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Electron Transport Chain in Respiratory Complex I

Introduction Every organism depends on energy to survive, in order to maintain an organized state, homeostasis, through metabolism and other biochemical reactions. Energy is generated in a number of different ways depending on the organism. Mammals create energy through the breakdown of organic molecules, such as carbohydrates, proteins and lipids, that yields other compounds that drives cellular processes. One such compound is ATP (Adenosine triphosphate) an essential energy-carrying molecule that is synthesised by respiration through a series of enzyme protein complexes found in the mitochondria. Complex I (NADH:ubiquinone oxidoreductase) is one of those essential protein complex embedded in mammalian mitochondria. NADH produced by the Krebs’ tricarboxylic acid cycle and β-oxidation of fatty acids, is oxidised to initiate the mechanistic pathway of Complex I, ultimately reducing ubiquinone and establish proton-motive force across the inner membrane of the mitochondria. It is this proton gradient that will support the generation of ATP from ATP synthase and other core processes.
Significant research has been conducted on Complex I, particularly from Bovine heart mitochondria, however to date many aspects of this enzyme is still poorly understood due to its complex structural arrangement and pathways undertaken. To decipher its mechanism, will eventually lead to a greater understanding in the role of Complex I in many diseases and dysfunctions.
Mitochondria Mitochondria are small sub-cellular organelles involved in a series of processes primarily with its role in the respiratory system. Occupying almost 10% to 30% of cell volumes of sizes ranging between 0.75 and 3μm, the unique shape of a mitochondrion allows the process to take place, with its key structural feature being a double membrane.1 These two membranes are separated by the intermembrane space and overall enclose the central matrix. Whereas the outer membrane is inundated by porins to facilitate the movement of solutes of about 12 kDa or less; the inner membrane is impermeable to solutes but presents the ideal environment for the establishment of an electrochemical proton gradient, by the presence of numerous protein complexes.
Additional compartments of the organelle include the cristae and the mitochondrial matrix, which comprises a plethora of enzymes involved in ATP metabolism.
Additionally, a range of studies have also indicated the ability of mitochondria to form dynamic networks of interconnected tubules that regulates the cell structure to adapt to its specific function when required. As a result, during disruption of such networks, cellular dysfunction can occur, leading to a number of neural related syndromes such as Parkinson’s and Alzheimer’s.2,3 Aside from the primary role of energy metabolism, the mitochondria also power other core cellular functions such as apoptosis, calcium handling and the formation of iron sulphur clusters.
The following sections discuss the main enzymes involved in the electron transport chain that lead to the generation of ATP, particularly respiratory complex I, which will be the main focus of this thesis.
Respiratory Complexes Complex II
Also known as succinate: ubiquinone oxidoreductase, complex II is a 120 kDa enzyme consisting of four nuclear-encoded subunits which are arranged in two domains.4 It is this distinctive arrangement which allows this enzyme to oxidise succinate to fumarate which is coupled to the production of ubiquinol through the reduction of ubiquinone in the mitochondrial inner membrane. While it is involved with cofactors, this enzyme complex does not directly contribute to the proton motive force in order to establish a chemical gradient.4,5
Succinate Q → Fumarate QH2
Equation 1:
Two of the enzyme’s subunits SdhA and SdhB form a hydrophilic, ‘succinate dehydrogenase’ subcomplex and forms the succinate/fumarate binding site whereas SdhB contains three iron-sulphur clusters which are embedded to the mitochondrial membrane by the remaining SdhC and SdhD subunits.4 These latter subunits contain a heam group and ubiqionone binding sites. When a flavin dinucleotide, which is ligated to SdhA, it oxidises succinate, the electrons produced in this process are passed down through the iron-sulphur clusters. The electrons subsequently allow the reduction of ubiquinone to ubiquinol.6,7
Complex III
Complex III or ubiquinol:cytochrome c oxidoreductase is an 240 kDa enzyme which is made up of 11 subunits. Its structure comprises of two ubiquinone binding sites; Qo, present towards the mitochondrial membrane, catalyses the oxidation of ubiquinol to ubiquinone and Qi, present towards the matrix, catalyses the reduction of ubiquinone to ubiquinol.8,9
Complexes I and II produces ubiquinol from the reduction of ubiquinone, which binds to the Qo site on complex III. During this process, an electron is passed along the iron-sulfur cluster reducing it and moving it towards cytochrome c1 and cytochrome c resulting in a conformational change. The change causes a second electron to be transferred through another pathway formed of cytochromes bL and bH towards to Qi binding site, in where it allows the formation of a semiquinone anion through the reduction of an already bound ubiquinone. Parallel to this, a second quinol is oxidised at Qo allowing the electron to be transferred through the first pathway of Rieske iron-sulphur cluster and cytochrome c1 and the second electron follows the second pathway mentioned above to Qi, reducing the semiquinone anion to ubiquinol.10 The oxidation at Qo releases four protons into the inter-membrane space of the mitochondria and the reduction at Qi results in the uptake of two protons from the matrix which are transferred into the inter-membrane space during ubiquinol oxidation. This complete cycle allows the reduction of two cytochrome c molecules.9
QH2 2 cyt c3 2H in → Q 2 cyt c2 4H out
Equation 2:
Oxidation and reduction cycles in Complex III results in the movement of four protons into the inter-membrane space maintains the proton motive force used by ATP synthase to synthesise ATP.8
Complex IV
Complex IV, also known as cytochrome c oxidase, is an enzyme, which comprises of 13 subunits, of which three are encoded by the mitochondrial genome. The enzyme catalyses the oxidation of cytochrome c which leads to the reduction of oxygen to water allowing the translocation of four protons across the mitochondrial inner membrane.11,12
The oxidation of cytochrome c produces electrons that are transferred to an active site where molecular oxygen is reduced. This reduction producing water releases free energy required for the pumping of four protons from the matrix of the mitochondria into its inner-membrane space. This movement of protons is facilitated through two known proton channels: the K-channel passes two protons for the reduction of oxygen and the D-channel allows the movement of newly translocated protons.13
O2 4 cyt c2 8H in → 2 H2O 4 cyt c3 4H out
Equation 3:
The translocated protons and the reduction of oxygen to water allows ATP synthase to generate ATP as this contributes to the proton motive force similar to Complex III.
Complex V
Primarily known as ATP Synthase, this enzyme complex operates by utilising the proton chemical gradient established in the intermembrane space by the preceding complexes, to drive the synthesis of ATP from ADP and inorganic Phosphate. With an average size of 580 kDa, the enzyme is composed of 16 subunits organised in two hydrophobic and hydrophilic domains; the hydrophobic domain forms a proton conductive pore through the inner membrane while the hydrophilic domain, containing three copies of α and β subunits, spreads into the matrix. The two domains are linked by an asymmetric central stalk and a peripheral stalk, which acts as a stator to prevent the F1 domain rotating freely during catalysis. The interfaces between the two subunits forms the binding sites for ADP and inorganic Phosphate. 14,15
ADP P nH in → ATP nH out
Equation 4
Complex I
Complex I, is the first and largest enzyme involved the electron transfer chain of the mitochondrion. Alternatively known as NADH:ubiquinone oxidoreductase, its primary role is to oxidise NADH and ultimately reduce ubiquinone.16
NADH H Q 4H in → NAD QH2 4H out
Equation 5:
Just like the other protein complexes, the potential energy released from the redox reaction within the complex, translocates four protons across the inner membrane for every molecule of oxidized NADH and removes two additional protons from the matrix for the reduction of quinone. The processes contribute to the overall electrochemical gradient which is to be used by ATP synthase to synthesise ATP.17
Structure To date, complex I has been found in a variety of species, including many prokaryotes. The complex I from bovine heart mitochondria is primarily used in studies due to its close sequential identity with the human complex I enzyme. The mammalian complex I is one of the most complex and largest enzymes known, with a combined mass of 980 kDA and composed of at least 45 different polypeptide subunits; with 14 strictly conserved core subunits that are necessary for function and also common across the among all known complex I.16 The significance of the additional subunits in complex I among different species still remain a mystery. It is known some be involved in protection against reactive oxygen species generation and some are required needed for proper assembly and stability of the enzyme.16,18
As observed by single-particle electron microscopy (EM) for both bacterial and mitochondrial enzymes, the determined structure of the enzyme closely resembles to an “L” shape, with seven hydrophobic core subunits that constitutes the membrane tail domain and seven hydrophilic core subunits that constitutes peripheral (hydrophilic) arm domain protruding into the mitochondrial matrix; which is known as the catalytic domain as it includes all redox centres and binding site while the membrane domain consists mostly of hydrophobic subunits. 16
While the full structure of the eukaryotic complex is not still well characterised, in 2006, Sazanov group successfully reported structure of the hydrophilic domain of complex I from Thermus thermophiles bacteria.20
The Peripheral Arm of complex I
The peripheral arm of the complex is composed of seven individual subunits, that together, houses the NADH-oxidizing dehydrogenase module, which provides electron input into a noncovalently-bound flavin mononucleotide (FMN) molecule. The molecule sequentially transfers the electron to a chain of nine iron-sulphur (Fe-S) clusters, eight of which are found in the bovine enzyme. Additionally, the hydrophilic arm also comprises of a Q-module, which conducts electrons to the quinone-binding site for quinol production. 16,20
All of these
Within the respiratory chain complexes, there are three different types of Fe-S clusters, two of which, are found in complex I; Two binuclear [2Fe-2S] and six tetranuclear [4Fe-4S] clusters.
As the name suggests, the binuclear clusters are composed of two iron atoms that function as bridged by two acid-labile sulphur atoms. Each iron atom is also coordinated by an additional two sulphur atoms found on the surrounding cysteine residues from the protein complex. In the tetranuclear Fe-S clusters, four iron atoms and four sulphur atoms are arranged in a cube with each iron atom also ligated to sulphur cysteine-residue on the surrounding protein, similar to binuclear Fe-S.22
Due to their conformational arrangements and redox capabilities provided by the iron atom, these clusters act as electron transfer agents or also known as ferrodoxins. The detection of these clusters can be achieved by EPR (electron paramagnetic resonance) which is successfully achieved in many studies. However, out of the two binuclear and six tetranuclear iron-sulfur clusters found in complex I, only two binuclear and four tetranuclear clusters are EPR active.22

Figure 1.: structures of the iron-sulphur clusters found in complex I.
As previously mentioned, seven of the eight clusters, form a 95 Å-long extensive chain directly from the flavin site to the quinone binding site on the interface of the membrane domain. Even though the distances between these chains may seem far apart, as much as 14 Å, distances are close enough to allow electron transfer to occur.23,24
However, the presence of the eight cluster is still not well understood. Cluster 2Fe[24] found on the opposite side of the Flavin site, is believed not to be involved in electron transfer pathway. While it was just a theory with no evidence, it has been proposed that this additional cluster functions as an electron store that accepts an electron from the flavosemiquinone species preventing the generation of reactive oxygen species during enzyme turnover.24
Membrane Domain of complex I
The membrane domain comprises the proton-translocating module which catalyses proton transport. With the exception of subunit ND1 and the quinone binding site, found on the interface of the peripheral arm, the membrane domain functions totally independently from the two arms of complex I.
Within the membrane domain, there are four structural subunits that have been identified to be possibly involved with proton translocation; these include subunits ND2, ND4 and ND5. There is also an additional transporter which believed to be either ND1, ND6 or ND4L. Each believed to be transporting one proton per catalytic cycle. Each individual subunits are composed of charged residues and helices that creates half-channels that allow the passage of proton to occur. The membrane structure is also held together by a long α-helix chain that spans across its entire length. Its feature is to maintain and support the integrity of the membrane domain.26
Overall Mechanism of complex I
The mammalian complex I includes 45 known proteins, out of which 14 “core” subunits comprises of both hydrophilic and hydrophobic domains as explained above.16
The mechanism through the electron transfer chain starts with a Flavin mononucleotide (FMN) molecule which is non-covalently bound to the 51kDa subunit through hydrogen bonds at the top of the hydrophilic domain. FMN molecule oxidises NADH leading to the reduction of iron-sulphur clusters (Fe-S) which transfers electrons from Flavin to the quinone-binding site {51}. This electron transfer distorts the conformation of the protein through changes in its redox state leading to alterations in pKa values of its side chains; these alterations allows four hydrogen ions being pumped out of the mitochondrial matrix.24
It is believed NADH gets oxidised to NAD through a hydride transfer avoiding the formation of the unstable NAD. Radical.24 This oxidation process occurs when the nicotinamide ring of the NADH lies above the flavin isoalloxazine system, allowing the electron donor hydride (C4 of the 27 nicotinamide ring) and acceptor (N5 of the flavin) to come within 3.5 Å of each other and transfer electrons.28

As explained above, NADH oxidation leads to transfer of electrons through seven iron-sulphur clusters chain between Flavin and quinone reduction binding site in the membrane.20 It is the final Fe-S cluster that donates the electrons to the bound ubiquinone substrate which is believed to be accessed through an entry point in the membrane to the binding site.21
These iron-sulphur clusters are best detected using a technique called electron paramagnetic resonance (EPR). Previous studies have observed five reduced Fe-S clusters through EPR from Bovine compliex I reduced by NADH, and their spectra are represented N1b, N2, N3, N4 and N5.25 This technique will be further explained throughout this thesis.
A much recent study by Roessler et al. (2010) used EPR to understand the tunnelling electron transfer pathway through these clusters. Previous studies have already established EPR signals N1b, N2 and N3 are detected from 2Fe cluster in the 75 kDa subunit (position 2), and from 4Fe clusters in the PSST (position7) and 51 kDa subunits (position 1) respectively along the clusters chain due to interactions with ubisemiquinones and flavosemiquinone. As the other EPR signals have yet failed to be assigned to a particular cluster, Roessler et al. (2010) went on to use double electron-electron resonance (DEER) spectroscopy to detect N4 and N5. Their results demonstrate that N4 is assigned to the first 4Fe cluster in the TYKY subunit (position 5), and N5 to the all-cysteine ligated 4Fe cluster in the 75 kDa subunit (position 3).25
The study propose an alternating energy potential profile for electron transfer along the chain between the actives sites, in B.taurus, which enhances the rate of a single electron travelling through the empty chain subsequently leading to more efficient energy conversion in complex I.25
Followed by the iron-sulfur cluster is the site of quinone reduction. A study performed by Sazanov and Hinchliffe has identified a supposed binding site for the quinone head group from T. thermophilus complex I hydrophilic domain between the 49 kDa and PSST subunits.20 This alleged site is close to the cluster where the ubiquinone substrate accepts electrons from the chain and it has also been acknowledged the 49 kDa and PSST subunits play an important role in quinone binding and catalysis.29
Nevertheless, it is believed that additional hydrophobic subunits may also be involved in quinone binding and these are still being investigated.
Even though the mechanism of NADH oxidation and ubiquinone reduction is relatively well understood, how this oxidoreduction leads to quinone reduction and subsequent protons pumping across the mitochondrial membrane from complex I still remain a mystery. A number of theories for complex I mechanism have been proposed based on the proton-pumping systems of the other mitochondrial respiratory complexes. These theories have been outlined below:
A direct coupling mechanism – as demonstrated by complex IV through cytochrome c oxidase where the proton transfer is determined by a gating reaction occurring at the same time as the electron transfer reaction that started it.30
An indirect coupling mechanism – as seen in complex V (ATP synthase) explained previously. A study performed by Efremov et al., suggests that within complex I, one proton is translocated by a directly coupled mechanism at the Fe-S clusters and the rest are moved when quinone reduction drives conformational changes to the four-helix bundle of Nqo4 and of Nqo6 in complex I, subsequently affecting the C-terminal helix of Nqo12. The C-terminal has been identified by the authors running parallel to the membrane. The effect on this helix consequently leads to the other three helices to tilt which results in proton translocation.31
A Q-cycle-like mechanism – as represented by complex III where quinol is used as a carrier to transport protons across the mitochondrial membrane. A study completed by Dutton and co-workers suggested the complete reverse of this mechanism for complex I featuring the presence of two ubiquinone binding sites; one facing the inter-membrane space, Qo, and the other facing the mitochondrial matrix, Qi. The quinone substrate would bind at Qi, and be reduced by one electron from a quinol already bound at Qo and another electron from the Fe-S cluster; subsequently leading to two protons being taken up from the matrix while the formed semiquinone specie is still bound at Qo. Following the uptake of the protons, semiquinone is oxidised to ubiquinone.32 Nevertheless, further studies conducted have found no evidence of ubiquinol oxidation signifying complex I do not work through this mechanism.30,33
While the first isolation of complex I from bovine heart mitochondria by Joe Hatefi et al occurred 40 years ago, information on its overall mechanism of action is still very limited particularly the mechanism of redox-proton coupling occurring in the membrane domain. To further understand this, new studies are being conducted to trap radical intermediates formed at the interface of the peripheral and membrane arm to establish the pathway that initiates proton translocation.
Semiquinone radicals Semiquinones are catalytic intermediates formed within complex I during the reduction of quinones at the quinone binding site and can exist in neutral or anionic form. Due to the presence of the unpaired electron, semiquinone intermediates can be studied using EPR spectroscopy.

There are numerous pathways in which the formation of semiquinones can occur from quinone. The scheme below, proposed by Roessler and Hirst, illustrates the three main possible routes taken to obtain quinol.
Pathways A and B involves with the generation of a neutral semiquinone radical specie based on the transferring of a proton and electron. On the other hand, pathway C which follows through pathway B involve with the generation of an anionic radical specie generated from an electron transfer. All pathways lead to formation of quinol by series of electron transfer and protons. The pathway shown in grey which occurs from the protonation of the neutral semiquinone radical specie will result in a 1-electron-2-centre bond which are energetically unstable.27
Aside from one study, majority of the studies till date, have proved the existence of semiquinones by observing EPR signals using submitochondrial particles (SMPs). As the name suggests, these are inverted membrane vesicles housing the entire electron transport chain containing all enzyme complexes.34 However, since quinone cofactors are used by majority of the other complexes, distinguishing the semiquinone signals with each complex, has been far from successful.
More recently, there has been a wave of research focusing on the identification of semiquinone radicals exclusively from complex I, however these have proved even more challenging as the organic intermediates produced very low intensity signals.
Within complex I, there are two species of semiquinone that have been identified; SQNf and SQNs.35,36 Based on their EPR properties, SQNf or fast relaxing semiquinones has been reported only during the presence of an established proton gradient across the membrane. On the other hand, SQNs or slow relaxing semiquinones, are not effected by proton gradient. The presence of two semiquinones has also lead to the possibility of complex I to contain two separate quinone binding sites; Due to SQNf having a spin-spin interaction with Fe-S cluster N2, it is theorised that SQNf binding site is located close to the cluster at around 12 Å estimated distance, in contrast, SQNs binding site is suggested to be located around 30 Å from N2 cluster.22,25,37
Within the complex, the SQNf is believed to be involved in proton pumping and its site aids the system by acting as bound co-factor site that facilitates the transfer of one electron from one site to another allowing the formation of a binding pocket for the SQNs in equilibrium with the ubiquinone pool of the membrane.22,25,32,35,38
The presence of two separate quinone binding sites still remains a mystery and cannot be totally ruled out even though it has been suggested that SQNf and SQNs signals are detected from the same semiquinone species located from different sites or present in catalysis states.39
A recent potential way of observing semiquinone intermediates via EPR is through the use of liposomes. Liposomes containing just Complex I or proteoliposomes, will facilitate the capture of semiqinone within its native environment and hopefully provide an insight in the mechanism of Complex I and the binding of Q10.
Liposomes Liposomes are spherical nanovesicles used in a variety of applications. Composed of a phospholipid bilayer, these small vesicles have an aqueous solution core surrounded by a hydrophobic membrane. Hydrophobic chemicals associate with the bilayer while the hydrophilic solutes dissolved in the core cannot readily pass through the bilayer; essentially mimicking the cellular phospholipid bilayer. Due to these features, liposomes can be loaded both with hydrophobic or hydrophilic molecules and are excellent drug carriers or in this case house protein complexes. Liposomes are also not naturally occurring and must be artificially generated using lipid extracts by aggregating them.40
As liposomes are formed from naturally occurring lipids of low intrinsic toxicity, they are biodegradable and non-toxic. The functionality of liposomes is dependent based on three main factors. These include: size, bilayer composition and liposome surface properties.40
Phospholipids are one the essential components in the formations of liposomes and can be divided into synthetic and natural phospholipids. They consist of two fatty acids hydrophobic chains linked to a hydrophilic (polar) head group, and they have either glycerol or sphingomyeline as the back bone. Having both hydrophobic and hydrophilic components, make phospholipids having amphipathic molecules.41 The diversity of the hydrophilic head group molecules and hydrophobic chains’ length allows the formation of different phospholipids which affects the surface charge and bilayer permeability of the liposomes.40
The length and degree of saturation of the hydrocarbon acyl chains determines the stability of the liposomal membrane, by affecting the temperature at which the membrane changes from a “closely packed gel phase” to a “fluid phase”. The surface charge of the liposomes is determined by the charge of the lipid forming it which can be altered by modifying lipids with hydrophilic moieties to membrane bilayers.40
Liposomes can be composed of naturally-derived phospholipids such as cholesterol, one of the commonly used lipids in liposome formation. It enhances the stability of the lipid bilayer and form highly ordered and rigid membrane with fluid like characteristics. Other phospholipids, synthetic and non-synthetic, can also be used for the formation of the liposomes such as pure surfactant components like DOPC (1,2-dioleoyl-sn-glycero-3-phosphocholine).42
Classifications of liposomes Liposomes are classified according to their morphological sizes and lamellarity, depending on their composition and method of formation.40
Multilamellar vesicles (MLVs) – consists several concentric phospholipid bilayers or lamellar ranging between 100nm to 20 µm in size depending on the method of preparation. These large bilayers allow the integration of lipophilic molecules and proteins.
Small unilamellar vesicles (SUVs) – single phospholipid bilayer and sized between 20 nm to 100nm. Ideal for encapsulation small compounds and proteins.
Large unilamellar vesicles (LUVs) – single phospholipid bilayer with size ranging from 100 nm to 1 µm. They are known to have larger aqueous core compared with or MLVs, making them suitable to useful to load with numerous compounds.
Oligolamellar vesicles (OLVs) – vesicles similarly structured to MLVs but consists of anywhere between two and five phospholipid bilayers.
Multivesicular liposomes (MVLs) – When a large liposome vesicle similar in size to an MLV, enclose a group of liposomes, then the subsequent vesicle is known as multivesicular liposome (MVL).

Figure 1.40
The current state of research on liposomes have primarily been focusing on the administration of drugs and other compounds to biological systems since it overcome challenges associated with reaching the target, making them very useful in the cosmetic and pharmaceutical industries.40
Furthermore, it should be noted, some surfactant based phospholipids can mimic the biological systems helping construct important model systems for the research on enzymes and membranes. Many recent publications concerning liposomes have been focused on using this mimetic chemistry, which deals with models, mimicking cellular membrane to facilitate the research into their structures as well as the mechanisms both in vivo and in vitro.40
Aims of Project The current state of research on complex I remain largely focused on the determination of the mechanism since only a fraction has been found. Fully understanding will help solve many diseases and other complication caused by complex I.
Whereas the mechanism of the reactions between NADH and iron sulphur clusters have been established, little is known about the mechanism of proton translocation as well as the role and existence of semiquinones that will lead into revealing more information into the function of the enzyme. The work described in the following records, using the best technique available, EPR, will aim to be using current studies of using liposomes to mimic cellular conditions, similar to the mitochondrial membrane, for complex I in order to obtain data regarding reduction of Q10 and proton translocation.

Materials Preparation of Complex I from Bovine Mitochondria
Preparation of Complex I proteoliposomes
Stock solutions of 25 mgmL-1 of POPC in chloroform was transferred to a glass homogeniser with the required amount of ubiquinone-10 contained in chloroform. The chloroform was removed under Argon. An alternative approach is to remove under vacuum using rotary evaporator. The resulting phospholipid film was resuspended in 675 μL of buffer (10 mM Tris-SO4 (pH 7.5) and 50 mM KCl), and extruded 25 times through a Whatman 0.1 μm pore membrane. The liposome mixture was solubilised with the addition of 160 μL of octyl-glucoside from an aqueous 10% stock solution, sonicated for 10 min, and further incubated on ice for 10 min. The following steps were carried out at 4 °C. 0.2 mg of AOX (50 μL of 7.8 mgmL-1) and 0.2 mg of complex I (10 μL of 20 mgmL-1) were added to the solubilised lipids and incubated for a further 10 min, followed by the addition of 100 μL of SM2 Biobeads. The mixture wa

Construction of Recombinant DNA in E Coli

In 1973 Stanley Cohen and Herbert Boyer pioneered the use of recombinant DNA technology for cloning and expression of genes in foreign organisms. They cloned DNA from the Salmonella typhimurium streptomycin resistance plasmid RSF1010 into the Escherichia coli plasmid pSC101 and observed tolerance to streptomycin among the transformants (Cohen et al., 1973). The first reported production of a human recombinant protein took place a few years later when the then newly started biotech company Genentech announced that they had managed to express the gene encoding human somatostatin in E. coli (Itakura et al., 1977). The value of the resulting bioactive substance was similar to that of somatostatin extracted from the brains of 500.000 sheep. In 1982 Genentech followed up this success with the product humulin, a recombinant insulin produced in E.coli and the first recombinant biotech drug to be accepted for market by the Food and Drug Administration. Today the production of recombinant proteins has become a huge global industry with an annual market volume exceeding $50 billion (Schmidt, 2004). At the start of the recombinant protein expression era the bacteria Escherichia coli and Bacillus spp. dominated as hosts for recombinant expression, but the realization that a protein may require a specific host physiology and biochemistry for optimal production stimulated a search for new hosts, both prokaryotic and eukaryotic. Parallel to this quest, recombinant DNA technology advanced tremendously thereby opening up possibilities for the use of novel organisms. As a consequence, many different expression systems for use in many different hosts are now available, including systems for use in yeasts (Gellissen et al., 2005), filamentous fungi (Nevalainen et al., 2005), insect and animal cell cultures (Wurm, 2004; Kost et al., 2005), gram-positive bacteria like Bacillus (Westers et al., 2004) and Streptomyces (Binnie et al., 1997), and gram-negative bacteria like Escherichia coli
Bacterial expression systems are the preferred choice for production of many prokaryotic and eukaryotic proteins. The reasons for this lie in the cost-effectiveness of bacteria, their well-characterized genetics, and the availability of many different bacterial expression systems. Among the hosts available for recombinant expression, Escherichia coli is in an exceptional position. This stems from the many decades of intense researchon its genetics as well as the broad scope of biotechnological tools available for genetic engineering of this organism. As a host for recombinant expression, E.coli is especially valued because of its rapid growth rate, capacity for continuous fermentation, low media costs and achievable high expression levels (Yin et al., 2007). One consequence of this popularity is that about 80% of all proteins used to solve three-dimensional structures submitted to the protein data bank (PDB) in 2003 were prepared in E.coli (Sørensen and Mortensen, 2005) and during 2003 and 2006, nine out of 31 approved therapeutic proteins were produced in E.coli (Walsh, 2006), among them important growth factors, insulins and interferons (Schmidt, 2004).
Green Fluorescent Protein (GFP) was isolated from the jellyfish Aequorea aequorea in 1962 (Shimomura et al., 1962) where it was found as a companion protein to aequorin, the well-known chemiluminescent protein of the same species. It was noticed that living A. aequorea tissue had an emission spectrum peaking at 508nm and looking green but pure aequorin peaked in the blue range, at 470nm (Tsien, 1998). This then led Shimomura’s group to discover GFP and suggest radiation-less energy transfer as the mechanism for exciting the protein. Its structure has been determined to consist of an 11 stranded β-barrel containing the chromophore made up of a single α helix as shown in Figure1.
Its use as a tool in molecular biology was not realised until 1992 when Prasher reported the cloning and sequence of GFP (Prasher et al., 1992). Since 1994 GFP has been used as a reporter protein (Chalfie et al., 1994) flagging its own presence and therefore also proteins under the same control, by emitting green light (λem = 508 nm) upon excitation with near ultraviolet light (around 395 nm) or blue light (around 470 nm) (Ito et al, 1999). Since then many mutations have been developed looking to improve the emission or to focus it to a single wavelength (Heim et al., 1995) or to change the color of the emitted light itself.
Recombinant DNA molecules usually contain a DNA fragment inserted into a bacterial vector.
Polymerase chain reaction (PCR), a specific gene or DNA region of interest is isolated and amplified by DNA polymerase extracted from a heat-tolerant bacteria. PCR “finds” the DNA region of interest (called the target DNA) by the complementary binding of specific short primers to the ends of that sequence. The long chromosome-size DNA molecules of genomic DNA must be cut into fragments of a much smaller size before they can be inserted into a vector. Most cutting is done with the use of bacterial restriction enzymes. These enzymes cut at specific DNA sequences, called restriction sites, and this property is one of the key features that make restriction enzymes suitable for DNA manipulation. These enzymes are examples of endonucleases that cleave a phosphodiester bond (Anthony, 2012). The key property of some restriction enzymes is that they make “sticky ends. The restriction enzyme EcoRI (from E.coli) recognizes the following sequence of six nucleotide pairs in the DNA of any organism:
5-GAATTC-3
3-CTTAAG-5
The enzyme EcoRI makes cuts only between the G and the A nucleotides on each strand of the palindrome (Figure.2).

The recombinant DNA molecules are transferred into bacterial cells, and, generally, only one recombinant molecule is taken up by each cell. The recombinant molecule is amplified along with the vector during the division of the bacterial cell. This process results in a clone of identical cells, each containing the recombinant DNA molecule, and so this technique of amplification is called DNA cloning. The next stage is to find the rare clone containing the DNA of interest.
Bacterial plasmids (vectors) are small circular DNA molecules that replicate their DNA independent of the bacterial chromosome. The plasmids routinely used as vectors carry a gene for drug resistance and a gene to distinguish plasmids with and without DNA inserts. These drug-resistance genes provide a convenient way to select for bacterial cells transformed by plasmids: those cells still alive after exposure to the drug must carry the plasmid vectors. However, not all the plasmids in these transformed cells will contain DNA inserts. For this reason, it is desirable to be able to identify bacterial colonies with plasmids containing DNA inserts. Such a feature is part of the pUC18 (or pUC19) plasmid vector shown in Figure 2; DNA inserts disrupt a gene (lacZ) in the plasmid that encodes an enzyme (-galactosidase) necessary to cleave a compound added to the agar (X-gal) so that it produces a blue pigment. Thus, the colonies that contain the plasmids with the DNA insert will be white rather than blue (they cannot cleave X-gal because they do not produce -galactosidase).

The following experiment outlines the construction of recombinant protein production in E.coli strain BL21 by using a bacterial plasmid vector pUC18/19 expressing Green Fluorescent Protein (GFP) to act as a recombinant protein product with the benefits of being easy to visualise and measure.
Materials and Methods Materials:
The experiment was carried out using the following materials and Equipments: 2µl EcoRI/HindIII cut and cleaned PUC19 vector, 5µl EcoRI/HindIII cut and cleaned GFP insert, 2µl 10xT4 ligase buffer, 2µl T4 ligase(0.5 U ml-1) , and 9µl sterile water (H2O) ]to make up to 20µl volume[ .
100µl of competent BL21 E.coli cells on ice, 42°C water bath, Ice bucket with ice, selective media plates (1.5% Luria broth (LB) Agar, 40µg mL-1 X-gal, .1 mM IPTG, 50µg mL-1 ampicillin), sterile tubes, shaking incubator, Spectrophotometer or similar device to measure optical density of the bacterial cultures, flasks, Microcentrifuge.
Methods:
It can be divided into three stages:
Ligation Reaction stage: in this stage 2µl EcoRI/HindIII cut and cleaned PUC19 vector, 5µl EcoRI/HindIII cut and cleaned GFP insert, 2µl 10xT4 ligase buffer, 2µl T4 ligase (0.5 U ml-1) , and 9µl sterile water (H2O) are mixed and kept at room temperature for at least 30 minutes.
Transformation of ligation into cloning host stage: this stage conducted by deforesting 100µl of competent BL21 E.coli cells on ice (with caution do not allow to warm to room temperature), then adding 10µl of the ligation reaction from the first stage to BL21 E.coli cells. They are then incubated for up to 30 minutes on ice. Next step, is done by taking out the transformation mixture out of the ice and heated in water bath at 42 °C for almost 75 seconds, then followed by return immediately into ice for a minimum of 2 mins. Then the cells were plated out on selective media plates (1.5% Luria broth (LB) Agar, 40µg mL-1 X-gal, .1 mM IPTG, 50µg mL-1 ampicillin). Lastly, the transformation mixture is incubated at 37 °C for 12-18 hours afterdriedd.
Picking of colonies for the protein expression stage: 2x5ml LB 50µg ml-1 ampicillin in 30ml sterile tubes were prepared, then 1xBlue individual colony and 1x white individual colony selected and inoculated in separate tubes. Then the tubes were incubated with shaking incubator throughout the night at 37°C , speed: 220rpm.
Subculture and Growth of Recombinant E.coli for Protein expression: At the beginning, 2x60ml sterile Luria-Bertani (LB), in 250ml conical flask were warmed , (1 per inoculums ) at 37 °C, Then aseptically the ampicillin was added to a last concentration of 50µg ml-1 ampicillin. Next 1 ml of media was removed and was put in a cuvette to act as blank (one blank is enough for both ouh), followed by addition of 600µl overnight to calture of each individual colony to separate flask (1:100 inoculum), the flasks were put back to the shaking incubator and incubated at 37°C, speed: 200rpm , after that blank spectrophotometer was placed against media at 600nm , after 45 minutes the samples were removed aseptically from flasks, then from every flask 1x 1mL was removed and added to a fresh clean cuvette (take to next step 8) and 1x1ml was added to clean Eppendrof (take to step 9) . The OD600nm of culture in cuvette was Measured and the result of growth curve was recorded (once the culture has reached an OD 600nm of 0.5, IPTG was added to final concentration 1Mm stock solution. Then samples were spun down in the Eeppendrof tube at max speed in Microcentrifuge for 5 minutes , ensure centrifuge is balanced before spinning , the supernatant was removed and pellet ,then the pellet was suspended in 200µl Cell lysis buffer (10mMl Tris PH8.0, 300Mm Nacl , 10mg ml-1 Lysozyme). Resuspended cells were frozen at -20 c to the next day. Lastly, sampling was continued until OD600nm is no longer rising for two successive samples or until 16:30 pm.
Results and discussion Although it is supposed to harvest between 30-300 colonies per plate (210- 2100 colonies for all groups), just three blue colonies were observed in plates between all groups, which mean that protein of interest (GFP protein ) was not expressed (inefficient) in BL21 E.coli cells due to some factors influenced the expression level or to some technical problems during the experiment which will be discussed.
The most popular strain, BL21 and its derivatives, which are good producing protein, are descended from E.coli B and thus is deficient in the Lon protease. Additionally, the BL21 background lacks the OmpT outer membrane protease. For expression work, BL21 cells should be taken from stock cultures that performed from fresh transforms. This step is crucial to insure that the clone does not change and that each expression run gives optimal performance.
Transformation frequency is affected by the purity of the DNA, how the cells are handled, and how the transformation was actually performed. In the impurities in the DNA usually spin columns can be used to purify DNA from PCR reactions, ligations, endonuclease digestions, or other treatments. In addition, the most common mistake when transforming E.coli is to put a lot of ligation mixture in the transformation.
Other factors that effect transformation with BL21 are the handling and the storage of the competent cells. Competent cells need to be reserved at -70°C to keep them at the peak .It is worthy of noting that 5-10-fold of efficiency usually lost if tube put back in the box and place in the freezer. Moreover, Cells must be thawed on ice, and the transformation should be started immediately after the cells are thawed. Incubating on ice is necessary for chemically competent cells. If you heat shock right away, the efficiencies will be down 10-fold. If incubate for only 15 minutes, it will be down 3-fold. In addition, time of heat shock (75 second) could be not enough , thus, affect the efficiency enough to transformation of E.coli. Moreover, water bath temperature may be not equilibrated (less than 42°C or a higher which decrease in transformation efficiency ( Smith, et al, 1992).
Also, the concentration of DNA has significant effect on the transformation efficiency , usually less amount of DNA is used. If using more, the result is fewer colonies because the impurities in the DNA will inhibit some of the cells from being transformed.
There are main factors to consider during induction conditions: Vector, Host Strain, and Growth Conditions. These three factors have the biggest impact on the expression of the protein of interest. First on the list of considerations is the vector that is used to express GFP protein. The first thing should be considered after cloning, the protein of interest is still in frame. It is recommended that before any experiment is carried out the first thing is should be done is cloned plasmid (or a few different clones) sequenced. This will show if the sequence you inserted into the expression vector is still correct and is still in frame. This is especially important if the construct contains any PCR fragments. If there are any point mutations or the sequence gets out of frame by even a few bases it can have dramatic effects on the protein that expressed. Another thing to check before expressing is if the GFP protein sequence contains long stretches of rare codons. This can cause the protein that is expressed to be truncated or non-functional. A few rare cordons spread around the protein are OK in most cases, but if there are a number of rare codons in a row, then it can have a big effect. The third sequence related step to optimize the protein production is to make sure there is not a high GC concentration at the 5′ end of GFP protein. This could potentially cause problems with the mRNA’s stability, and could prevent it from being translated correctly, which would also lead to truncated or non-functional proteins. If your sequence is GC heavy at this end, you can try to make a few silent mutations to break up long stretches to try and help stability.
After the plasmid is sequence verified, the next factor is the bacterial host that is used. There are almost as may hosts as there are expression vectors, with certain hosts excelling in producing different types of proteins. For example if you have a toxic protein, or a protein that could potentially cause genomic rearrangement, you will want a vector that gives you very tight control over the induction of your protein. There can be “leaky” expression (i.e. expression of your protein without the addition of your inducer) that can potentially have adverse effects on the cells growth or even prevent your cells from over-expressing your protein in the first place. If you’re utilizing the T7 polymerase system, then look for a host containing the pLysS plasmid, as this will code for T7 lysozyme, which will suppress the T7 polymerase and can greatly reduce the level of background expression. If as stated before you have a protein that contains a large number of rare codons, then look for a host with the genes for the necessary tRNA’s already present, which should allow your protein to express correctly. Sometimes simply changing hosts can have a dramatic effect on the amount of protein produced and the stability of the protein that is made, so if one host isn’t giving you the results you need, then feel free to switch your host up.
The third and final factor to consider when expressing a protein is growth conditions. When first starting out with the protein induction it is very important to run an expression time course, where you take a fresh colony from a streaked plate, and grow the culture to stationary phase. Next, dilute the overnight culture 1/100 and grow to mid log phase, then add the inducer and induce your protein for a number of hours, taking 1mL samples every hour or so. Once these samples are lysed, you can run an SDS-PAGE gel to determine your protein production levels. You might get great induction the first time, or you may have to tweak your conditions in order to get really good expression levels. Other factors that may need to be controlled for are the bacterial growth rate (determined by taking OD measurements during the induction process), and the temperature during induction. Some constructs will express perfectly fine at 37°C, while others need to be bumped down to 30°C to induce correctly. The concentration of the inducer too will have an effect, as many inducers (IPTG) can be toxic to the cells that they are inducing. Using freshly made inducer is good step to making sure you always have consistent results. Only through experimentation can you determine what will be best for your construct, and give you the most robust expression levels.
Transformation efficiency:
Transformation efficiency is a measure of the ability of cells to be transformed. Transformation efficiency is expressed as the number of transforms per microgram of pUC19.
By using the following formula:
Colonies on plate / ng of control DNA X 1000ng/µg = (transformation (T) / µg plasmid DNA)
100 μL equivalent to 0.01 ng DNA in the plate.
Growth curve
In general growth curve shows the S- shaped when plotted in log linear format as shown in figure 4, that separated into four phases:

Lag phase; the initial period when no increase in cell number is seen.
Log phase; when cells are growing at the maximumm rate.
Stationary phase; growth decreases as a nutrient are depleted and waste products accumulate.
Death phase; this is the result of prolonged starvation and toxicity.
Conclusion The main goal for the experiment was to express the protein of interest (GFP). However, factors influencing transformation efficiency include technique errors, the temperature and length of the incubation period, the growth stage of the cells, and using the correct mass of plasmid DNA. Escherichia coli is one of the most important hosts in modern day recombinant protein production. Throughout academia and industry its uses are widespread and with sequence data available for some of the most common strains of the bacteria it has been a favourite organism for many metabolic engineering and metabolic modelling projects in the past (Berry, 1996; Koffas et al., 1999).

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