting

from 10.6 to 14.5 mL) was pooled, concentrated and dialysed into water to yield

bioconjugate 16-GFP (7.74 nmol, 11%). MS (MALDI) m/z: 27 246 ([M – 2PF6]+

requires 27 239).

8.5.8 12-GFP

A solution of anthraquinone 12 (0.567 mg, crude) in N,N-dimethylformamide (353 L)

was added to a solution of 200 mM sodium bicarbonate, pH 8.3 (1.87 mL) in water

(44 L). Acropora millepora green fluorescent protein (amFP497)4 (2.00 mg,

0.0746 mol) was added at room temperature in darkness and stirred for 16.5 h. The

reaction mixture was then concentrated to ca. 1 mL and desalted by size exclusion

chromatography (SEC, HiTrapTM desalting, GE Healthcare) using 20 mM sodium

dihydrogen phosphate, pH 7.0 at 0.5 mL/min. The product fraction (eluting from 1.7 to

3.45 mL) was pooled, concentrated and dialysed into water to yield bioconjugate

12-GFP (20.1 nmol, 27%). MS (MALDI) m/z: 27 037 ([M]+ requires 27 041, singly

labelled) and 27 282 ([M]+ requires 27 275, doubly labelled).

8.5.9 17-GFP

A solution of N-propargyl maleimide 17 (0.068 mg, 0.502 mol) in

N,N-dimethylformamide (251 L) was added to a solution of 94 mM sodium dihydrogen

phosphate, 94 mM ethylenediaminetetraacetic acid, pH 7.0 (1.07 mL) in water (203 L).

Acropora millepora green fluorescent protein (amFP497)4 (1.346 mg, 0.0502 mol) was

added at room temperature in darkness and stirred for 16.5 h. The reaction mixture was

then concentrated to ca. 1 mL and desalted by size exclusion chromatography (SEC,

HiTrapTM desalting, GE Healthcare) using 20 mM sodium dihydrogen phosphate, pH 7.0


8-26

at 0.5 mL/min. The product fraction (eluting from 1.55 to 2.55 mL) was pooled,

concentrated and dialysed into water to yield bioconjugate 17-GFP (24.9 nmol, 50%).

MS (MALDI) m/z: 26 967 ([M + Na]+ requires 26 965).

8.5.10 Attempted ‘click’ synthesis of 13-triazole-GFP

A solution of tetrakis(acetonitrile)copper(I) hexafluorophosphate (0.146 mg,

0.392 mol) and anthraquinone azide 13 (0.100 mg, 0.299 mol) in

N,N-dimethylformamide (98 L) was added to 17-GFP (0.671 mg, 0.0249 mol) in

100 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 7.0 (393 L)

and water (925 L). Reaction mixture allowed to stir for 19 h in the dark with a nitrogen

overlay at 32 oC. Size exclusion chromatography and UV-Vis analysis did not show

evidence of formation of 13-triazole-GFP.

8.5.11 cyt c-Oregon Green 488

A solution of Oregon Green® 488 maleimide (0.46 mg, 0.99 mol) in

dimethylformamide (50 L) was mixed with iso-1 cytochrome c (1.3 mg, 1.0 mol) in

20 mM sodium dihydrogen phosphate, 20 mM ethylenediaminetetraacetic acid, 5%

dimethylformamide, pH 7.0 (1 mL). The mixture was stirred at room temperature for

21.5 h and concentrated. The conjugate was then purified in 20 mM sodium dihydrogen

phosphate buffer, pH 7.0 using size-exclusion chromatography (SEC, HiTrapTM

desalting, GE Healthcare), concentrated and dialysed extensively into water (molecular

weight cut-off 3 000 Da) yielding conjugate cyt c-Oregon Green 488 (13 nmol, 13%).

MS (MALDI) m/z 13 173 ([M + H]+ requires 13 169).


8-27

8.5.12 Cytochrome c Oxidase-Oregon Green 488

A solution of Oregon Green® 488 carboxylic acid, succinimidyl ester *5-isomer*

(0.12 mg, 0.236 mol) in N,N-dimethylformamide (50 L) was mixed with

cytochrome c oxidase (4.04 mg, 0.0202 mol) in 50 mM sodium bicarbonate, pH 8.3

(950 L). The mixture was stirred at room temperature for 160 min and concentrated.

The conjugate was then purified in 20 mM sodium dihydrogen phosphate buffer, pH 7.0

using size-exclusion chromatography (SEC, HiTrapTM desalting, GE Healthcare),

concentrated and dialysed extensively into water (molecular weight cut-off 3 000 Da)

yielding cytochrome c oxidase-Oregon Green 488. The concentration of dye labelled

cytochrome c oxidase was not determined.

8.6 Vesicle Formation and Encapsulation

8.6.1 Phospholipid vesicles (liposomes)

8.6.1.1 Preparation of large unilamellar vesicles

Liposomes were prepared using the film hydration method. In a typical experiment, an

organic solution (7 mg, 350 L) of egg phosphatidylcholine (PC) or 1-palmitoyl-2-

oleoyl-sn-glycero-3-phosphocholine (POPC) from a stock solution in chloroform

(20 mg/mL) was added to a 5 mL round bottom flask. Chloroform was evaporated

under a nitrogen stream and residual solvent was removed under high vacuum for 4 h.

The lipid was then dispersed at room temperature by adding 20 mM sodium dihydrogen

phosphate buffer, pH 7.0 (470 L) to a final lipid concentration of 15 mg/mL and

hydrated for 1 h with regular vortexing. To prepare unilamellar vesicles from the

resulting multilamellar liposome suspension, liposomes were treated according to the

methods below.


8-28

Extrusion method

Liposomal suspension (15 mg/mL) was extruded by passing the sample ten times15

through a 100 nm pore (single or double stacked) polycarbonate filters at room

temperature to size and homogenise the vesicle aggregates.15-16

Freeze-thaw method

Liposomal suspension (15 mg/mL) was subjected to five cycles of freezing (liquid

nitrogen) and thawing (40 oC water bath) followed by passing the sample ten times

through a 100 nm pore (single or double stacked) polycarbonate filter at room

temperature to size and homogenise the vesicle aggregates.17

Dehydration-rehydration method

Liposomal suspension (15 mg/mL) was freeze-dried, rehydrated in water for 1 h with

regular vortexing to a final concentration of 15 mg/mL at room temperature. Sample

was then extruded by passing the sample ten times through a 100 nm pore (single or

double stacked) polycarbonate filter at room temperature to size and homogenise the

vesicle aggregates.18

8.6.1.2 Preparation of enzyme encapsulated lipid vesicles

In a typical experiment, using the method described in Chapter 8.6.1.1, enzymes were

loaded into vesicles by rehydration of the dried lipid by adding cytochrome c (100 M)

or cyt c-Oregon Green 488 (5 M) in 20 mM sodium dihydrogen phosphate buffer,

pH 7.0. Suspension was freeze-dried, rehydrated and non-entrapped enzyme was

removed by size exclusion chromatography (SEC, Superdex 75, GE Healthcare) in

20 mM sodium dihydrogen phosphate buffer, pH 7.0 at 0.5 mL/min. The liposome


8-29

containing fractions eluted from 5.0 to 7.0 mL. To determine encapsulation efficiency

(EE%), liposomes were treated with Triton X-100 (2%, v/v) with a sample to surfactant

ratio of 1:1 (v/v).

8.6.1.3 Dynamic light scattering

The average hydrodynamic diameter (Dh) and size distribution of the prepared

aggregates in an aqueous solution (1 mg/mL) was measured using a Malvern

ZetasizerNano ZS instrument equipped with a 4 mV He-Ne laser operating at

= 632 nm, an avalanche photodiode detector with high quantum efficiency, and an

ALV/LSE-5003 multiple- digital correlator electronics system. Phospholipid based

aggregates were filtered to remove dust by extrusion using a (0.1 m) polycarbonate

membrane prior to measurement. Three measurements were made after stabilisation of

the temperature at 25 oC and the average Dh of the aggregates in these runs were

calculated.

8.6.1.4 Liposome staining for electron microscopy

Vesicles were diluted to a concentration of 1 mg/mL for transmission electron

microscopy (TEM) and measurements were performed within 1 day of preparation. A

droplet of liposomes (20 L) were applied to a formvar-coated copper grid for 1 min

and excess sample was blotted away with filter paper. Liposomes were subsequently

negatively stained according to the methods below.

2% Phosphotungstic acid

Aqueous phosphotungstic acid (2%, 20 L) was applied to the TEM grid for 30 s and

excess stain was blotted away.


8-30

2% Uranyl acetate

Aqueous uranyl acetate (2%, 20 L) was applied to the TEM grid for 45 s and excess

stain was blotted away.

8.6.2 Polymer vesicles (polymersomes)

8.6.2.1 Slow addition

Ultrapure water (0.3 mL) was added dropwise (16 L per 10 min) over a period of 3 h

to a solution of polystyrene140-b-poly(acrylic acid)48 (10 mg) dissolved in either

dioxane or tetrahydrofuran (1 mL), a common solvent for both blocks of the copolymer,

at room temperature. A turbid solution formed and was dialysed extensively (molecular

weight cut-off 4 000 Da) into water to remove dioxane/tetrahydrofuran.

8.6.2.2 Injection method

A solution of 1 mg/mL of polystyrene140-b-poly(acrylic acid)48 dissolved in dioxane or

tetrahydrofuran was injected into water at room temperature or 40 oC with continuous

vortexing to a final solvent to water ratio of either 1:2, 1:6, 1:10 (v/v) and allowed to

equilibrate for 2 days.

8.6.2.3 Enzyme induced polymersomes

In a typical experiment, a 1 mg/mL solution of PS140-b-PAA48 in tetrahydrofuran

(33 l) was injected into an enzyme solution (5 M, 200 L) in sodium dihydrogen

phosphate buffer (20 mM, pH 7.0) or phosphate buffered saline (150 mM, pH 7.2). The

solution was allowed to equilibrate for at least 24 h and extensively dialysed against

water using a 50 kDa molecular weight cut-off membrane over 24 h to remove non-

encapsulated enzymes.


8-31

8.6.2.4 Proton pumping proteo-polymersomes

In a typical experiment, a 1 mg/mL solution of PS140-b-PAA48 in tetrahydrofuran

(33 L) was injected into a 8-cyt c (7.5 M), CcOx (0.75 M),

8-hydroxypyrene-1,3,6-trisulfonic acid (5 M) solution (200 L) in phosphate buffered

saline (PBS, 150 mM, pH 7.5) at 25 oC. The buffered enzyme solution was allowed to

equilibrate for 10 min prior to PS140-b-PAA48 in tetrahydrofuran injection. After

injection, the solution was allowed to equilibrate for at least 24 h and extensively

dialysed into ultrapure water using a 50 kDa molecular weight cut-off membrane over

24 h to remove non-encapsulated enzymes and fluorescent dye at 25 oC. This also

removes both the external and internally encapsulated 150 mM PBS salt buffer from the

proteo-polymersome sample. The concentration of 8-cyt c loaded proteo-polymersomes

was estimated by UV-Vis absorbance ( 410 = 97.6 mM-1cm-1).1b

8.7 Enzyme Activity and Photoreaction Experiments

8.7.1 General photo-induced light reaction equipment

Ruthenium(II)-protein biohybrid photo-reactions were performed using a 16 LED Blue

(465 nm) Flashlight (LDP LLC) and Iridium(III)-protein photoreactions were performed

using a 16 LED White (372 nm) Flashlight (LDP LLC). Power measurements of LED

Flashlights were made using a Newport Power Meter (Model 1918-C). Where indicated,

samples were exposed to light (equipped with a UV filter) from an Oriel Basic Power

Supply (model 68806, 50 – 200 watts) fitted with a xenon arc lamp. amGFP photo-

conversion reactions were made using a custom built LED source consisting of an array

of 3 Cyan Lambertian Luxeon-VTM diodes (Phillips, LXHL-LE5C) emitting in a

490±30 nm band connected to an electrical circuit that was fed a power supply at 2.5 A

(30 W). The array was water-cooled with a tightly attached hard disk drive heat sink.


8-32

8.7.2 Cytochrome c/Cytochrome c oxidase studies

8.7.2.1 Measurement of catalytic activity by ABTS assay

The catalytic activity of yeast iso-1 cytochrome c was measured using 2,2’-azino-bis(3-

ethylbenzothiazoline-6-sulphonic acid) (ABTS). A stock solution of 100 mM hydrogen

peroxide was prepared by dilution of 9.4 L of 30% hydrogen peroxide to 1 mL with

ultra pure water. A second stock solution of 20 mM ABTS was prepared by dissolution

of 6.9 mg ABTS in 629 L of 20 mM sodium dihydrogen phosphate, pH 7.0. Activity of

solution (bulk) cytochrome c and encapsulated cytochrome c (PS140-b-PAA48) were

measured at target concentrations of 200 M ABTS, 0.5 M cytochrome c/encapsulated

cytochrome c and 1 mM hydrogen peroxide by adding 1 L of 20 mM ABTS, 54 L of

0.93 M cytochrome c/encapsulated cytochrome c to 44 L of 20 mM sodium

dihydrogen phosphate, pH 7.0 and initiating the reaction with 1 L of 1 mM hydrogen

peroxide. The reactions were monitored by increasing UV absorbance at 415 nm.19

8.7.2.2 Measurement of biological activity by CcOx assay

Biological activity was measured using horse heart cytochrome c oxidase (CcOx). A

stock solution of cytochrome c oxidase was prepared at 1 mg/mL in 20 mM sodium

dihydrogen phosphate buffer, pH 7.0. A catalytic amount (0.25 M) of cytochrome c

oxidase was added to 2.3 M reduced cytochrome c or bioconjugates (bulk and

encapsulated) in 5 mM sodium dihydrogen phosphate, 5 mM ethylenediaminetetraacetic

acid (EDTA), pH 7.0 by twenty-fold dilution and oxidation of cytochrome c was

monitored by decreasing UV absorbance at 550 nm.


8-33

8.7.2.3 Photo-induced electron transfer studies

Room temperature photo-induced electron transfer measurements of a non-covalent

mixture of complex 6 and cytochrome c and 8-cyt c bioconjugate were conducted in

specialised small volume quartz cuvettes designed for protein samples, allowing

complete exposure to irradiation with a constant area for all experiments

(1.0 cm × 0.3 cm). A solution (80 L) of 5 mM sodium dihydrogen phosphate buffer,

5 mM ethylenediaminetetraacetic acid, pH 7.0 was prepared containing either a 1:1

mixture of cytochrome c (2.3 M) and complex 6 (2.3±0.1 M) or bioconjugate (2.3 M)

in bulk or membrane encapsulated (see Chapter 8.6.2.3) samples. Prior to irradiation,

cuvettes were degassed for 30 min at 0 oC under reduced pressure (120 mbar) and

overlayed with nitrogen in the dark. Samples were irradiated in a nitrogen purged

UV-Vis spectrometer with a 465 nm light (LED) source, placed 2.5 cm from sample

and cytochrome c reduction was monitored by UV absorbance at 550 nm.

8.7.2.4 Photo-induced proton pumping studies

In a typical experiment, a solution of salt-free proteo-polymersomes (see Chapter

8.6.2.4) (1.1±0.1 M 8-cyt c, 113.6 L) was adjusted to a final buffered condition of

5 mM NaH2PO4 and 5 mM EDTA by addition of a stock solution of 94 mM NaH2PO4

and 94 mM EDTA (6.4 L) at a desired pH. The sample was degassed for 20 min at

0 oC under reduced pressure (120 mbar) and overlayed with nitrogen in the dark to

remove oxygen quenching the excited state of Ru(II)-bisterpyridine complex as well as

allowing the equilibration of the buffer in the internal proteo-polymersome

compartment due to porosity of the membrane. A 465 nm light (LED) source was

placed 2.5 cm from the sample over a 1.0 cm × 0.3 cm sample area and irradiated in a

nitrogen purged fluorescence spectrometer. Fluorescence emission measurements were


8-34

monitored over 2 min intervals with an ex = 460 nm and em = 510 nm. Fluorescence

excitation measurements were made with a em = 510 nm. Excitation and emission slit

widths were 5 nm. After irradiation, a drop of aqueous sodium hydroxide (2 L, 1 M)

was added to determine the maximum emission intensity (pH > 9) to calibrate the

8-hydroxypyrene-1,3,6-trisulfonic acid pH titration curve (see Chapter 7).

8.7.2.5 8-hydroxypyrene-1,3,6-trisulfonic acid pH titration

Bulk 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS) (480 M, 3 L) and polymersome

encapsulated HPTS (200 M in lumen, 3 L) (see Chapter 8.6.2.3) was diluted into

20 mM sodium dihydrogen phosphate buffer (3 mL) at a desired pH value and pH of

phosphate buffers were adjusted using aqueous sodium hydroxide (1 M) or hydrochloric

acid (1 M). Titrations were measured from acidic to basic pH and fluorescence emission

intensities were measured with ex = 460 nm and em = 510 nm.

8.7.2.6 Nitrite reductase activity of cytochrome c

Equine heart cytochrome c (1.06 mM, 189 L) was incubated with 100 nm 1-palmitoyl-

2-oleoyl-sn-glycero-3-phosphocholine (POPC) vesicles (15 mg/mL, 100 L) and

bis(4’-(4-hydroxymethylphenyl)-2,2’:6’:2”-terpyridine)iridium(III)tris(hexafluorophosp

hate) (6 mM, 33 L) in 25 mM sodium dihydrogen phosphate, 5 mM

ethylenediaminetetraacetic acid, pH 5.4 (450 L) under reduced pressure (120 mbar) for

10 min. Sodium nitrite (100 M, 50 L) in 25 mM sodium dihydrogen phosphate, 5 mM

ethylenediaminetetraacetic acid, pH 5.4 was subsequently added and sample irradiated

using a 372 nm light (LED) from a distance of 2.0 cm at room temperature for 3 h

followed by incubation for a further 13 h 45 min under anaerobic conditions

(120 mbar). Solution centrifuged at 13 800 rpm for 30 min using a 3 000 molecular


8-35

weight cut-off centrifuge concentrator and filtrate analysed using the Griess assay

(Griess reagent to sample, 1:1, v/v).

8.7.2.7 Griess Assay

To an aqueous solution containing nitrite (100 L), saturated aqueous solution (50 L)

of sulphanilamide (10 mM) in 20 mM sodium dihydrogen phosphate buffer, pH 7.0 and

orthophosphoric acid (1%, v/v) was added. An aqueous solution (50 L) of

N-(1-Napthyl)ethylenediamine dihydrochloride (400 M) in 20 mM sodium dihydrogen

phosphate buffer, pH 7.0 was added to the mixture and incubated for 10 min. Assay was

monitored by measuring UV absorbance at 540 nm and nitrite concentration was

estimated based on a nitrite calibration curve (see Appendix C).

8.7.3 Green fluorescent protein studies

8.7.3.1 Time-resolved laser spectroscopy

Room temperature fluorescence measurements were performed in the laboratory of Dr.

Timothy W. Schmidt at the University of Sydney with Dr. Raphaël G. C. R. Clady and

Mr. Murad Tayebjee. Time-resolved fluorescence lifetime measurements were recorded

by ultrafast (femtosecond) photoexcitation of sample cuvette (non-degassed) using a

tuneable output TOPAS OPA laser (476 nm) pumped by a Clark-MXR femtosecond

laser operating at 1 kHz. The ~1 mm2 fluorescent spot on the front face of the cuvette

was entirely imaged, with a lens, through the slits of a spectrograph and detected with

an iCCD camera (Acton/Princeton) synchronised to the laser output. The kinetics of

delayed fluorescence were measured in 5 ps slices from 5 to 2180 ps delay for ultrafast

time-resolved photoluminescence and in 50 ps slices for time-correlated single photon


8-36

counting. All spectra are baseline corrected. Fitting of the delayed fluorescence signal

was performed with OriginPro 8.

8.7.3.2 Xenon arc lamp irradiation studies

Room temperature photo-induced electron transfer measurements on a 16-GFP

bioconjugate were conducted in a specialised small volume quartz cuvette. A solution

(80 L) of 20 mM sodium dihydrogen phosphate buffer, 20 mM

ethylenediaminetetraacetic acid, pH 7.0 was prepared containing bioconjugate 16-GFP

(13 M). Prior to irradiation, cuvettes were degassed for 30 min at 0 oC under reduced

pressure (120 mbar) and overlayed with nitrogen in the dark. Samples were irradiated

using a xenon lamp fitted with a 2 mm iris from a distance of 10 cm. Samples were

irradiated for 30 min and monitored by UV-Vis spectroscopy at 600 nm.

8.7.3.3 Cyan LED irradiation studies

A solution (120 L) of a non-covalent mixture of green fluorescent protein (20 M) and

4,4’-bipyridinium-N,N-di(maleimidopropyl) hexafluorophosphate, p-benzoquinone,

anthraquinone-2-carboxylic acid (1mM) or Anthraquinone-GFP (12-GFP, 18 M) in

20 mM sodium dihydrogen phosphate buffer, pH 7.0 were irradiated using a custom

built 3 × 490 nm LED source for 3 h at a distance of 2.0 cm. Photoconversion was

followed by UV-Vis and fluorescence (ca. 1.6 M) spectroscopy post-irradiation.

8.8 References

(1) (a) Peterson, J. R.; Smith, T. A.; Thordarson, P. Org. Biomol. Chem. 2010, 8,151. (b) Peterson, J. R.; Thordarson, P. Chiang Mai J. Sci. 2009, 26, 236.

(2) Hvasanov, D.; Wiedenmann, J.; Braet, F.; Thordarson, P. Chem. Commun. 2011,47, 6314.


8-37

(3) (a) Iancu, C. V.; Tivol, W. F.; Schooler, J. B.; Dias, D. P.; Henderson, G. P.; Murphy, G. E.; Wright, E. R.; Li, Z.; Yu, Z.; Briegel, A.; Gan, L.; He, Y.; Jensen, G. J. Nat. Protocols 2007, 1, 2813. (b) Frederik, P. M.; Hubert, D. H. W. Methods Enzymol. 2005, 391, 431.

(4) Angelo, C.; Denzel, A.; Vogt, A.; Matz, M. V.; Oswald, F.; Salih, A.; Nienhaus, G. U.; Wiedenmann, J. Mar. Ecol. Prog. Ser. 2008, 364, 97.

(5) Wiedenmann, J.; Schenk, A.; Röcker, C.; Girod, A.; Spindler, K.-D.; Nienhaus, G. U. Proc. Natl. Acad. Soc. U. S. A. 2002, 99, 11646.

(6) Bruker APEX2 Suite; Bruker AXS Inc.: Madison, WI, USA, 2007.(7) Sheldrick, G. Acta Crystallogr. Sect. A: Found. Crystallogr. 2008, 64, 112.(8) Kabsch, W. J. Appl. Crystallogr. 1993, 26, 795.(9) Mukkala, V.-M.; Helenius, M.; Hemmilä, I.; Kankare, J.; Takalo, H. Helv.

Chim. Acta 1993, 76, 1361.(10) Mikel, C.; Potvin, P. G. Polyhedron 2002, 21, 49.(11) (a) Lainé, P.; Bedioui, F.; Ochsenbein, P.; Marvaud, V.; Bonin, M.; Amouyal, E.

J. Am. Chem. Soc. 2002, 124, 1364. (b) Peterson, J. R. PhD Thesis, The University of Sydney, 2009.

(12) (a) Sullivan, B. P.; Calvert, J. M.; Meyer, T. J. Inorg. Chem. 1980, 19, 1404. (b) Hofmeier, H.; Andres, P. R.; Hoogenboom, R.; Herdtweck, E.; Schubert, U. S. Aust. J. Chem. 2004, 57, 419.

(13) Ng, W. Y.; Gong, X.; Chan, W. K. Chem. Mater. 1999, 11, 1165.(14) Lewis, W. G.; Magallon, F. G.; Fokin, V. V.; Finn, M. G. J. Am. Chem. Soc.

2004, 126, 9152.(15) Mui, B.; Chow, L.; Hope, M. J. Methods Enzymol. 2003, 367, 3.(16) Das, N.; Gupta, S.; Mazumdar, S. Biochem. Biophys. Res. Commun. 2001, 286,

311.(17) (a) Walde, P.; Marzetta, B. Biotechnol. Bioeng. 1998, 57, 216. (b) Blocher, M.;

Walde, P.; Dunn, I. J. Biotechnol. Bioeng. 1999, 62, 36.(18) Kirby, C.; Gregoriadis, G. Nat. Biotech. 1984, 2, 979.(19) Peterson, J. R.; Smith, T. A.; Thordarson, P. Chem. Commun. 2007, 1899.

Chapter 9

Conclusions and Future Work

Chapter 9 Conclusions and Future Work

9-2

9 Conclusions and Future Work

The development of nanoreactors and artificial cells has gained considerable interest

over the last two decades in order to better understand the origin of life.1 In a landmark

article, Szostak and co-workers have proposed three requirements for a model protocell

as (1) bearing an informational substance (DNA or RNA), (2) a catalyst and (3) a

compartment.2 However, implicitly, the requirement for energy production in a model

protocell system is essential to drive these processes. Nearly all organisms on Earth

source energy directly or indirectly from the Sun. In nature, photosynthesis in the

chloroplasts (thylakoid membrane) of plants achieve this energy production by

converting light energy from the Sun into chemical energy in the form of a

transmembrane electrochemical gradient which produces ATP.3 This Thesis describes

the development of a primitive chloroplast – a system that converts light energy into

chemical energy in the form of an electrochemical gradient ( ). Such a system can in

principle be used as a “nanoreactor battery” where the light induced proton gradient is

used to drive chemical reactions or biochemical processes.

In order to develop a primitive artificial chloroplast, a light-activated

donor-acceptor system based on a Ru(II)-bisterpyridine cytochrome c bioconjugate was

synthesised which has been reported in the Thordarson group.4 Terpyridine ligands

bearing amines with 4’-aryl functionalisation as metal complex precursors were

synthesised based on a modified Kröhnke synthesis with isolation of the

azachalcone/pyridinium iodide precursors. Light-activated chromophores of

Ru(II)-bisterpyridine complexes were prepared in ethylene glycol based on conditions

adapted from Ir(III)-bisterpyridine literature.5 Homo and heteroleptic aniline complexes

7 and 9 were synthesised to allow chemical modification by introducing maleimide

functionality using peptide coupling chemistry. Asymmetric and symmetric maleimide


9-3

complexes 8 and 10 were synthesised such that these electron donating photosensitisers

could be attached to cysteine bearing proteins/enzymes. As a reference compound,

complex 6 was prepared for control room temperature photoreduction studies with yeast

iso-1 cytochrome c (cyt c). Single crystals of complexes 7 and 8 were isolated and

analysed by X-ray crystallography to confirm their structural properties which was

conducted by Dr. Mohan Bhadbhade (UNSW) with analysis of aniline 7 performed

in-house at the UNSW Analytical Centre and maleimide 8 at the Australian Synchotron

facility. At the time of writing, complex 8 is the first reported single crystal X-ray

structure of a Ru(II)-bistepyridine complex with maleimide functionality.

The synthesis of a light-activated donor-acceptor system based on an

asymmetric terpyridine cytochrome c bioconjugate 8-cyt c was prepared for electron

transfer studies in membranes and for use as a component in a polymer membrane

reconstituted electron transfer chain as a photosynthetic-respiratory hybrid. The

synthesis of 8-cyt c was achieved by reacting maleimide functionalised asymmetric

complex 8 with the single cysteine residue (CYS102) of yeast iso-1 cytochrome c under

benign conditions at physiological pH 7. The purification of Ru(II)-bisterpyridine based

bioconjugate could be achieved using immobilised metal affinity chromatography

(IMAC, Ni2+) in high purity with 6% yield following procedures previously developed

in the Thordarson group by Peterson et al.4a, although the exact mechanism remains

unknown. The limitation of low yielding asymmetric bioconjugate 8-cyt c is a result of

the poor water solubility.

As an extension of the this work, the preparation methods of high molecular

weight dimeric bioconjugates were explored in order to improve yields. Several

bioconjugate dimers have been prepared based on combinations of cytochrome c, green

fluorescent protein (Acropora millepora) and bovine serum albumin to probe factors


9-4

affecting dimerisation yield. The role and importance of protein charge to analyse

protein function after amino acid modification has been employed in the literature, such

as the use of protein charge ladders.6 However, the exploitation of global protein charge

to facilitate dimer bioconjugate synthesis via supramolecular interactions has been

neglected. It was found that a high molecular weight heterodimer of complementary

charge cyt c-10-BSA (>80 kDa) was prepared in up to 30% yield. Additionally, it is

noteworthy that like-charged homo and heterodimers were prepared in extremely low

yields of less than 1%. Based on these studies, it is proposed that global protein charge

can be utilised to induce higher dimer yields by electrostatic supramolecular ionic

attraction. However, the absolute purification of dimers proved to be difficult due to the

low yields of the dimers. The bioconjugates were characterised by MALDI-TOF mass

spectrometry, UV-Vis spectroscopy and gel electrophoresis.

Following the work of synthetic donor-biological acceptor systems based on

Ru(II)-cyt c, the alternative biological donor-synthetic acceptor construct was

investigated using green fluorescent protein (GFP) for light-induced electron transfer

studies. Anthraquinone precursors were prepared to function as electron acceptors after

functionalising to the light-activated biological electron donor, GFP. Azide bearing

anthraquinone 13 was prepared for potential Cu(I)-catalysed [3+2] Huisgen

cycloaddition reactions (click reactions) of azides with terminal alkynes to introduce

maleimide functionality and subsequently site-specifically modify GFP at the single

CYS119 residue. However, attempts to functionalise azide 13 with maleimides using

click chemistry proved to be unsuccessful. Instead, the N-hydroxysuccinimide ester of

anthraquinone-2-carboxylic acid (anthraquinone 12) was prepared to non-specifically

modify GFP via amine residues. Electron transfer studies were conducted using

viologen-GFP (16-GFP) and anthraquinone-GFP (12-GFP) bioconjugates, which were


9-5

prepared via cysteine-maleimide coupling (reacting to single CYS119 residue) and

amine modification (non-specific), respectively.

The purification of the resulting bioconjugates was achieved using Ni2+-IMAC

chromatography, resulting in yields of 11% and 27%, respectively. Based on

steady-state UV-Vis and fluorescence spectroscopy studies, evidence of electron

transfer was observed by red photoconversion of the chromophore with p-benzoquinone

and anthraquinone acceptor molecules as a non-covalent mixture. To further confirm

electron/energy transfer processes of non-covalent and bioconjugate GFP mixtures,

time-resolved fluorescence spectroscopy measurements were conducted. Lifetime

measurements showed a GFP fluorescence lifetime of 1.65±0.06 ns. Furthermore, the

fluorescence lifetimes of GFP were quenched to 0.14±0.01 and 0.23±0.04 ns in the

presence of excess (non-covalent) p-benzoquinone and anthraquinone-2-carboxylic

acid, respectively, corresponding to a rate constant of forward electron transfer of

ket = 6.5±0.5 × 109 and 3.7±0.7 × 109 s-1. The donor-acceptor viologen bioconjugate

16-GFP showed no evidence of photoinduced electron transfer as viologen acts as a

poor electron acceptor in 16-GFP, which is consistent with non-covalent steady state

UV-Vis and fluorescence spectroscopy measurements. In contrast, fluorescence lifetime

studies of the anthraquinone bioconjugate 12-GFP demonstrated light-induced

electron/energy transfer processes, with a quenched GFP fluorescence lifetime of

0.28±0.04 ns and an electron transfer rate of ket = 2.9±0.4 × 109 s-1.

Vesicles based on phospholipids and diblock copolymers were investigated to

evaluate their feasibility as compartments for the encapsulation of donor-acceptor

8-cyt c. The first compartment candidate was based on lipid vesicles (liposomes). Lipid

vesicles were formed from either naturally sourced egg L- -phosphatidylcholine (egg

PC) or synthetically equivalent L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine


9-6

(POPC), using the film hydration method and sized via extrusion to produce

monodisperse unilamellar vesicles of 100 nm in diameter. The validation of

monodispersity and average diameter was determined using dynamic light scattering

techniques. Additionally, unilamellar vesicles were characterised using traditional TEM

microscopy and cryo-TEM techniques and it was found that structural characterisation

of lamellar features using conventional TEM microscopy introduced artificacts as a

result of the necessity of negative stains and drying of specimen. The possibility of

liposomes based on egg PC and POPC as potential compartments were determined by

the encapsulation of model protein cytochrome c, which is used as the redox

metalloprotein for light-induced electron transfer. Encapsulation was confirmed using

confocal laser-scanning microscopy (CLSM) of fluorescently labelled cytochrome c.

However, low encapsulation efficiencies of 1.5±0.1% prevented their further use for

bioconjugate encapsulation studies.

Polymersomes were also formed as potential compartments as an alternative

candidate, using the polyelectrolyte diblock copolymer, polystyrene140-b-poly(acrylic

acid)48 (PS140-b-PAA48). A novel method of polymersome preparation was

demonstrated using the syringe injection method which induces polymersome

aggregates in a facile and biologically friendly manner. Concomitantly, encapsulation of

positively charged biomolecules occurs within the coronal block. It is proposed that the

formation of polymersomes is in a 2:1 polymer:protein ratio which was investigated by

determining the relative ratio of polymersome/micelle formation as a function of

positive charge (enzyme) composition. Additionally, this induced formation is versatile

and can be applied to various positively charged biomolecules with high encapsulation

efficiencies allowing for potential applications as nanoreactors7 or protein therapeutic

vessels including use of polycationic copolymers as non-viral vectors for DNA/RNA


9-7

gene therapy8 and simplified preparation compared to polyelectrolyte microcapsules.9

Polymersomes were characterised by conventional and cryo-TEM microscopy as well

as CLSM. The encapsulated enzymes were demonstrated to remain biologically active

after encapsulation in the presence of organic solvent (tetrahydrofuran), using

spectroscopy studies and the ABTS assay. Due to the high encapsulation efficiencies of

55±1% and 66±7% in 20 mM NaH2PO4 and PBS, respectively, for cytochrome c and

35±1% for GFP, polymersomes were selected as the most suitable compartments for

enzyme encapsulation.

Using the formation of PS140-b-PAA48 polymersomes which allows the

encapsulation of positively charged enzymes within the coronal block of the membrane,

photoinduced electron transfer studies were performed using 8-cyt c. Initially, the

photophysical properties of bulk Ru(II)-cyt c bioconjugate 8-cyt c was investigated. The

photoreduction of 8-cyt c could be followed by monitoring the increase in the 550 nm

absorption band corresponding to heme reduction using steady-state UV-Vis absorption

spectroscopy measurements. It was observed that Ru(II) 8-cyt c could be fully reduced

in 50 min via irradiation with a 465 nm LED light source. The quantum efficiency ( )

of bioconjugate 8-cyt c in bulk solution was estimated to be 5.9±1.5 10-4%.

Subsequently, in order to determine the effect of membrane encapsulation for use as a

component in a semi-synthetic electron transport chain, bioconjugate 8-cyt c was

membrane encapsulated in PS140-b-PAA48. Induced 8-cyt c polymersome formation

was confirmed by TEM microscopy with an average diameter of 290±132 nm. The

encapsulated conjugate 8-cyt c, exhibited a two-fold enhancement of the initial rate of

heme reduction with an estimated of 1.1±0.3 × 10-3%. Molecular modelling of 8-cyt c

showed that electron transfer is occurring over a maximum distance between the

ruthenium and heme centre of 32 Å.


9-8

Expanding on photoreduction of cytochrome c, electron transfer studies between

a non-covalent mixture of Ir(III)-bisterpyridine complex 18 and horse heart

cytochrome c using a 372 nm UV light source was investigated to induce nitrite

reductase mimicry behaviour. It was shown that in the presence of zwitterionic

phospholipid (POPC) liposomes (100 nm), a pentacoordinate heme state could be

induced allowing reactions with nitrite anions to form nitric oxide. The photo-activated

nitrite reductase activity of cytochrome c exhibited an 82% conversion of nitrite anions

into nitric oxide after irradiation under anaerobic conditions in an acidic phosphate

buffered environment (pH 5.4). Due to experimental limitations, the exact irradiation

power could not be determined due scattering and absorption by the vacuum flask and

nitrite levels could only measured at a single time point.

After investigation of the photophysical properties of bulk and membrane

encapsulated PS140-b-PAA48 polymersomes, a synthetic hybrid

photosynthetic-respiratory system that generates a proton potential upon photoexcitation

was constructed via self-assembly. Reconstitution of the hybrid enzyme cascade

consisting of bioconjugate 8-cyt c, cytochrome c oxidase in the polymersome

membrane of polyelectrolyte PS140-b-PAA48 was characterised by TEM and CLSM.

Irradiation with visible light (465 nm) converts photon energy into an electrochemical

potential via electron transfer. Photoinduced electron transfer occurs across an

artificial electron transport chain within the polymersome membrane coupled with

overall vectorial proton translocation and resulting pH changes were monitored by

steady-state fluorescence spectroscopy using internally encapsulated HPTS. Following

fluorescence spectroscopy studies, it was observed that cytochrome c oxidase appears to

have the same orientation (average >50% native orientation) in the polymersome

membrane as encountered in the mitochondria. The resulting biased orientation is


9-9

attributed to the alignment of cytochrome c oxidase with the curvature of the

polymersomes. The photosynthetic-respiratory system is capable of storing chemical

energy in the form of a transmembrane proton potential with a proton pumping rate of

3.3 × 103 H+/s at physiological pH 7.2 ( pH 0.2 units) correlating to a of

1.1±0.3 × 10-9%. Proton translocation rates were investigated using pH-dependent

studies showing that a four-fold increase in proton pumping rate is observed under

acidic conditions of pH 6.5 despite the increased proton "back-pressure".

In conclusion, this research has shown that the construction of the synthetic

photoactive electron transport chain capable of storing chemical energy as a proton

gradient brings vesicle and polymersome chemists towards satisfying one of the three

requirements2 for an artificial model protocell, which include (1) catalyst (storing of

chemical energy), (2) self-reproducing vesicles1a and (3) self-reproducing informational

substance (genetic information).1a The semi-synthetic nature of the electron transport

chain, which is reconstituted in an artificial polyelectrolyte membrane demonstrates the

construction of a robust and reproducible primitive chloroplast. This system opens

opportunities for further study including possibly utilising this system to drive

biomimetic processes such as generation of ATP10 and ATP-linked processes by

potentially reconstituting transmembrane ATP synthase or alternatively inducing

pH-sensitive reactions within the lumen.

9.1 References

(1) (a) Kurihara, K.; Tamura, M.; Shohda, K.-i.; Toyota, T.; Suzuki, K.; Sugawara, T. Nat. Chem. 2011, 3, 775. (b) Vriezema, D. M.; Garcia, P. M. L.; SanchoOltra, N.; Hatzakis, N. S.; Kuiper, S. M.; Nolte, R. J. M.; Rowan, A. E.; vanHest, J. C. M. Angew. Chem. Int. Ed. 2007, 46, 7378.

(2) Szostak, J. W.; Bartel, D. P.; Luisi, P. L. Nature 2001, 409, 387.(3) Cordes, M.; Giese, B. Chem. Soc. Rev. 2009, 38, 892.(4) (a) Peterson, J. R.; Smith, T. A.; Thordarson, P. Chem. Commun. 2007, 1899. (b)

Peterson, J. R.; Smith, T. A.; Thordarson, P. Org. Biomol. Chem. 2010, 8, 151.


9-10

(5) Hofmeier, H.; Schubert, U. S. Chem. Soc. Rev. 2004, 33, 373.(6) (a) Gitlin, I.; Carbeck, J. D.; Whitesides, G. M. Angew. Chem. Int. Ed. 2006, 45,

3022. (b) Gitlin, I.; Mayer, M.; Whitesides, G. M. J. Phys. Chem. B 2003, 107,1466.

(7) Minten, I. J.; Claessen, V. I.; Blank, K.; Rowan, A. E.; Nolte, R. J. M.; Cornelissen, J. J. L. M. Chem. Sci. 2011, 2, 358.

(8) Ropert, C. Braz. J. Med. Biol. Res. 1999, 32, 163.(9) Zelikin, A. N.; Becker, A. L.; Johnston, A. P. R.; Wark, K. L.; Turatti, F.;

Caruso, F. ACS Nano 2007, 1, 63.(10) (a) Choi, H.-J.; Montemagno, C. D. Nano Lett. 2005, 5, 2538. (b) Steinberg-

Yfrach, G.; Rigaud, J.-L.; Durantini, E. N.; Moore, A. L.; Gust, D.; Moore, T. A. Nature 1998, 392, 479.

Appendices

Appendices

Appendix A

A.1 Supplementary data for X-ray diffraction analysis of Ru(II) complex 7

Table A.1. Data collection for Ru(II) complex 7

Radiation source: fine-focus sealed tube 7849 independent reflections32252 measured reflections 5932 reflections with I > 2 (I)Graphite monochromator Rint = 0.056

scans, and scans with offsets max = 25.0o, max = 2.7o

Absorption correction: Multi-scan h = -10 10SADABS (Bruker, 2001) k = -10 10Tmin = 0.924, Tmax = 0.974 l = -33 33

Table A.2. Refinement for Ru(II) complex 7

Refinement on F2 Primary atom site location: Structure-invariant direct methods

Least-squares matrix: Full Secondary atom site location: Difference Fourier map

R[F2 > 2 (F2)] = 0.041 Hydrogen site location: Inferred from neighbouring sites

wR(F2) = 0.125 H-atom parameters constrainedS = 0.83 w = 1/[ 2(Fo

2) + (0.1P)2 + 0.7828P] where P =(Fo

2 + 2Fc2)/3

7849 reflections ( / )max = 0.002672 parameters max = 0.62 e Å-3

12 restraints min = -0.52 e Å-3

Table A.3. Fractional atomic coordinates and isotropic or equivalent isotropic displacement parameters (Å2)

x y z U iso*/Ueq Occ. (<1)P1A 0.60250 (16) 0.34843 (13) 0.85755 (4) 0.0454 (3)F1A 0.713 (3) 0.487 (2) 0.8553 (8) 0.108 (9) 0.33 (2)F2A 0.745 (3) 0.257 (3) 0.8420 (10) 0.140 (13) 0.33 (2)F3A 0.499 (3) 0.205 (2) 0.8582 (8) 0.120 (12) 0.33 (2)F4A 0.479 (4) 0.460 (3) 0.8727 (8) 0.110 (10) 0.33 (2)F5A 0.545 (3) 0.349 (3) 0.8037 (4) 0.074 (7) 0.33 (2)F6A 0.664 (4) 0.323 (5) 0.9102 (5) 0.076 (8) 0.33 (2)F1A' 0.642 (3) 0.5117 (7) 0.8666 (5) 0.162 (7) 0.67 (2)F2A' 0.7719 (7) 0.3036 (14) 0.8525 (4) 0.089 (3) 0.67 (2)F3A' 0.5591 (16) 0.1818 (6) 0.8471 (6) 0.108 (5) 0.67 (2)F4A' 0.4243 (10) 0.375 (2) 0.8606 (5) 0.127 (5) 0.67 (2)F5A' 0.5874 (14) 0.3886 (15) 0.8031 (2) 0.064 (3) 0.67 (2)F6A' 0.615 (2) 0.317 (3) 0.9119 (3) 0.091 (5) 0.67 (2)Ru1 0.78387 (3) 0.83275 (3) 0.721565 (10) 0.01946 (11)N1A 0.7035 (3) 1.0282 (3) 0.69429 (11) 0.0243 (7)N2A 0.8119 (3) 0.7998 (3) 0.65361 (11) 0.0255 (7)

Appendices

N3A 0.8671 (3) 0.6207 (3) 0.72202 (12) 0.0256 (7)C1A 0.6451 (4) 1.1439 (4) 0.71774 (15) 0.0269 (9)H1A 0.6418 1.1394 0.7510 0.032*C2A 0.5903 (5) 1.2678 (5) 0.69557 (16) 0.0360 (10)H2A 0.5479 1.3461 0.7132 0.043*C3A 0.5978 (5) 1.2766 (5) 0.64765 (17) 0.0400 (11)H3A 0.5618 1.3616 0.6316 0.048*C4A 0.6584 (5) 1.1603 (5) 0.62311 (16) 0.0384 (11)H4A 0.6645 1.1652 0.5899 0.046*C5A 0.7104 (4) 1.0367 (4) 0.64652 (14) 0.0275 (9)C6A 0.7768 (5) 0.9085 (4) 0.62331 (13) 0.0295 (9)C7A 0.8093 (6) 0.8921 (5) 0.57639 (15) 0.0447 (12)H7A 0.7853 0.9680 0.5547 0.054*C8A 0.8770 (6) 0.7638 (6) 0.56164 (17) 0.0506 (13)H8A 0.8990 0.7509 0.5295 0.061*C9A 0.9133 (5) 0.6529 (5) 0.59337 (16) 0.0450 (12)H9A 0.9609 0.5650 0.5833 0.054*C10A 0.8788 (4) 0.6729 (4) 0.63990 (14) 0.0307 (9)C11A 0.9062 (4) 0.5707 (4) 0.67858 (15) 0.0292 (9)C12A 0.9646 (5) 0.4299 (5) 0.67295 (18) 0.0404 (11)H12A 0.9911 0.3951 0.6428 0.048*C13A 0.9835 (5) 0.3413 (5) 0.71157 (19) 0.0458 (13)H13A 1.0277 0.2465 0.7083 0.055*C14A 0.9388 (5) 0.3891 (5) 0.75472 (18) 0.0403 (11)H14A 0.9468 0.3268 0.7812 0.048*C15A 0.8818 (4) 0.5306 (4) 0.75859 (16) 0.0310 (9)H15A 0.8519 0.5649 0.7884 0.037*N1B 0.9959 (3) 0.9272 (3) 0.74028 (10) 0.0192 (6)N2B 0.7655 (3) 0.8670 (3) 0.78959 (10) 0.0206 (7)N3B 0.5658 (3) 0.7491 (3) 0.73025 (11) 0.0238 (7)N4B 0.6978 (5) 0.9776 (4) 1.08262 (12) 0.0456 (10)H4B1 0.6412 0.9168 1.0985 0.055*H4B2 0.7489 1.0509 1.0973 0.055*C1B 1.1115 (4) 0.9561 (4) 0.71184 (13) 0.0224 (8)H1B 1.0993 0.9270 0.6796 0.027*C2B 1.2456 (4) 1.0258 (4) 0.72752 (14) 0.0279 (9)H2B 1.3249 1.0434 0.7066 0.034*C3B 1.2640 (5) 1.0702 (4) 0.77424 (14) 0.0314 (9)H3B 1.3544 1.1219 0.7857 0.038*C4B 1.1479 (4) 1.0379 (4) 0.80391 (13) 0.0271 (9)H4B 1.1595 1.0648 0.8363 0.033*C5B 1.0154 (4) 0.9666 (4) 0.78651 (12) 0.0195 (8)C6B 0.8845 (4) 0.9287 (4) 0.81554 (13) 0.0229 (8)C7B 0.8775 (4) 0.9458 (4) 0.86366 (13) 0.0249 (8)H7B 0.9625 0.9877 0.8817 0.030*C8B 0.7447 (5) 0.9013 (4) 0.88597 (14) 0.0271 (9)C9B 0.6238 (4) 0.8385 (4) 0.85746 (14) 0.0288 (9)H9B 0.5323 0.8070 0.8713 0.035*

Appendices

C10B 0.6358 (4) 0.8219 (4) 0.80993 (13) 0.0216 (8)C11B 0.5229 (4) 0.7531 (4) 0.77566 (14) 0.0253 (9)C12B 0.3835 (4) 0.6913 (5) 0.78767 (16) 0.0335 (10)H12B 0.3540 0.6950 0.8193 0.040*C13B 0.2894 (5) 0.6250 (5) 0.75311 (17) 0.0409 (11)H13B 0.1938 0.5824 0.7607 0.049*C14B 0.3338 (5) 0.6201 (5) 0.70739 (17) 0.0397 (11)H14B 0.2697 0.5736 0.6833 0.048*C15B 0.4704 (4) 0.6825 (4) 0.69718 (15) 0.0311 (9)H15B 0.5001 0.6792 0.6655 0.037*C16B 0.7339 (5) 0.9204 (4) 0.93706 (14) 0.0309 (9)C17B 0.8121 (5) 1.0334 (5) 0.96206 (14) 0.0347 (10)H17B 0.8753 1.0995 0.9458 0.042*C18B 0.8010 (5) 1.0529 (5) 1.00985 (15) 0.0383 (11)H18B 0.8573 1.1306 1.0261 0.046*C19B 0.7063 (5) 0.9579 (5) 1.03462 (14) 0.0364 (11)C20B 0.6280 (5) 0.8444 (5) 1.01012 (15) 0.0393 (11)H20B 0.5643 0.7787 1.0264 0.047*C21B 0.6411 (5) 0.8250 (5) 0.96246 (14) 0.0365 (10)H21B 0.5866 0.7459 0.9464 0.044*C1D 0.7073 (7) 0.4443 (7) 0.0391 (3) 0.0815 (19)H1D1 0.6725 0.3416 0.0423 0.122*H1D2 0.7287 0.4623 0.0062 0.122*H1D3 0.6271 0.5112 0.0489 0.122*C2D 0.9311 (9) 0.6126 (8) 0.0632 (3) 0.114 (3)H2D1 0.8642 0.6937 0.0718 0.171*H2D2 0.9550 0.6205 0.0301 0.171*H2D3 1.0265 0.6183 0.0830 0.171*N1D 0.8538 (6) 0.4723 (5) 0.07032 (19) 0.0703 (14)C3D 0.8830 (9) 0.3857 (8) 0.1053 (2) 0.086 (2)H3D 0.9706 0.4112 0.1252 0.103*O1D 0.8107 (6) 0.2744 (4) 0.11575 (15) 0.0827 (14)C2E 0.7544 (8) 0.3853 (9) 0.5056 (2) 0.100 (3)H2E1 0.7862 0.4875 0.4999 0.151*H2E2 0.8435 0.3209 0.5046 0.151*H2E3 0.7113 0.3799 0.5367 0.151*C1E 0.5886 (7) 0.1866 (6) 0.4704 (2) 0.0687 (17)H1E1 0.5338 0.1609 0.4403 0.103*H1E2 0.5197 0.1728 0.4961 0.103*H1E3 0.6776 0.1226 0.4750 0.103*N1E 0.6394 (5) 0.3376 (5) 0.46995 (14) 0.0471 (10)C3E 0.5763 (7) 0.4307 (6) 0.4394 (2) 0.0630 (15)H3E 0.6131 0.5301 0.4413 0.076*O1E 0.4765 (4) 0.4022 (4) 0.40915 (12) 0.0604 (10)P1B 0.18071 (13) 0.17081 (13) 0.58486 (4) 0.0365 (3)F1B 0.3310 (3) 0.1981 (3) 0.55689 (10) 0.0581 (8)F2B 0.1036 (3) 0.3115 (3) 0.56081 (10) 0.0577 (8)F3B 0.0298 (3) 0.1440 (3) 0.61339 (10) 0.0574 (8)

Appendices

F4B 0.2581 (4) 0.0315 (3) 0.60954 (10) 0.0708 (9)F5B 0.1129 (3) 0.0685 (3) 0.54266 (10) 0.0582 (8)F6B 0.2477 (3) 0.2766 (3) 0.62676 (9) 0.0537 (7)

Table A.4. Atomic displacement parameters (Å2)U11 U22 U33 U12 U13 U23

P1A 0.0654 (9) 0.0326 (6) 0.0395 (7) 0.0080 (6) 0.0109 (6) 0.0064 (5)F1A 0.171 (17) 0.091 (17) 0.064 (11) 0.052 (10)F2A 0.23 (3) 0.098 (14) 0.106 (16) 0.131 (18) 0.062 (16) 0.050 (12)F3A 0.098 (17) 0.19 (3) 0.066 (9) (17) 0.037 (12)F4A 0.16 (2) 0.102 (17) 0.078 (13) 0.072 (14) 0.071 (14) 0.030 (11)F5A 0.119 (19) 0.068 (11) 0.033 (7) 0.002 (7) 0.012 (5)F6A 0.078 (18) 0.112 (14) 0.037 (8) 0.014 (9)F1A' 0.40 (2) 0.017 (3) 0.068 (8)F2A' 0.043 (4) 0.144 (9) 0.079 (5) 0.005 (3)F3A' 0.092 (7) 0.039 (4) 0.184 (12) 0.023 (4)F4A' 0.085 (6) 0.177 (12) 0.124 (8) 0.071 (7) 0.032 (5) 0.036 (8)F5A' 0.065 (5) 0.078 (7) 0.051 (4) 0.004 (3) 0.020 (3)F6A' 0.097 (11) 0.120 (9) 0.059 (5) 0.020 (8) 0.024 (4) 0.048 (5)Ru1 0.01729 (17) 0.01850 (17) 0.02258 (18) 0.00113 (12) 0.00061 (11)N1A 0.0179 (16) 0.0248 (17) 0.0301 (19) 0.0004 (13) 0.0019 (14)N2A 0.0212 (17) 0.0288 (17) 0.0260 (18)N3A 0.0177 (16) 0.0220 (17) 0.037 (2)C1A 0.020 (2) 0.0219 (19) 0.039 (2) 0.0018 (16) 0.0025 (17) 0.0018 (17)C2A 0.025 (2) 0.030 (2) 0.053 (3) 0.0033 (18) 0.000 (2) 0.005 (2)C3A 0.028 (2) 0.036 (2) 0.055 (3) 0.017 (2)C4A 0.034 (2) 0.046 (3) 0.034 (3) 0.013 (2)C5A 0.021 (2) 0.033 (2) 0.027 (2) 0.0049 (17)C6A 0.030 (2) 0.037 (2) 0.020 (2) 0.0008 (17)C7A 0.060 (3) 0.050 (3) 0.024 (2) 0.000 (2)C8A 0.063 (3) 0.061 (3) 0.028 (3) 0.006 (2)C9A 0.044 (3) 0.048 (3) 0.043 (3) 0.005 (2)C10A 0.027 (2) 0.029 (2) 0.035 (2) 0.0035 (18)C11A 0.0145 (19) 0.028 (2) 0.045 (3) 0.0035 (17)C12A 0.027 (2) 0.028 (2) 0.066 (3) 0.006 (2)C13A 0.027 (2) 0.023 (2) 0.088 (4) 0.0007 (18) 0.001 (2)C14A 0.030 (2) 0.027 (2) 0.064 (3) 0.000 (2) 0.007 (2)C15A 0.022 (2) 0.024 (2) 0.047 (3) 0.0059 (19)N1B 0.0194 (16) 0.0175 (15) 0.0205 (17) 0.0011 (12) 0.0009 (12)N2B 0.0183 (16) 0.0180 (15) 0.0255 (17) 0.0001 (12) 0.0002 (13) 0.0031 (13)N3B 0.0218 (17) 0.0199 (16) 0.0297 (19) 0.0016 (13) 0.0005 (13)N4B 0.057 (3) 0.058 (3) 0.023 (2) 0.004 (2) 0.0054 (18) 0.0053 (17)C1B 0.025 (2) 0.0214 (18) 0.021 (2) 0.0026 (16) 0.0034 (16) 0.0039 (15)C2B 0.021 (2) 0.032 (2) 0.031 (2) 0.0070 (17) 0.0038 (17)C3B 0.023 (2) 0.036 (2) 0.035 (2) 0.0026 (18)C4B 0.027 (2) 0.034 (2) 0.020 (2) 0.0015 (16)C5B 0.0191 (19) 0.0173 (18) 0.022 (2) 0.0009 (15) 0.0040 (15)

Appendices

C6B 0.024 (2) 0.0195 (18) 0.026 (2) 0.0020 (16) 0.0022 (16) 0.0037 (15)C7B 0.025 (2) 0.024 (2) 0.026 (2) 0.0044 (16) 0.0019 (16)C8B 0.033 (2) 0.0212 (19) 0.028 (2) 0.0038 (17) 0.0061 (18) 0.0035 (16)C9B 0.025 (2) 0.028 (2) 0.033 (2) 0.0080 (18) 0.0074 (17)C10B 0.0177 (19) 0.0162 (17) 0.032 (2) 0.0015 (15) 0.0067 (16) 0.0066 (15)C11B 0.021 (2) 0.0183 (19) 0.037 (2) 0.0038 (15) 0.0030 (17) 0.0066 (16)C12B 0.022 (2) 0.035 (2) 0.044 (3) 0.0001 (18) 0.0064 (19) 0.0106 (19)C13B 0.020 (2) 0.043 (3) 0.060 (3) 0.002 (2) 0.003 (2)C14B 0.024 (2) 0.042 (3) 0.052 (3)C15B 0.025 (2) 0.029 (2) 0.038 (3) 0.0005 (18)C16B 0.029 (2) 0.034 (2) 0.030 (2) 0.0041 (18) 0.0038 (18) 0.0041 (18)C17B 0.039 (3) 0.035 (2) 0.031 (2) 0.0088 (19) 0.0025 (18)C18B 0.046 (3) 0.038 (2) 0.031 (2) 0.004 (2) 0.006 (2)C19B 0.038 (3) 0.046 (3) 0.026 (2) 0.019 (2) 0.0053 (19) 0.011 (2)C20B 0.043 (3) 0.049 (3) 0.027 (2) 0.000 (2) 0.009 (2) 0.012 (2)C21B 0.040 (3) 0.040 (2) 0.030 (2) 0.0021 (19) 0.0070 (19)C1D 0.063 (4) 0.066 (4) 0.114 (6)C2D 0.090 (5) 0.068 (4) 0.180 (9) 0.051 (5)N1D 0.077 (4) 0.047 (3) 0.088 (4) 0.013 (3) 0.007 (3)C3D 0.129 (6) 0.072 (4) 0.056 (4) 0.004 (4) 0.004 (3)O1D 0.131 (4) 0.043 (2) 0.078 (3) 0.051 (3) 0.008 (2)C2E 0.097 (6) 0.134 (7) 0.066 (4) 0.035 (4)C1E 0.093 (5) 0.056 (3) 0.057 (4) 0.022 (3) 0.001 (3)N1E 0.045 (2) 0.054 (3) 0.041 (2) 0.011 (2)C3E 0.080 (4) 0.056 (3) 0.051 (3) 0.010 (3)O1E 0.072 (3) 0.064 (2) 0.044 (2) 0.0093 (18)P1B 0.0381 (7) 0.0418 (7) 0.0300 (6) 0.0029 (5) 0.0057 (5)F1B 0.0526 (18) 0.072 (2) 0.0517 (18) 0.0038 (15) 0.0221 (14)F2B 0.070 (2) 0.0518 (17) 0.0508 (18) 0.0158 (15) 0.0033 (14)F3B 0.0541 (18) 0.0559 (17) 0.0631 (19) 0.0229 (14)F4B 0.103 (3) 0.067 (2) 0.0461 (18) 0.0372 (18) 0.0189 (17) 0.0144 (15)F5B 0.069 (2) 0.0598 (18) 0.0450 (17) 0.0037 (14)F6B 0.0385 (15) 0.079 (2) 0.0426 (16) 0.0006 (14) 0.0002 (12)

Table A.5. Geometric parameters (Å, o)P1A—F1A' 1.525 (6) C2B—H2B 0.9500P1A—F2A' 1.551 (6) C3B—C4B 1.383 (5)P1A—F4A 1.555 (9) C3B—H3B 0.9500P1A—F3A' 1.567 (6) C4B—C5B 1.380 (5)P1A—F3A 1.569 (10) C4B—H4B 0.9500P1A—F6A' 1.575 (7) C5B—C6B 1.483 (5)P1A—F1A 1.576 (9) C6B—C7B 1.380 (5)P1A—F4A' 1.583 (7) C7B—C8B 1.407 (5)P1A—F2A 1.584 (9) C7B—H7B 0.9500P1A—F6A 1.586 (10) C8B—C9B 1.403 (6)P1A—F5A 1.587 (9) C8B—C16B 1.469 (5)P1A—F5A' 1.597 (6) C9B—C10B 1.368 (5)

Appendices

Ru1—N2B 1.971 (3) C9B—H9B 0.9500Ru1—N2A 1.980 (3) C10B—C11B 1.472 (5)Ru1—N3A 2.062 (3) C11B—C12B 1.391 (5)Ru1—N1A 2.062 (3) C12B—C13B 1.372 (6)Ru1—N3B 2.064 (3) C12B—H12B 0.9500Ru1—N1B 2.066 (3) C13B—C14B 1.377 (6)N1A—C1A 1.351 (5) C13B—H13B 0.9500N1A—C5A 1.368 (5) C14B—C15B 1.358 (6)N2A—C6A 1.348 (5) C14B—H14B 0.9500N2A—C10A 1.356 (5) C15B—H15B 0.9500N3A—C15A 1.335 (5) C16B—C17B 1.385 (6)N3A—C11A 1.370 (5) C16B—C21B 1.407 (6)C1A—C2A 1.376 (5) C17B—C18B 1.377 (6)C1A—H1A 0.9500 C17B—H17B 0.9500C2A—C3A 1.373 (6) C18B—C19B 1.406 (6)C2A—H2A 0.9500 C18B—H18B 0.9500C3A—C4A 1.380 (6) C19B—C20B 1.382 (7)C3A—H3A 0.9500 C20B—C21B 1.376 (6)C4A—C5A 1.382 (6) C20B—H20B 0.9500C4A—H4A 0.9500 C21B—H21B 0.9500C5A—C6A 1.467 (6) C1D—N1D 1.533 (8)C6A—C7A 1.386 (6) C1D—H1D1 0.9800C7A—C8A 1.379 (7) C1D—H1D2 0.9800C7A—H7A 0.9500 C1D—H1D3 0.9800C8A—C9A 1.390 (7) C2D—N1D 1.450 (7)C8A—H8A 0.9500 C2D—H2D1 0.9800C9A—C10A 1.383 (6) C2D—H2D2 0.9800C9A—H9A 0.9500 C2D—H2D3 0.9800C10A—C11A 1.462 (6) N1D—C3D 1.295 (8)C11A—C12A 1.389 (6) C3D—O1D 1.229 (8)C12A—C13A 1.376 (7) C3D—H3D 0.9500C12A—H12A 0.9500 C2E—N1E 1.445 (7)C13A—C14A 1.372 (7) C2E—H2E1 0.9800C13A—H13A 0.9500 C2E—H2E2 0.9800C14A—C15A 1.385 (6) C2E—H2E3 0.9800C14A—H14A 0.9500 C1E—N1E 1.429 (7)C15A—H15A 0.9500 C1E—H1E1 0.9800N1B—C1B 1.351 (5) C1E—H1E2 0.9800N1B—C5B 1.359 (5) C1E—H1E3 0.9800N2B—C6B 1.350 (5) N1E—C3E 1.327 (6)N2B—C10B 1.357 (5) C3E—O1E 1.212 (6)N3B—C15B 1.351 (5) C3E—H3E 0.9500N3B—C11B 1.364 (5) P1B—F1B 1.587 (3)N4B—C19B 1.381 (5) P1B—F5B 1.587 (3)N4B—H4B1 0.8800 P1B—F4B 1.595 (3)N4B—H4B2 0.8800 P1B—F6B 1.596 (3)C1B—C2B 1.370 (5) P1B—F2B 1.597 (3)C1B—H1B 0.9500 P1B—F3B 1.600 (3)

Appendices

C2B—C3B 1.383 (6)

F1A'—P1A—F2A' 94.0 (9) C15A—C14A—H14A 120.9F1A'—P1A—F4A 58.7 (10) N3A—C15A—C14A 122.6 (4)F2A'—P1A—F4A 151.6 (15) N3A—C15A—H15A 118.7F1A'—P1A—F3A' 178.5 (7) C14A—C15A—H15A 118.7F2A'—P1A—F3A' 86.7 (5) C1B—N1B—C5B 118.3 (3)F4A—P1A—F3A' 120.8 (13) C1B—N1B—Ru1 127.4 (3)F1A'—P1A—F3A 155.6 (11) C5B—N1B—Ru1 114.2 (2)F2A'—P1A—F3A 109.1 (12) C6B—N2B—C10B 121.1 (3)F4A—P1A—F3A 97.3 (14) C6B—N2B—Ru1 119.5 (2)F3A'—P1A—F3A 24.3 (10) C10B—N2B—Ru1 119.3 (2)F1A'—P1A—F6A' 91.3 (10) C15B—N3B—C11B 118.5 (3)F2A'—P1A—F6A' 91.2 (8) C15B—N3B—Ru1 127.2 (3)F4A—P1A—F6A' 82.5 (11) C11B—N3B—Ru1 114.1 (2)F3A'—P1A—F6A' 90.0 (10) C19B—N4B—H4B1 120.0F3A—P1A—F6A' 80.5 (12) C19B—N4B—H4B2 120.0F1A'—P1A—F1A 27.8 (10) H4B1—N4B—H4B2 120.0F2A'—P1A—F1A 67.9 (12) N1B—C1B—C2B 122.7 (3)F4A—P1A—F1A 86.0 (13) N1B—C1B—H1B 118.6F3A'—P1A—F1A 152.4 (9) C2B—C1B—H1B 118.6F3A—P1A—F1A 176.7 (12) C1B—C2B—C3B 119.2 (4)F6A'—P1A—F1A 100.7 (12) C1B—C2B—H2B 120.4F1A'—P1A—F4A' 92.7 (7) C3B—C2B—H2B 120.4F2A'—P1A—F4A' 173.2 (7) C2B—C3B—C4B 118.5 (4)F4A—P1A—F4A' 35.2 (11) C2B—C3B—H3B 120.7F3A'—P1A—F4A' 86.6 (6) C4B—C3B—H3B 120.7F3A—P1A—F4A' 64.5 (10) C5B—C4B—C3B 120.1 (4)F6A'—P1A—F4A' 89.7 (8) C5B—C4B—H4B 119.9F1A—P1A—F4A' 118.5 (11) C3B—C4B—H4B 119.9F1A'—P1A—F2A 112.4 (13) N1B—C5B—C4B 121.1 (3)F2A'—P1A—F2A 20.3 (15) N1B—C5B—C6B 115.0 (3)F4A—P1A—F2A 171.0 (14) C4B—C5B—C6B 123.9 (3)F3A'—P1A—F2A 68.1 (12) N2B—C6B—C7B 120.5 (3)F3A—P1A—F2A 91.7 (13) N2B—C6B—C5B 112.2 (3)F6A'—P1A—F2A 98.9 (11) C7B—C6B—C5B 127.3 (3)F1A—P1A—F2A 85.0 (14) C6B—C7B—C8B 120.1 (4)F4A'—P1A—F2A 153.1 (13) C6B—C7B—H7B 120.0F1A'—P1A—F6A 86.5 (17) C8B—C7B—H7B 120.0F2A'—P1A—F6A 76.5 (14) C9B—C8B—C7B 117.3 (4)F4A—P1A—F6A 93.0 (17) C9B—C8B—C16B 121.7 (4)F3A'—P1A—F6A 94.9 (17) C7B—C8B—C16B 121.0 (4)F3A—P1A—F6A 91.0 (18) C10B—C9B—C8B 120.9 (4)F6A'—P1A—F6A 15.9 (18) C10B—C9B—H9B 119.6F1A—P1A—F6A 89.5 (18) C8B—C9B—H9B 119.6F4A'—P1A—F6A 105.1 (14) N2B—C10B—C9B 120.2 (4)F2A—P1A—F6A 86.8 (17) N2B—C10B—C11B 112.3 (3)F1A'—P1A—F5A 101.4 (10) C9B—C10B—C11B 127.5 (3)F2A'—P1A—F5A 99.8 (13) N3B—C11B—C12B 120.9 (4)

Appendices

F4A—P1A—F5A 93.7 (16) N3B—C11B—C10B 115.2 (3)F3A'—P1A—F5A 77.2 (10) C12B—C11B—C10B 123.9 (4)F3A—P1A—F5A 83.0 (13) C13B—C12B—C11B 119.0 (4)F6A'—P1A—F5A 162.4 (10) C13B—C12B—H12B 120.5F1A—P1A—F5A 96.1 (13) C11B—C12B—H12B 120.5F4A'—P1A—F5A 77.7 (12) C12B—C13B—C14B 119.9 (4)F2A—P1A—F5A 87.5 (16) C12B—C13B—H13B 120.0F6A—P1A—F5A 172 (2) C14B—C13B—H13B 120.0F1A'—P1A—F5A' 86.2 (6) C15B—C14B—C13B 119.2 (4)F2A'—P1A—F5A' 90.5 (6) C15B—C14B—H14B 120.4F4A—P1A—F5A' 94.8 (9) C13B—C14B—H14B 120.4F3A'—P1A—F5A' 92.5 (6) N3B—C15B—C14B 122.6 (4)F3A—P1A—F5A' 101.2 (10) N3B—C15B—H15B 118.7F6A'—P1A—F5A' 177.0 (10) C14B—C15B—H15B 118.7F1A—P1A—F5A' 77.7 (11) C17B—C16B—C21B 117.3 (4)F4A'—P1A—F5A' 88.9 (6) C17B—C16B—C8B 121.7 (4)F2A—P1A—F5A' 83.4 (10) C21B—C16B—C8B 121.0 (4)F6A—P1A—F5A' 164.5 (13) C18B—C17B—C16B 122.0 (4)F5A—P1A—F5A' 18.6 (9) C18B—C17B—H17B 119.0N2B—Ru1—N2A 177.57 (13) C16B—C17B—H17B 119.0N2B—Ru1—N3A 99.94 (12) C17B—C18B—C19B 120.0 (4)N2A—Ru1—N3A 79.52 (13) C17B—C18B—H18B 120.0N2B—Ru1—N1A 101.75 (12) C19B—C18B—H18B 120.0N2A—Ru1—N1A 78.86 (12) N4B—C19B—C20B 122.0 (4)N3A—Ru1—N1A 158.28 (13) N4B—C19B—C18B 119.4 (4)N2B—Ru1—N3B 79.03 (12) C20B—C19B—C18B 118.6 (4)N2A—Ru1—N3B 103.32 (12) C21B—C20B—C19B 120.9 (4)N3A—Ru1—N3B 89.68 (12) C21B—C20B—H20B 119.5N1A—Ru1—N3B 93.17 (12) C19B—C20B—H20B 119.5N2B—Ru1—N1B 78.97 (12) C20B—C21B—C16B 121.2 (4)N2A—Ru1—N1B 98.68 (12) C20B—C21B—H21B 119.4N3A—Ru1—N1B 93.30 (11) C16B—C21B—H21B 119.4N1A—Ru1—N1B 92.08 (11) N1D—C1D—H1D1 109.5N3B—Ru1—N1B 157.98 (12) N1D—C1D—H1D2 109.5C1A—N1A—C5A 118.1 (3) H1D1—C1D—H1D2 109.5C1A—N1A—Ru1 127.9 (3) N1D—C1D—H1D3 109.5C5A—N1A—Ru1 114.0 (2) H1D1—C1D—H1D3 109.5C6A—N2A—C10A 122.3 (3) H1D2—C1D—H1D3 109.5C6A—N2A—Ru1 119.2 (3) N1D—C2D—H2D1 109.5C10A—N2A—Ru1 118.2 (3) N1D—C2D—H2D2 109.5C15A—N3A—C11A 119.1 (3) H2D1—C2D—H2D2 109.5C15A—N3A—Ru1 127.6 (3) N1D—C2D—H2D3 109.5C11A—N3A—Ru1 113.3 (3) H2D1—C2D—H2D3 109.5N1A—C1A—C2A 122.9 (4) H2D2—C2D—H2D3 109.5N1A—C1A—H1A 118.6 C3D—N1D—C2D 125.0 (6)C2A—C1A—H1A 118.6 C3D—N1D—C1D 118.8 (6)C3A—C2A—C1A 118.9 (4) C2D—N1D—C1D 115.0 (5)C3A—C2A—H2A 120.5 O1D—C3D—N1D 127.5 (7)

Appendices

C1A—C2A—H2A 120.5 O1D—C3D—H3D 116.2C2A—C3A—C4A 119.0 (4) N1D—C3D—H3D 116.2C2A—C3A—H3A 120.5 N1E—C2E—H2E1 109.5C4A—C3A—H3A 120.5 N1E—C2E—H2E2 109.5C3A—C4A—C5A 120.3 (4) H2E1—C2E—H2E2 109.5C3A—C4A—H4A 119.8 N1E—C2E—H2E3 109.5C5A—C4A—H4A 119.8 H2E1—C2E—H2E3 109.5N1A—C5A—C4A 120.6 (4) H2E2—C2E—H2E3 109.5N1A—C5A—C6A 115.4 (3) N1E—C1E—H1E1 109.5C4A—C5A—C6A 124.0 (4) N1E—C1E—H1E2 109.5N2A—C6A—C7A 119.7 (4) H1E1—C1E—H1E2 109.5N2A—C6A—C5A 112.4 (3) N1E—C1E—H1E3 109.5C7A—C6A—C5A 127.8 (4) H1E1—C1E—H1E3 109.5C8A—C7A—C6A 119.0 (4) H1E2—C1E—H1E3 109.5C8A—C7A—H7A 120.5 C3E—N1E—C1E 120.5 (5)C6A—C7A—H7A 120.5 C3E—N1E—C2E 122.0 (5)C7A—C8A—C9A 120.6 (4) C1E—N1E—C2E 117.4 (5)C7A—C8A—H8A 119.7 O1E—C3E—N1E 126.8 (5)C9A—C8A—H8A 119.7 O1E—C3E—H3E 116.6C10A—C9A—C8A 118.9 (4) N1E—C3E—H3E 116.6C10A—C9A—H9A 120.6 F1B—P1B—F5B 89.46 (16)C8A—C9A—H9A 120.6 F1B—P1B—F4B 90.09 (17)N2A—C10A—C9A 119.5 (4) F5B—P1B—F4B 90.71 (17)N2A—C10A—C11A 112.9 (3) F1B—P1B—F6B 90.40 (16)C9A—C10A—C11A 127.6 (4) F5B—P1B—F6B 178.85 (18)N3A—C11A—C12A 120.4 (4) F4B—P1B—F6B 90.43 (17)N3A—C11A—C10A 115.8 (3) F1B—P1B—F2B 90.13 (16)C12A—C11A—C10A 123.8 (4) F5B—P1B—F2B 90.04 (16)C13A—C12A—C11A 119.2 (4) F4B—P1B—F2B 179.22 (18)C13A—C12A—H12A 120.4 F6B—P1B—F2B 88.82 (16)C11A—C12A—H12A 120.4 F1B—P1B—F3B 179.57 (18)C14A—C13A—C12A 120.3 (4) F5B—P1B—F3B 90.97 (16)C14A—C13A—H13A 119.8 F4B—P1B—F3B 89.90 (17)C12A—C13A—H13A 119.8 F6B—P1B—F3B 89.17 (14)C13A—C14A—C15A 118.2 (4) F2B—P1B—F3B 89.88 (17)C13A—C14A—H14A 120.9

A.2 Supplementary data for X-ray diffraction analysis of Ru(II) complex 8

Table A.6. Data collection for Ru(II) complex 8

Tmin = 0.935, Tmax = 0.978 9736 independent reflections65736 measured reflections 8515 reflections with I > 2 (I)Absorption correction: Multi-scan Rint = 0.049

Appendices

Table A.7. Refinement for Ru(II) complex 8

R[F2 > 2 (F2)] = 0.059 180 restraintswR(F2) = 0.144 H atoms treated by a mixture of independent

and constrained refinementS = 1.16 max = 0.97 e Å-3

9736 reflections min = -0.60 e Å-3

814 parameters

Table A.8. Fractional atomic coordinates and isotropic or equivalent isotropic displacement parameters (Å2)

x y z U iso*/Ueq Occ. (<1)Ru1 0.48331 (4) 0.56103 (2) 0.123505 (7) 0.02499 (14)N1A 0.2711 (4) 0.5947 (2) 0.14104 (8) 0.0274 (8)N2A 0.5066 (4) 0.5149 (3) 0.16776 (8) 0.0312 (8)N3A 0.7029 (4) 0.5075 (3) 0.12376 (8) 0.0308 (8)C1A 0.1528 (5) 0.6378 (3) 0.12592 (10) 0.0289 (9)H1A 0.1607 0.6524 0.1044 0.035*C2A 0.0205 (5) 0.6612 (3) 0.14081 (11) 0.0358 (11)H2A 0.6915 0.1296 0.043*C3A 0.0063 (5) 0.6391 (3) 0.17252 (11) 0.0386 (11)H3A 0.6547 0.1830 0.046*C4A 0.1247 (5) 0.5937 (4) 0.18847 (10) 0.0379 (11)H4A 0.1163 0.5776 0.2098 0.045*C5A 0.2565 (5) 0.5718 (3) 0.17255 (10) 0.0319 (10)C6A 0.3894 (5) 0.5267 (3) 0.18784 (10) 0.0348 (10)C7A 0.4081 (6) 0.4999 (4) 0.21942 (11) 0.0478 (13)H7A 0.3274 0.5056 0.2331 0.057*C8A 0.5481 (7) 0.4644 (4) 0.23041 (12) 0.0510 (14)H8A 0.5632 0.4483 0.2519 0.061*C9A 0.6661 (7) 0.4526 (4) 0.20970 (12) 0.0471 (13)H9A 0.7606 0.4291 0.2171 0.057*C10A 0.6410 (5) 0.4764 (3) 0.17783 (11) 0.0360 (11)C11A 0.7503 (5) 0.4682 (3) 0.15227 (11) 0.0342 (10)C12A 0.8889 (5) 0.4234 (3) 0.15600 (13) 0.0405 (12)H12A 0.9175 0.3971 0.1755 0.049*C13A 0.9840 (6) 0.4179 (4) 0.13069 (14) 0.0450 (13)H13A 1.0770 0.3875 0.1328 0.054*C14A 0.9390 (6) 0.4583 (3) 0.10227 (13) 0.0420 (12)H14A 1.0026 0.4560 0.0850 0.050*C15A 0.7999 (5) 0.5024 (3) 0.09930 (11) 0.0334 (10)H15A 0.7717 0.5294 0.0799 0.040*O1B 0.1203 (5) 0.7942 (3) 0.0655 (12)O2B 0.6872 (6) 0.7083 (4) 0.0835 (15)O3B 0.7345 (5) 0.8553 (3) 0.0544 (10)N1B 0.5611 (4) 0.6855 (3) 0.13178 (8) 0.0278 (8)N2B 0.4601 (4) 0.6111 (2) 0.07960 (8) 0.0250 (8)N3B 0.3951 (4) 0.4566 (2) 0.09776 (8) 0.0271 (8)

Appendices

N4B 0.3551 (5) 0.8387 (3) 0.0422 (10)H4B 0.4333 0.8704 0.051*N5B 0.6643 (6) 0.7793 (3) 0.0515 (12)C1B 0.6103 (5) 0.7216 (3) 0.16021 (10) 0.0351 (11)H1B 0.6110 0.6878 0.1787 0.042*C2B 0.6590 (6) 0.8059 (3) 0.16289 (11) 0.0388 (11)H2B 0.6914 0.8284 0.1829 0.047*C3B 0.6594 (6) 0.8570 (4) 0.13562 (11) 0.0426 (12)H3B 0.6933 0.9140 0.1370 0.051*C4B 0.6086 (5) 0.8220 (3) 0.10620 (11) 0.0371 (11)H4B1 0.6079 0.8554 0.0876 0.045*C5B 0.5591 (5) 0.7373 (3) 0.10478 (9) 0.0287 (9)C6B 0.4986 (5) 0.6938 (3) 0.07498 (9) 0.0268 (9)C7B 0.4786 (5) 0.7313 (3) 0.04503 (9) 0.0281 (9)H7B 0.5040 0.7892 0.0422 0.034*C8B 0.4199 (5) 0.6817 (3) 0.01888 (9) 0.0268 (9)C9B 0.3820 (5) 0.5956 (3) 0.02439 (9) 0.0281 (9)H9B 0.3439 0.5613 0.0074 0.034*C10B 0.4006 (5) 0.5605 (3) 0.05516 (9) 0.0264 (9)C11B 0.3622 (5) 0.4730 (3) 0.06542 (9) 0.0267 (9)C12B 0.2963 (5) 0.4108 (3) 0.04535 (10) 0.0333 (10)H12B 0.2729 0.4233 0.0238 0.040*C13B 0.2650 (6) 0.3298 (3) 0.05741 (11) 0.0389 (11)H13B 0.2191 0.2876 0.0441 0.047*C14B 0.3025 (6) 0.3119 (3) 0.08947 (11) 0.0380 (11)H14B 0.2852 0.2573 0.0979 0.046*C15B 0.3662 (5) 0.3772 (3) 0.10872 (10) 0.0316 (10)H15B 0.3901 0.3652 0.1303 0.038*C16B 0.3999 (5) 0.7219 (3) 0.0288 (9)C17B 0.4868 (5) 0.7935 (3) 0.0342 (10)H17B 0.5591 0.8157 0.041*C18B 0.4683 (6) 0.8327 (3) 0.0383 (11)H18B 0.5260 0.8815 0.046*C19B 0.3621 (6) 0.7983 (3) 0.0358 (11)C20B 0.2744 (6) 0.7274 (3) 0.0373 (11)H20B 0.2029 0.7051 0.045*C21B 0.2926 (5) 0.6891 (3) 0.0338 (10)H21B 0.2330 0.6411 0.041*C22B 0.2419 (7) 0.8338 (4) 0.0490 (14)C23B 0.2775 (8) 0.8813 (4) 0.0587 (16)H23A 0.3642 0.9204 0.070*H23B 0.1882 0.9154 0.070*C24B 0.3177 (8) 0.8160 (4) 0.0576 (15)H24A 0.4041 0.7803 0.069*H24B 0.2294 0.7785 0.069*C25B 0.3604 (8) 0.8632 (4) 0.0581 (16)H25A 0.4530 0.8974 0.070*H25B 0.2771 0.9027 0.070*

Appendices

C26B 0.3897 (8) 0.8024 (5) 0.0612 (16)H26A 0.2958 0.7696 0.073*H26B 0.4112 0.8370 0.073*C27B 0.5226 (8) 0.7397 (4) 0.0600 (16)H27A 0.4903 0.6960 0.072*H27B 0.5441 0.7107 0.072*C28B 0.7393 (8) 0.7591 (4) 0.0606 (16)C29B 0.8819 (8) 0.8073 (5) 0.0668 (18)H29B 0.9527 0.8072 0.080*C30B 0.8959 (7) 0.8521 (4) 0.0571 (15)H30B 0.9764 0.8896 0.069*C31B 0.7618 (7) 0.8318 (4) 0.0482 (13)P1A 0.93516 (15) 0.68546 (12) 0.03687 (3) 0.0486 (4)F1A 0.8758 (3) 0.7075 (2) 0.07160 (7) 0.0564 (9)F2A 1.0358 (3) 0.6073 (2) 0.05201 (7) 0.0553 (9)F3A 1.0797 (4) 0.7481 (3) 0.04297 (9) 0.0727 (11)F4A 0.9921 (4) 0.6611 (3) 0.00219 (7) 0.0718 (12)F5A 0.8334 (4) 0.7626 (3) 0.02169 (9) 0.0751 (12)F6A 0.7905 (3) 0.6209 (3) 0.03097 (7) 0.0565 (9)P1B 0.97262 (16) 0.89067 (10) 0.23038 (3) 0.0446 (4)F1B 0.7997 (14) 0.8619 (12) 0.2363 (4) 0.066 (4) 0.56 (3)F2B 0.9861 (16) 0.8163 (10) 0.2048 (4) 0.083 (5) 0.56 (3)F3B 0.908 (3) 0.9526 (9) 0.2022 (4) 0.077 (4) 0.56 (3)F4B 1.143 (2) 0.926 (2) 0.2242 (9) 0.065 (5) 0.44 (3)F5B 0.9643 (16) 0.9714 (10) 0.2542 (4) 0.052 (4) 0.44 (3)F6B 1.041 (2) 0.8326 (8) 0.2589 (4) 0.097 (5) 0.56 (3)F1B' 0.7961 (18) 0.883 (2) 0.2357 (6) 0.094 (7) 0.44 (3)F2B' 0.965 (3) 0.7989 (11) 0.2167 (7) 0.114 (8) 0.44 (3)F3B' 0.925 (3) 0.9322 (18) 0.1977 (3) 0.088 (6) 0.44 (3)F4B' 1.1495 (18) 0.904 (2) 0.2258 (7) 0.082 (7) 0.56 (3)F5B' 0.964 (3) 0.9770 (13) 0.2491 (6) 0.121 (7) 0.56 (3)F6B' 0.999 (3) 0.861 (2) 0.2658 (3) 0.116 (7) 0.44 (3)O1D 0.7385 (5) 0.0191 (3) 0.0741 (13)N1D 0.9267 (7) 0.0640 (4) 0.03604 (15) 0.0696 (15)C1D 0.9186 (11) 0.0556 (2) 0.112 (3)H1D1 0.8688 0.0750 0.168*H1D2 1.0216 0.0607 0.168*H1D3 0.8600 0.0437 0.168*C2D 1.0363 (9) 0.1294 (5) 0.0468 (2) 0.085 (2)H2D1 1.0257 0.1788 0.0329 0.127*H2D2 1.1397 0.1068 0.0463 0.127*H2D3 1.0166 0.1462 0.0684 0.127*C3D 0.8372 (7) 0.0734 (4) 0.01008 (17) 0.0613 (16)H3D 0.8471 0.1240 0.074*O1E 0.3699 (8) 1.0700 (4) 0.11749 (15) 0.104 (2)N1E 0.2293 (9) 0.9481 (5) 0.12085 (19) 0.104 (2)C1E 0.182 (3) 0.9696 (15) 0.1514 (4) 0.313 (11)H1E1 0.2634 1.0010 0.1629 0.470*

Appendices

H1E2 0.1597 0.9177 0.1629 0.470*H1E3 0.0905 1.0049 0.1493 0.470*C2E 0.1935 (13) 0.8633 (6) 0.1096 (4) 0.168 (5)H2E1 0.0865 0.8608 0.1020 0.253*H2E2 0.2116 0.8226 0.1268 0.253*H2E3 0.2581 0.8491 0.0923 0.253*C3E 0.3286 (9) 0.9985 (5) 0.10787 (16) 0.0691 (18)H3E 0.3727 0.9782 0.0894 0.083*O1W 0.5174 (6) 0.9460 (3) 0.04109 (10) 0.0619 (11)H1W 0.600 (3) 0.974 (3) 0.0291 (10) 0.060*H2W 0.415 (2) 0.957 (4) 0.0296 (10) 0.060*

Table A.9. Atomic displacement parameters (Å2)U11 U22 U33 U12 U13 U23

Ru1 0.02127 (19) 0.0393 (2) 0.01389 (18) 0.00076 (15)N1A 0.0242 (18) 0.041 (2) 0.0164 (16)N2A 0.0287 (19) 0.045 (2) 0.0194 (17) 0.0003 (17)N3A 0.0271 (19) 0.036 (2) 0.0284 (19)C1A 0.025 (2) 0.039 (3) 0.021 (2)C2A 0.027 (2) 0.050 (3) 0.030 (2) 0.003 (2)C3A 0.026 (2) 0.056 (3) 0.034 (2) 0.002 (2) 0.0006 (18)C4A 0.035 (3) 0.058 (3) 0.021 (2) 0.0035 (18)C5A 0.030 (2) 0.048 (3) 0.0171 (19) 0.000 (2)C6A 0.031 (2) 0.050 (3) 0.023 (2) (2)C7A 0.052 (3) 0.072 (4) 0.019 (2) 0.004 (3) 0.000 (2) 0.004 (2)C8A 0.059 (3) 0.070 (4) 0.023 (2) 0.005 (3) 0.009 (2)C9A 0.049 (3) 0.057 (4) 0.033 (3) 0.004 (3) 0.006 (2)C10A 0.032 (2) 0.048 (3) 0.027 (2) 0.000 (2) (18) 0.000 (2)C11A 0.027 (2) 0.040 (3) 0.034 (2)C12A 0.030 (2) 0.041 (3) 0.049 (3) 0.004 (2) 0.004 (2)C13A 0.027 (2) 0.046 (3) 0.061 (3) 0.006 (2)C14A 0.028 (2) 0.041 (3) 0.058 (3) 0.013 (2)C15A 0.028 (2) 0.038 (3) 0.035 (2) 0.0036 (18)O1B 0.060 (3) 0.098 (4) 0.037 (2) 0.020 (2)O2B 0.096 (4) 0.101 (4) 0.054 (3) 0.004 (3) 0.011 (2) 0.039 (3)O3B 0.066 (3) 0.069 (3) 0.0280 (18) 0.003 (2) 0.0022 (16) 0.0094 (17)N1B 0.0205 (17) 0.044 (2) 0.0185 (16) 0.0029 (16)N2B 0.0195 (17) 0.037 (2) 0.0178 (16)N3B 0.0265 (18) 0.036 (2) 0.0190 (16) 0.0027 (16) 0.0019 (13)N4B 0.054 (3) 0.049 (3) 0.0234 (19) 0.0069 (17)N5B 0.063 (3) 0.057 (3) 0.034 (2) 0.003 (2) 0.003 (2) 0.009 (2)C1B 0.026 (2) 0.059 (3) 0.019 (2) 0.002 (2)C2B 0.038 (3) 0.050 (3) 0.027 (2)C3B 0.045 (3) 0.047 (3) 0.035 (3)C4B 0.037 (3) 0.048 (3) 0.026 (2)C5B 0.022 (2) 0.043 (3) 0.020 (2)C6B 0.021 (2) 0.038 (3) 0.021 (2) 0.0001 (18)

Appendices

C7B 0.025 (2) 0.040 (3) 0.019 (2) 0.0009 (19)C8B 0.022 (2) 0.040 (3) 0.0187 (19) 0.0012 (19) 0.0019 (15)C9B 0.025 (2) 0.043 (3) 0.0161 (19) 0.0015 (19)C10B 0.023 (2) 0.039 (2) 0.0178 (19) 0.0007 (19)C11B 0.021 (2) 0.040 (2) 0.0196 (19) 0.0021 (18) 0.0004 (15)C12B 0.033 (2) 0.046 (3) 0.021 (2) 0.0005 (17)C13B 0.042 (3) 0.041 (3) 0.033 (2) 0.000 (2)C14B 0.043 (3) 0.036 (3) 0.035 (2) 0.004 (2)C15B 0.032 (2) 0.038 (3) 0.025 (2) 0.001 (2) 0.0023 (17)C16B 0.029 (2) 0.038 (3) 0.0186 (19) 0.0052 (19)C17B 0.034 (2) 0.044 (3) 0.024 (2)C18B 0.044 (3) 0.045 (3) 0.026 (2) 0.0043 (19)C19B 0.043 (3) 0.043 (3) 0.022 (2) 0.005 (2) 0.0002 (18) 0.0006 (19)C20B 0.038 (3) 0.051 (3) 0.023 (2) 0.001 (2)C21B 0.036 (2) 0.042 (3) 0.022 (2) 0.0001 (19)C22B 0.055 (3) 0.063 (4) 0.029 (3) 0.000 (3) 0.001 (2) 0.007 (2)C23B 0.080 (4) 0.066 (4) 0.029 (3) 0.001 (3) 0.013 (3)C24B 0.068 (4) 0.071 (4) 0.034 (3) 0.004 (3) 0.010 (3)C25B 0.065 (4) 0.072 (4) 0.037 (3) 0.010 (3) 0.006 (3) 0.012 (3)C26B 0.065 (4) 0.081 (5) 0.039 (3) 0.005 (3) 0.004 (3)C27B 0.073 (4) 0.062 (4) 0.047 (3) 0.011 (3)C28B 0.071 (4) 0.071 (4) 0.040 (3) 0.007 (3) 0.007 (3) 0.012 (3)C29B 0.069 (4) 0.093 (5) 0.037 (3) 0.000 (4) 0.008 (3)C30B 0.059 (4) 0.073 (4) 0.039 (3) 0.006 (3)C31B 0.057 (3) 0.060 (4) 0.028 (3) 0.003 (3) 0.002 (2) 0.005 (2)P1A 0.0273 (6) 0.0907 (12) 0.0276 (6) 0.0016 (7) 0.0096 (7)F1A 0.0407 (16) 0.095 (3) 0.0334 (15) 0.0078 (17) 0.0015 (12)F2A 0.0363 (16) 0.094 (3) 0.0347 (15) 0.0112 (17) 0.0000 (16)F3A 0.0401 (18) 0.101 (3) 0.077 (2) 0.006 (2)F4A 0.0397 (17) 0.149 (4) 0.0276 (15) 0.0069 (12) 0.0082 (19)F5A 0.0459 (19) 0.109 (3) 0.070 (2) 0.011 (2) 0.038 (2)F6A 0.0336 (15) 0.105 (3) 0.0303 (15)P1B 0.0431 (7) 0.0605 (9) 0.0292 (6)F1B 0.065 (6) 0.095 (8) 0.037 (6) 0.007 (5)F2B 0.053 (5) 0.088 (7) 0.104 (9) 0.019 (5)F3B 0.073 (7) 0.052 (6) 0.100 (10) 0.016 (5)F4B 0.037 (7) 0.097 (11) 0.059 (10) 0.002 (6)F5B 0.036 (6) 0.075 (7) 0.046 (6) 0.007 (4)F6B 0.108 (10) 0.053 (6) 0.120 (10) 0.035 (5)F1B' 0.059 (8) 0.160 (18) 0.065 (10) 0.027 (7)F2B' 0.133 (14) 0.086 (8) 0.117 (15)F3B' 0.073 (9) 0.168 (16) 0.020 (5) 0.023 (7)F4B' 0.041 (6) 0.149 (19) 0.056 (10)F5B' 0.136 (13) 0.120 (11) 0.107 (12) 0.017 (9)F6B' 0.128 (14) 0.176 (17) 0.039 (6) 0.034 (8)O1D 0.056 (3) 0.073 (3) 0.094 (4) (2) 0.003 (2) 0.001 (3)N1D 0.059 (3) 0.074 (4) 0.076 (4) 0.002 (3) 0.010 (3)C1D 0.103 (7) 0.108 (7) 0.122 (8) 0.053 (6)

Appendices

C2D 0.062 (4) 0.086 (6) 0.105 (6) 0.003 (4) 0.001 (5)C3D 0.050 (3) 0.058 (4) 0.078 (4) 0.010 (3)O1E 0.118 (5) 0.099 (4) 0.097 (4) 0.031 (4)N1E 0.116 (6) 0.088 (5) 0.113 (5) 0.052 (4)C1E 0.42 (2) 0.33 (2) 0.218 (16) 0.233 (17)C2E 0.106 (8) 0.065 (6) 0.332 (17) 0.002 (8)C3E 0.075 (4) 0.077 (5) 0.056 (4) 0.005 (3)O1W 0.078 (3) 0.057 (3) 0.051 (2) 0.003 (2) 0.000 (2)

Table A.10. Geometric parameters (Å, o)Ru1—N2A 1.976 (4) C14B—H14B 0.9300Ru1—N2B 1.985 (3) C15B—H15B 0.9300Ru1—N3B 2.060 (4) C16B—C17B 1.386 (7)Ru1—N1B 2.061 (4) C16B—C21B 1.406 (6)Ru1—N3A 2.068 (4) C17B—C18B 1.387 (6)Ru1—N1A 2.076 (4) C17B—H17B 0.9300N1A—C1A 1.346 (5) C18B—C19B 1.399 (7)N1A—C5A 1.371 (5) C18B—H18B 0.9300N2A—C10A 1.350 (6) C19B—C20B 1.374 (7)N2A—C6A 1.361 (6) C20B—C21B 1.389 (6)N3A—C15A 1.356 (6) C20B—H20B 0.9300N3A—C11A 1.376 (6) C21B—H21B 0.9300C1A—C2A 1.378 (6) C22B—C23B 1.524 (7)C1A—H1A 0.9300 C23B—C24B 1.539 (9)C2A—C3A 1.376 (7) C23B—H23A 0.9700C2A—H2A 0.9300 C23B—H23B 0.9700C3A—C4A 1.379 (7) C24B—C25B 1.531 (7)C3A—H3A 0.9300 C24B—H24A 0.9700C4A—C5A 1.391 (6) C24B—H24B 0.9700C4A—H4A 0.9300 C25B—C26B 1.536 (8)C5A—C6A 1.458 (6) C25B—H25A 0.9700C6A—C7A 1.380 (6) C25B—H25B 0.9700C7A—C8A 1.382 (8) C26B—C27B 1.516 (9)C7A—H7A 0.9300 C26B—H26A 0.9700C8A—C9A 1.383 (8) C26B—H26B 0.9700C8A—H8A 0.9300 C27B—H27A 0.9700C9A—C10A 1.382 (7) C27B—H27B 0.9700C9A—H9A 0.9300 C28B—C29B 1.438 (9)C10A—C11A 1.467 (7) C29B—C30B 1.332 (8)C11A—C12A 1.383 (7) C29B—H29B 0.9300C12A—C13A 1.375 (7) C30B—C31B 1.471 (8)C12A—H12A 0.9300 C30B—H30B 0.9300C13A—C14A 1.375 (8) P1A—F3A 1.588 (4)C13A—H13A 0.9300 P1A—F5A 1.591 (4)C14A—C15A 1.380 (7) P1A—F4A 1.597 (3)C14A—H14A 0.9300 P1A—F2A 1.597 (4)C15A—H15A 0.9300 P1A—F1A 1.598 (3)

Appendices

O1B—C22B 1.225 (7) P1A—F6A 1.606 (4)O2B—C28B 1.229 (7) P1B—F2B' 1.530 (15)O3B—C31B 1.213 (6) P1B—F3B' 1.540 (12)N1B—C1B 1.355 (5) P1B—F5B' 1.550 (14)N1B—C5B 1.380 (6) P1B—F6B' 1.552 (12)N2B—C6B 1.339 (6) P1B—F1B' 1.557 (14)N2B—C10B 1.361 (5) P1B—F4B' 1.564 (13)N3B—C15B 1.338 (6) P1B—F2B 1.578 (8)N3B—C11B 1.384 (5) P1B—F6B 1.579 (10)N4B—C22B 1.341 (7) P1B—F3B 1.589 (11)N4B—C19B 1.416 (6) P1B—F1B 1.590 (10)N4B—H4B 0.8600 P1B—F5B 1.598 (11)N5B—C31B 1.387 (7) P1B—F4B 1.601 (14)N5B—C28B 1.394 (7) O1D—C3D 1.254 (8)N5B—C27B 1.447 (8) N1D—C3D 1.301 (8)C1B—C2B 1.372 (7) N1D—C2D 1.438 (9)C1B—H1B 0.9300 N1D—C1D 1.466 (10)C2B—C3B 1.384 (7) C1D—H1D1 0.9600C2B—H2B 0.9300 C1D—H1D2 0.9600C3B—C4B 1.387 (6) C1D—H1D3 0.9600C3B—H3B 0.9300 C2D—H2D1 0.9600C4B—C5B 1.377 (7) C2D—H2D2 0.9600C4B—H4B1 0.9300 C2D—H2D3 0.9600C5B—C6B 1.480 (5) C3D—H3D 0.9300C6B—C7B 1.377 (6) O1E—C3E 1.221 (9)C7B—C8B 1.403 (6) N1E—C3E 1.299 (9)C7B—H7B 0.9300 N1E—C1E 1.398 (13)C8B—C9B 1.393 (7) N1E—C2E 1.420 (10)C8B—C16B 1.485 (6) C1E—H1E1 0.9600C9B—C10B 1.392 (6) C1E—H1E2 0.9600C9B—H9B 0.9300 C1E—H1E3 0.9600C10B—C11B 1.463 (6) C2E—H2E1 0.9600C11B—C12B 1.375 (6) C2E—H2E2 0.9600C12B—C13B 1.382 (7) C2E—H2E3 0.9600C12B—H12B 0.9300 C3E—H3E 0.9300C13B—C14B 1.384 (7) O1W—H1W 0.993 (2)C13B—H13B 0.9300 O1W—H2W 0.993 (2)C14B—C15B 1.383 (6)

N2A—Ru1—N2B 178.23 (15) H23A—C23B—H23B 108.2N2A—Ru1—N3B 102.57 (15) C25B—C24B—C23B 110.5 (5)N2B—Ru1—N3B 79.10 (14) C25B—C24B—H24A 109.5N2A—Ru1—N1B 99.56 (15) C23B—C24B—H24A 109.5N2B—Ru1—N1B 78.77 (14) C25B—C24B—H24B 109.5N3B—Ru1—N1B 157.87 (13) C23B—C24B—H24B 109.5N2A—Ru1—N3A 78.79 (15) H24A—C24B—H24B 108.1N2B—Ru1—N3A 101.87 (14) C24B—C25B—C26B 113.7 (6)N3B—Ru1—N3A 90.05 (14) C24B—C25B—H25A 108.8N1B—Ru1—N3A 94.73 (14) C26B—C25B—H25A 108.8

Appendices

N2A—Ru1—N1A 78.93 (15) C24B—C25B—H25B 108.8N2B—Ru1—N1A 100.42 (14) C26B—C25B—H25B 108.8N3B—Ru1—N1A 94.12 (14) H25A—C25B—H25B 107.7N1B—Ru1—N1A 89.60 (14) C27B—C26B—C25B 114.1 (5)N3A—Ru1—N1A 157.71 (14) C27B—C26B—H26A 108.7C1A—N1A—C5A 117.9 (4) C25B—C26B—H26A 108.7C1A—N1A—Ru1 128.4 (3) C27B—C26B—H26B 108.7C5A—N1A—Ru1 113.7 (3) C25B—C26B—H26B 108.7C10A—N2A—C6A 121.6 (4) H26A—C26B—H26B 107.6C10A—N2A—Ru1 119.3 (3) N5B—C27B—C26B 114.4 (5)C6A—N2A—Ru1 119.0 (3) N5B—C27B—H27A 108.7C15A—N3A—C11A 117.1 (4) C26B—C27B—H27A 108.7C15A—N3A—Ru1 128.6 (3) N5B—C27B—H27B 108.7C11A—N3A—Ru1 114.2 (3) C26B—C27B—H27B 108.7N1A—C1A—C2A 123.1 (4) H27A—C27B—H27B 107.6N1A—C1A—H1A 118.5 O2B—C28B—N5B 123.6 (6)C2A—C1A—H1A 118.5 O2B—C28B—C29B 128.6 (6)C3A—C2A—C1A 119.2 (4) N5B—C28B—C29B 107.8 (5)C3A—C2A—H2A 120.4 C30B—C29B—C28B 108.7 (5)C1A—C2A—H2A 120.4 C30B—C29B—H29B 125.6C2A—C3A—C4A 119.0 (4) C28B—C29B—H29B 125.6C2A—C3A—H3A 120.5 C29B—C30B—C31B 108.1 (6)C4A—C3A—H3A 120.5 C29B—C30B—H30B 125.9C3A—C4A—C5A 119.8 (4) C31B—C30B—H30B 125.9C3A—C4A—H4A 120.1 O3B—C31B—N5B 124.2 (5)C5A—C4A—H4A 120.1 O3B—C31B—C30B 129.1 (5)N1A—C5A—C4A 121.0 (4) N5B—C31B—C30B 106.7 (4)N1A—C5A—C6A 115.4 (4) F3A—P1A—F5A 91.0 (2)C4A—C5A—C6A 123.5 (4) F3A—P1A—F4A 90.5 (2)N2A—C6A—C7A 119.6 (4) F5A—P1A—F4A 90.5 (2)N2A—C6A—C5A 112.9 (4) F3A—P1A—F2A 89.6 (2)C7A—C6A—C5A 127.4 (4) F5A—P1A—F2A 179.3 (2)C6A—C7A—C8A 119.2 (5) F4A—P1A—F2A 89.5 (2)C6A—C7A—H7A 120.4 F3A—P1A—F1A 90.8 (2)C8A—C7A—H7A 120.4 F5A—P1A—F1A 90.1 (2)C7A—C8A—C9A 120.4 (5) F4A—P1A—F1A 178.5 (2)C7A—C8A—H8A 119.8 F2A—P1A—F1A 89.97 (18)C9A—C8A—H8A 119.8 F3A—P1A—F6A 179.1 (2)C10A—C9A—C8A 118.9 (5) F5A—P1A—F6A 89.8 (2)C10A—C9A—H9A 120.6 F4A—P1A—F6A 89.50 (19)C8A—C9A—H9A 120.6 F2A—P1A—F6A 89.5 (2)N2A—C10A—C9A 120.0 (5) F1A—P1A—F6A 89.15 (17)N2A—C10A—C11A 112.9 (4) F2B'—P1B—F3B' 93.0 (12)C9A—C10A—C11A 127.0 (5) F2B'—P1B—F5B' 170.1 (19)N3A—C11A—C12A 122.1 (4) F3B'—P1B—F5B' 93.9 (17)N3A—C11A—C10A 114.6 (4) F2B'—P1B—F6B' 94.5 (11)C12A—C11A—C10A 123.3 (4) F3B'—P1B—F6B' 169.3 (14)C13A—C12A—C11A 119.7 (5) F5B'—P1B—F6B' 77.7 (19)

Appendices

C13A—C12A—H12A 120.2 F2B'—P1B—F1B' 87.7 (14)C11A—C12A—H12A 120.2 F3B'—P1B—F1B' 86.7 (14)C12A—C13A—C14A 118.7 (5) F5B'—P1B—F1B' 85.7 (14)C12A—C13A—H13A 120.7 F6B'—P1B—F1B' 86.1 (13)C14A—C13A—H13A 120.7 F2B'—P1B—F4B' 95.7 (18)C13A—C14A—C15A 120.2 (5) F3B'—P1B—F4B' 92.9 (15)C13A—C14A—H14A 119.9 F5B'—P1B—F4B' 91.0 (14)C15A—C14A—H14A 119.9 F6B'—P1B—F4B' 93.8 (16)N3A—C15A—C14A 122.2 (5) F1B'—P1B—F4B' 176.6 (18)N3A—C15A—H15A 118.9 F2B'—P1B—F2B 22.2 (13)C14A—C15A—H15A 118.9 F3B'—P1B—F2B 74.5 (11)C1B—N1B—C5B 117.5 (4) F5B'—P1B—F2B 167.3 (15)C1B—N1B—Ru1 127.9 (3) F6B'—P1B—F2B 114.4 (12)C5B—N1B—Ru1 114.6 (3) F1B'—P1B—F2B 98.5 (11)C6B—N2B—C10B 121.9 (4) F4B'—P1B—F2B 84.6 (13)C6B—N2B—Ru1 119.5 (3) F2B'—P1B—F6B 76.1 (11)C10B—N2B—Ru1 118.6 (3) F3B'—P1B—F6B 166.8 (11)C15B—N3B—C11B 117.9 (4) F5B'—P1B—F6B 97.8 (14)C15B—N3B—Ru1 127.7 (3) F6B'—P1B—F6B 23.8 (13)C11B—N3B—Ru1 114.4 (3) F1B'—P1B—F6B 100.2 (12)C22B—N4B—C19B 128.3 (5) F4B'—P1B—F6B 81.0 (16)C22B—N4B—H4B 115.9 F2B—P1B—F6B 93.2 (7)C19B—N4B—H4B 115.9 F2B'—P1B—F3B 106.2 (11)C31B—N5B—C28B 108.2 (5) F3B'—P1B—F3B 14.5 (14)C31B—N5B—C27B 125.5 (5) F5B'—P1B—F3B 80.1 (13)C28B—N5B—C27B 125.4 (5) F6B'—P1B—F3B 155.2 (14)N1B—C1B—C2B 122.8 (4) F1B'—P1B—F3B 81.2 (13)N1B—C1B—H1B 118.6 F4B'—P1B—F3B 97.5 (16)C2B—C1B—H1B 118.6 F2B—P1B—F3B 88.7 (8)C1B—C2B—C3B 119.4 (4) F6B—P1B—F3B 177.4 (9)C1B—C2B—H2B 120.3 F2B'—P1B—F1B 77.3 (14)C3B—C2B—H2B 120.3 F3B'—P1B—F1B 92.6 (11)C2B—C3B—C4B 119.1 (5) F5B'—P1B—F1B 95.4 (11)C2B—C3B—H3B 120.4 F6B'—P1B—F1B 81.7 (11)C4B—C3B—H3B 120.4 F1B'—P1B—F1B 11.8 (17)C5B—C4B—C3B 119.4 (5) F4B'—P1B—F1B 171.3 (16)C5B—C4B—H4B1 120.3 F2B—P1B—F1B 90.4 (8)C3B—C4B—H4B1 120.3 F6B—P1B—F1B 92.2 (9)C4B—C5B—N1B 121.8 (4) F3B—P1B—F1B 89.5 (10)C4B—C5B—C6B 124.2 (4) F2B'—P1B—F5B 162.5 (16)N1B—C5B—C6B 114.0 (4) F3B'—P1B—F5B 101.8 (14)N2B—C6B—C7B 120.8 (4) F5B'—P1B—F5B 8.2 (17)N2B—C6B—C5B 113.1 (4) F6B'—P1B—F5B 69.6 (16)C7B—C6B—C5B 126.1 (4) F1B'—P1B—F5B 84.0 (11)C6B—C7B—C8B 119.7 (4) F4B'—P1B—F5B 92.8 (12)C6B—C7B—H7B 120.2 F2B—P1B—F5B 175.3 (11)C8B—C7B—H7B 120.2 F6B—P1B—F5B 90.3 (10)C9B—C8B—C7B 118.2 (4) F3B—P1B—F5B 87.8 (10)

Appendices

C9B—C8B—C16B 122.2 (4) F1B—P1B—F5B 92.7 (9)C7B—C8B—C16B 119.7 (4) F2B'—P1B—F4B 106 (2)C10B—C9B—C8B 120.5 (4) F3B'—P1B—F4B 85.5 (17)C10B—C9B—H9B 119.7 F5B'—P1B—F4B 81.9 (17)C8B—C9B—H9B 119.7 F6B'—P1B—F4B 99.7 (17)N2B—C10B—C9B 118.9 (4) F1B'—P1B—F4B 165 (2)N2B—C10B—C11B 113.3 (3) F4B'—P1B—F4B 12 (3)C9B—C10B—C11B 127.8 (4) F2B—P1B—F4B 91.9 (15)C12B—C11B—N3B 121.2 (4) F6B—P1B—F4B 90.3 (16)C12B—C11B—C10B 124.2 (4) F3B—P1B—F4B 87.9 (16)N3B—C11B—C10B 114.6 (4) F1B—P1B—F4B 176.5 (16)C11B—C12B—C13B 119.7 (4) F5B—P1B—F4B 84.9 (15)C11B—C12B—H12B 120.2 C3D—N1D—C2D 122.0 (6)C13B—C12B—H12B 120.2 C3D—N1D—C1D 120.6 (7)C12B—C13B—C14B 119.4 (4) C2D—N1D—C1D 117.5 (7)C12B—C13B—H13B 120.3 N1D—C1D—H1D1 109.5C14B—C13B—H13B 120.3 N1D—C1D—H1D2 109.5C15B—C14B—C13B 118.5 (5) H1D1—C1D—H1D2 109.5C15B—C14B—H14B 120.7 N1D—C1D—H1D3 109.5C13B—C14B—H14B 120.7 H1D1—C1D—H1D3 109.5N3B—C15B—C14B 123.2 (4) H1D2—C1D—H1D3 109.5N3B—C15B—H15B 118.4 N1D—C2D—H2D1 109.5C14B—C15B—H15B 118.4 N1D—C2D—H2D2 109.5C17B—C16B—C21B 118.0 (4) H2D1—C2D—H2D2 109.5C17B—C16B—C8B 120.6 (4) N1D—C2D—H2D3 109.5C21B—C16B—C8B 121.5 (4) H2D1—C2D—H2D3 109.5C16B—C17B—C18B 121.5 (4) H2D2—C2D—H2D3 109.5C16B—C17B—H17B 119.3 O1D—C3D—N1D 124.8 (7)C18B—C17B—H17B 119.3 O1D—C3D—H3D 117.6C17B—C18B—C19B 119.5 (5) N1D—C3D—H3D 117.6C17B—C18B—H18B 120.3 C3E—N1E—C1E 118.0 (10)C19B—C18B—H18B 120.3 C3E—N1E—C2E 123.6 (9)C20B—C19B—C18B 120.1 (4) C1E—N1E—C2E 116.8 (12)C20B—C19B—N4B 123.3 (4) N1E—C1E—H1E1 109.5C18B—C19B—N4B 116.5 (4) N1E—C1E—H1E2 109.5C19B—C20B—C21B 120.0 (4) H1E1—C1E—H1E2 109.5C19B—C20B—H20B 120.0 N1E—C1E—H1E3 109.5C21B—C20B—H20B 120.0 H1E1—C1E—H1E3 109.5C20B—C21B—C16B 121.0 (4) H1E2—C1E—H1E3 109.5C20B—C21B—H21B 119.5 N1E—C2E—H2E1 109.5C16B—C21B—H21B 119.5 N1E—C2E—H2E2 109.5O1B—C22B—N4B 124.0 (5) H2E1—C2E—H2E2 109.5O1B—C22B—C23B 121.8 (5) N1E—C2E—H2E3 109.5N4B—C22B—C23B 114.1 (5) H2E1—C2E—H2E3 109.5C22B—C23B—C24B 110.0 (5) H2E2—C2E—H2E3 109.5C22B—C23B—H23A 109.7 O1E—C3E—N1E 126.4 (7)C24B—C23B—H23A 109.7 O1E—C3E—H3E 116.8C22B—C23B—H23B 109.7 N1E—C3E—H3E 116.8

Appendices

C24B—C23B—H23B 109.7 H1W—O1W—H2W 109.4 (3)

N2A—Ru1—N1A—C1A 178.9 (4) N3A—Ru1—N3B—C15B 76.8 (4)N2B—Ru1—N1A—C1A 0.6 (4) N1A—Ru1—N3B—C15BN3B—Ru1—N1A—C1A N2A—Ru1—N3B—C11B 177.8 (3)N1B—Ru1—N1A—C1A 79.1 (4) N2B—Ru1—N3B—C11BN3A—Ru1—N1A—C1A N1B—Ru1—N3B—C11BN2A—Ru1—N1A—C5A 1.7 (3) N3A—Ru1—N3B—C11BN2B—Ru1—N1A—C5A N1A—Ru1—N3B—C11B 98.2 (3)N3B—Ru1—N1A—C5A 103.7 (3) C5B—N1B—C1B—C2BN1B—Ru1—N1A—C5A Ru1—N1B—C1B—C2BN3A—Ru1—N1A—C5A 3.4 (6) N1B—C1B—C2B—C3BN2B—Ru1—N2A—C10A C1B—C2B—C3B—C4B 0.8 (8)N3B—Ru1—N2A—C10A 89.7 (4) C2B—C3B—C4B—C5BN1B—Ru1—N2A—C10A C3B—C4B—C5B—N1BN3A—Ru1—N2A—C10A 2.2 (4) C3B—C4B—C5B—C6B 178.3 (4)N1A—Ru1—N2A—C10A C1B—N1B—C5B—C4B 1.6 (6)N2B—Ru1—N2A—C6A 67 (5) Ru1—N1B—C5B—C4BN3B—Ru1—N2A—C6A C1B—N1B—C5B—C6BN1B—Ru1—N2A—C6A 85.9 (4) Ru1—N1B—C5B—C6B 1.0 (4)N3A—Ru1—N2A—C6A 178.8 (4) C10B—N2B—C6B—C7B 0.3 (6)N1A—Ru1—N2A—C6A Ru1—N2B—C6B—C7BN2A—Ru1—N3A—C15A 177.1 (4) C10B—N2B—C6B—C5B 179.2 (4)N2B—Ru1—N3A—C15A Ru1—N2B—C6B—C5B 1.3 (5)N3B—Ru1—N3A—C15A 74.3 (4) C4B—C5B—C6B—N2B 179.0 (4)N1B—Ru1—N3A—C15A N1B—C5B—C6B—N2BN1A—Ru1—N3A—C15A 175.3 (4) C4B—C5B—C6B—C7BN2A—Ru1—N3A—C11A 1.1 (3) N1B—C5B—C6B—C7B 177.4 (4)N2B—Ru1—N3A—C11A 179.4 (3) N2B—C6B—C7B—C8BN3B—Ru1—N3A—C11A C5B—C6B—C7B—C8B 180.0 (4)N1B—Ru1—N3A—C11A 99.9 (3) C6B—C7B—C8B—C9B 0.7 (6)N1A—Ru1—N3A—C11A C6B—C7B—C8B—C16BC5A—N1A—C1A—C2A 1.4 (7) C7B—C8B—C9B—C10B 0.7 (6)Ru1—N1A—C1A—C2A C16B—C8B—C9B—C10BN1A—C1A—C2A—C3A C6B—N2B—C10B—C9B 1.1 (6)C1A—C2A—C3A—C4A Ru1—N2B—C10B—C9B 179.1 (3)C2A—C3A—C4A—C5A 0.8 (8) C6B—N2B—C10B—C11BC1A—N1A—C5A—C4A Ru1—N2B—C10B—C11BRu1—N1A—C5A—C4A 176.6 (4) C8B—C9B—C10B—N2BC1A—N1A—C5A—C6A C8B—C9B—C10B—C11B 177.8 (4)Ru1—N1A—C5A—C6A C15B—N3B—C11B—C12B 2.5 (6)C3A—C4A—C5A—N1A Ru1—N3B—C11B—C12BC3A—C4A—C5A—C6A 177.7 (5) C15B—N3B—C11B—C10BC10A—N2A—C6A—C7A 0.8 (7) Ru1—N3B—C11B—C10B 1.9 (4)Ru1—N2A—C6A—C7A N2B—C10B—C11B—C12B 177.9 (4)C10A—N2A—C6A—C5A 178.2 (4) C9B—C10B—C11B—C12BRu1—N2A—C6A—C5A 1.6 (6) N2B—C10B—C11B—N3BN1A—C5A—C6A—N2A C9B—C10B—C11B—N3B 179.5 (4)C4A—C5A—C6A—N2A N3B—C11B—C12B—C13B

Appendices

N1A—C5A—C6A—C7A 177.0 (5) C10B—C11B—C12B—C13B 179.8 (4)C4A—C5A—C6A—C7A C11B—C12B—C13B—C14BN2A—C6A—C7A—C8A 2.5 (8) C12B—C13B—C14B—C15B 1.9 (7)C5A—C6A—C7A—C8A C11B—N3B—C15B—C14BC6A—C7A—C8A—C9A Ru1—N3B—C15B—C14B 178.1 (3)C7A—C8A—C9A—C10A C13B—C14B—C15B—N3BC6A—N2A—C10A—C9A C9B—C8B—C16B—C17BRu1—N2A—C10A—C9A 172.6 (4) C7B—C8B—C16B—C17B 23.9 (6)C6A—N2A—C10A—C11A 178.7 (4) C9B—C8B—C16B—C21B 24.2 (6)Ru1—N2A—C10A—C11A C7B—C8B—C16B—C21BC8A—C9A—C10A—N2A 3.7 (8) C21B—C16B—C17B—C18B 0.7 (7)C8A—C9A—C10A—C11A C8B—C16B—C17B—C18BC15A—N3A—C11A—C12A C16B—C17B—C18B—C19BRu1—N3A—C11A—C12A 175.1 (4) C17B—C18B—C19B—C20B 1.5 (7)C15A—N3A—C11A—C10A 179.7 (4) C17B—C18B—C19B—N4BRu1—N3A—C11A—C10A C22B—N4B—C19B—C20B 19.8 (8)N2A—C10A—C11A—N3A 5.5 (6) C22B—N4B—C19B—C18BC9A—C10A—C11A—N3A (5) C18B—C19B—C20B—C21BN2A—C10A—C11A—C12A N4B—C19B—C20B—C21B 177.3 (5)C9A—C10A—C11A—C12A 9.4 (8) C19B—C20B—C21B—C16B 0.1 (7)N3A—C11A—C12A—C13A 0.5 (8) C17B—C16B—C21B—C20B 0.0 (7)C10A—C11A—C12A—C13A 179.2 (5) C8B—C16B—C21B—C20B 179.7 (4)

C11A—C12A—C13A—C14A 0.7 (8) C19B—N4B—C22B—O1B 3.0 (10)

C12A—C13A—C14A—C15A C19B—N4B—C22B—C23B

C11A—N3A—C15A—C14A 1.3 (7) O1B—C22B—C23B—C24BRu1—N3A—C15A—C14A N4B—C22B—C23B—C24B 105.3 (6)C13A—C14A—C15A—N3A C22B—C23B—C24B—C25BN2A—Ru1—N1B—C1B C23B—C24B—C25B—C26BN2B—Ru1—N1B—C1B 178.5 (4) C24B—C25B—C26B—C27BN3B—Ru1—N1B—C1B 177.8 (3) C31B—N5B—C27B—C26BN3A—Ru1—N1B—C1B C28B—N5B—C27B—C26B 120.3 (6)N1A—Ru1—N1B—C1B 77.8 (4) C25B—C26B—C27B—N5BN2A—Ru1—N1B—C5B C31B—N5B—C28B—O2BN2B—Ru1—N1B—C5B C27B—N5B—C28B—O2BN3B—Ru1—N1B—C5B C31B—N5B—C28B—C29B 5.4 (7)N3A—Ru1—N1B—C5B 100.9 (3) C27B—N5B—C28B—C29B 175.1 (6)N1A—Ru1—N1B—C5B O2B—C28B—C29B—C30B 177.6 (7)N2A—Ru1—N2B—C6B 19 (5) N5B—C28B—C29B—C30BN3B—Ru1—N2B—C6B 179.1 (3) C28B—C29B—C30B—C31BN1B—Ru1—N2B—C6B C28B—N5B—C31B—O3B 174.5 (6)N3A—Ru1—N2B—C6B C27B—N5B—C31B—O3B 4.8 (10)N1A—Ru1—N2B—C6B 86.9 (3) C28B—N5B—C31B—C30BN2A—Ru1—N2B—C10B C27B—N5B—C31B—C30BN3B—Ru1—N2B—C10B 1.1 (3) C29B—C30B—C31B—O3BN1B—Ru1—N2B—C10B C29B—C30B—C31B—N5B 5.1 (7)

Appendices

N3A—Ru1—N2B—C10B 88.9 (3) C2D—N1D—C3D—O1DN1A—Ru1—N2B—C10B C1D—N1D—C3D—O1D 0.7 (11)N2A—Ru1—N3B—C15B C1E—N1E—C3E—O1EN2B—Ru1—N3B—C15B 178.9 (4) C2E—N1E—C3E—O1EN1B—Ru1—N3B—C15B 179.6 (3)

Appendices

Appendix B

Estimation of dimer bioconjugate yield by gel electrophoresis

A typical calculation for the dimer bioconjugate example cyt c-10-cyt c. After collection

of product band containing a mixture of unreacted protein, dimer and monomer

bioconjugate (determined by MALDI-TOF mass spectrometry), the concentration of

mixture was estimated by Beer-Lambert law,

[Proteincyt c] = 0.841 a.u./(97.6 mM-1cm-1 1 cm) (cyt c: 410 = 97.6 mM-1cm-1)

= 8.62 10-3 mM

Amount of mixture = 8.62 10-3 mM 220 L

= 1.89 10-9 mol

To estimate the fraction of dimer in the mixture, gel electrophoresis was performed to

separate the components. The estimation of relative concentration of bands within the

lane was determined using the ‘gel analysis’ function in ImageJ v1.42q as shown in

Figure B.1.

Figure B.1. Estimated relative composition of bands in purified bioconjugate mixture cyt c-10-cyt c.

Therefore, amount of dimer = composition amount of mixture

= 0.119 1.89 10-9 mol

= 2.25 10-10 mol

Yield = (amount of dimer/starting ligand concentration) 100%

Appendices

= (2.25 10-10 mol/3.1325 10-10 mol) 100%

= 0.7%

1%

Appendices

Appendix C

Standard nitrite curve using the Griess assay

Table C.1. Nitrite anion standard curved based on absorbance using Griess assay.Nitrite concentration / M Average absorbance /a.u.a

0 0.0032±0.00261 0.0386±0.00182 0.0765±0.00023 0.1118±0.00074 0.1488±0.00155 0.1950±0.00566 0.2111±0.0020

a Monitored absorbance at 540 nm. Error bars are standard deviation.

Figure C.1. Standard nitrite anion concentration standard curve.

Appendices

Appendix D

Theoretical enzyme encapsulation efficiency of egg PC liposomes

In a representative calculation for egg L- -phosphatidylcholine liposomes,

Areahead = 69 Å2 (=0.69 nm2)1 and vesicle d = 105 nm

Surface areavesicle = 4 r2 = 4 (105/2)2 = 34636.0 nm2

Therefore, number of phospholipid molecules = surface areavesicle/Areahead 2 because

of a bilayer.

= (34636.0/0.69) 2 molecules/vesicle

= 100394.2 molecules/vesicle (MW of egg PC = 770.12 g/mol)

Hence, one vesicle = number of phospholipid molecules/NA

= 100394.2/6.023 1023 mol

= 1.67 10-19 mol

Stock (100 L): 100 M iso-1 cyt c = 1 10-8 mol; 15 mg/mL egg PC = 1.5 mg

Moles of phospholipid = mass of lipid/MWegg PC

= 1.5 10-3/770.12 mol

= 1.95 10-6 mol

Number of vesicles = moles of total phospholipid/moles of phospholipid for one vesicle

= 1.95 10-6 mol/1.67 10-19 mol

= 1.16 1013 vesicles

Volumevesicle = (4/3) r3 = 606131 nm3

= 6.06 10-16 cm3

= 6.06 10-16 mL

Therefore, total lumen volume = volumevesicle number of vesicles

= 6.06 10-16 mL 1.16 1013 vesicles

= 7.03 10-6 L

Appendices

[iso-1 cyt c]lumen = 100 M

Therefore, total proteins in lumen = [iso-1 cyt c]lumen total lumen volume

= 100 10-6 M 7.03 10-6 L

= 7.03 10-10 mol

Hence, encapsulation efficiency (%) = proteins in lumen/proteins total

= 7.03 10-10 mol/1 10-8 mol 100%

= 7.0%

Determination of encapsulation efficiency via fluorescence

With the following representative calculation, the encapsulation efficiency (EE%) was

determined for dye labelled cytochrome c and amGFP.

EE% = total number of enzymes encapsulated/number of enzymes added

For 5 M of cyt c added for a 200 L buffer solution:

ncyt c added = (200 × 10-6 L) × (5 × 10-6 M) = 1 × 10-9 mol

Now, enzymes encapsulated determined by estimating total fluorescence of sample after

dialysis; for 1 a.u. fluorescence emission intensity ( ex = 488 nm, em= 513 nm),

equivalent to 4.2 × 10-3 M cyt c-dye (determined experimentally). Therefore, since

sample has fluorescence intensity of 849 a.u. in PBS (166 L) after dialysis,

nencapsulated = 849 × (4.2 × 10-9 M) × (166 × 10-6 L) = 5.9 × 10-10 mol

EE% = (nencapsulated/ncyt c added) × 100 = (5.9 × 10-10/1 × 10-9) × 100 = 59%

Appendices

Appendix E

pH titration of 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS)

Figure E.1. pH titration showing the changes in the excitation spectra ( em = 510 nm) of pyranine encapsulated in a diluted (0.003 mg/mL) PS140-b-PAA48 polymer micelles in 20 mM sodium dihydrogen phosphate with 20 mM ethylenediaminetetraacetic acid. Legend corresponds to pH values.

Appendices

Photoreduction scattering by EDTA by-products of HPTS

Figure E.2. Photoreduction scattering from EDTA by-products under various conditions. (a) Excitation spectra of HPTS encapsulated in a poly-L-lysine: PS140-b-PAA48 polymersome in 5 mM sodium dihydrogen phosphate, 5 mM EDTA at pH 7.2.This experiment shows the characteristic (ratiometric) changes in the HPTS excitation spectra with the 405 nm maxima moving up and the 460 nm maxima moving down following slight acidification after irradiation of this system in the absence of Ru(II)-bisterpyridine chromophore and hence no EDTA by-product formation after irradiation. (b) Excitation spectra of 8-cyt c:CcOx:polymersome in 50 mM unbuffered potassium chloride, 5 mM EDTA, pH 7.2. In this experiment, there is an overall increase in excitation spectra baseline due to scattering caused by EDTA by-products from photoreduction resulting in increase in both the 405 nm and 460 nm excitation maxima for HPTS. (c) UV-Vis spectra of bulk 8-cyt c after photoreduction in 5 mM sodium dihydrogen phosphate, 5 mM EDTA, pH 7.2 showing increased scattering due to EDTA by-products. (d) UV-Vis spectra of 8-cyt c:polymersome after photoreduction in 5 mMsodium dihydrogen phosphate, 5 mM EDTA, pH 7.2 showing increased scattering due to EDTA by-products.

Appendices

Appendix F

F.1 Optical density correction

The number of photons absorbed by a chromophore can be estimated based on

transmittance using the formula TA log where A is absorbance and T is

transmittance.2

Figure F.1. Transmittance of light based on absorbance of chromophore absorption maxima.

In a typical calculation, for a Ru(II)-bisterpyridine chromophore with an absorbance of

0.0115 a.u, an equivalent 97.9% transmittance of photons is observed. Therefore, 2.1%

of incident photons are absorbed by the complex participating in photoexcitation, which

is the quantity used for subsequent quantum efficiency calculations.

F.2 Quantum efficiency of photoreduced 8-cyt c bioconjugates

In a typical calculation, assume that the initial rate of heme reduction is within the linear

region.

Absorbancephotoreduction = 0.007373 a.u. ( 550 = 24.3 mM-1cm-1)

Appendices

Abs = b[8-cyt c]

[8-cyt c] = 3.03 10-7 M (in an 80 L solution)

moles of reduced 8-cyt c = 2.427 10-11 mol

= 2.427 10-11 NA electrons in 1860 s

= 1.46 1013 electrons in 1860 s

Therefore, rate of heme reduction is 7.85 109 electrons/s

%efficiency = (rate of heme reduction/incident photons) 100% ( incident =

1.4 1016 photons/s)

= (7.85 109/1.4 1016) 100%

= 5.59 10-5 %

= %100)correctiondensity opticalphotons(incident

reductionhemeofrate

= (7.85 109/[1.4 1016 0.095]) 100% (For 0.046 a.u., 9.5% photons

contributing to excitation)

= 5.9 10-4%

F.3 Determinatin of proton pumping rate (H+/s)

With the following representative calculation, proton pumping rate was determined.

Vout

Vin

Appendices

Estimation of number of polymer molecules within membrane of one polymersome:

Vmembrane = Vout – Vin

Vout = 3

34

outd where dout = diameter of vesicles (dout = 367±185 nm) and

membrane thickness = 98±35 nm.

Vout = 2.06 × 10-19 m3

Vin = 3

34

ind

Vout = 8.09 × 10-20 m3

Hence, Vmembrane = 1.25 × 10-19 m3

Therefore, one polymersome occupies total volume of

Vmembrane = 1.25 × 10-19 m3.

As a result, number of polymer molecules in one polymersome nPS-b-PAA;

nPS-b-PAA =PAAbPS

membrane

VV

= 4.76 × 107 molecules

= 7.90 × 10-17 mol

mpolymersome = 1.42 × 10-12 g (for one polymersome).

Estimation of number of polymersomes in reaction sample:

Now, the mass of PS-b-PAA in 120 mL for a 1:6 THF:Buffer solution with

1 mg/mL polymer dissolved in THF and MWPS-b-PAA = 18054 g/mol is 0.02 mg

(6.67 × 1014 molecules).

Total number of polymersomes (Npolymersomes) in reaction sample is then;

Npolymersomes = 6.67 × 1014 molecules/4.76 × 107 molecules

= 1.40 × 107 polymersomes

Estimation of total encapsulated volume in reaction sample:

As a result, total encapsulated volume in a reaction solution (Vtotal inside);

Vtotal inside = Npolymersomes × Vin

= 1.13 × 10-12 m3

=1.13 × 10-9 L

dPS-b-PAA 0.27 nm

lPS-b-PAA 46 nm

VPS-b-PAA = ld 2)2

(

VPS-b-PAA = 2.63 × 10-27 m3

Appendices

Estimation of proton pumping rate:

Initial average internal pH for pH 7.2 buffered experiment was pH 7.28 and

increased to pH 7.44 over 3360 s of irradiation.

Hence, [H+] = 1.63 × 10-8 M protons pumped.

Therefore, nH+ = [H+] × Vtotal inside

= 1.85 × 10-17 mol

= 1.12 × 107 H+ pumped across membrane over 3360 s.

Rate of proton pumping = 3.33 × 103 H+/s.

F.4 Quantum efficiency of proton pumping

= %100)correctiondensity opticalphotons(incident

ratepumpingproton ( incident =

1.4 1016 photons/s)

= (3333/[1.4 1016 0.021]) 100% (For 0.0115 a.u., 2.1% photons contributing to

excitation)

= 1.1 10-9%

Appendices

Appendix G

Abbreviations

ABTS 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid)

alpha

beta

br broad

ca. circa

NA Avogadro constant

J coupling constantoC degrees Celsius

d deuterium

s singlet (NMR)

s strong (IR)

d doublet

t triplet

q quartet

p pentet

m medium (IR)

m multiplet (NMR)

w weak (IR)

g grams

mg milligrams

h hours

Hz hertz

MHz megahertz

nuclear magnetic resonance chemical shift (ppm)

ppm parts per million

mp melting point

m/z mass to charge ratio

MS mass spectrometry

IR infrared

UV-Vis ultraviolet visible spectroscopy

L litre

Appendices

mL millilitre

L microlitre

M molar

mM millimolar

M micromolar

nM nanomolar

mol mole

mmol millimole

mol micromole

nmol nanomole

min minutes

molar absorptivity

r.t. room temperature

cmd centimeters diameter (chromatography column diameter)

cmh centimeters height (chromatography column height)

CLSM confocal laser-scanning microscopy

TEM transmission electron microscopy

FPLC fast protein liquid chromatography

HPLC high-performance liquid chromatography

ESI electrospray ionisation

PBS phosphate buffered saline

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

DIPEA N,N-diisopropylethylamine

DMF N,N-dimethylformamide

CH2Cl2 dichloromethane

CDCl3 chloroform

CH3CN acetonitrile

DMSO dimethyl sulfoxide

MeOH methanol

EtOH ethanol

THF tetrahydrofuran

TLC thin layer chromatography

Appendices

MALDI-TOF matrix assisted laser desorption ionisation time of flight mass

spectrometry

UFTRPL ultrafast time-resolved photoluminescence

TCSPC time-correlated single photon counting

ESI electrospray ionisation

SEC size exclusion chromatography

CEX strong cation exchange chromatography

IMAC immobilised metal affinity chromatography

PC L- -phosphatidylcholine

POPC L-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine

PS-b-PAA polystyrene-b-poly(acrylic acid)

EDTA ethylenediaminetetraacetic acid

BSA bovine serum albumin

cyt c cytochrome c

CcOx cytochrome c oxidase

GFP green fluorescent protein

bpy 2,2'-bipyridine

tpy 2,2':6',2"-terpyridine

HPTS 8-hydroxy-1,3,6-pyrenetrisulfonate

HATU O-(7-azabenzotriazole-1-yl)-1,1,3,3-tetramethyluronium

hexafluorophosphate

HOAc glacial acetic acid

DTT dithiothreitol

References

(1) Huang, C.; Mason, J. T. Proc. Natl. Acad. Soc. U. S. A. 1978, 75, 308.(2) Skoog, D. A.; West, D. M.; Holler, J. F.; Crouch, S. R. Fundamentals of

Analytical Chemistry 8th ed.; Thomson Brooks/Cole: Belmont, California, 2004.

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