Cyanobacteria constitute the most basic beings ever to appear on our planet. Their ability to use water as an electron source in photosynthesis contributes largely to the balance between CO2 and O2 in the atmosphere. Carbon dioxide (CO2) is not only a greenhouse gas increasing in the Earth’s atmosphere, but also a key metabolite in living organisms. It plays an essential role in such fundamental biological processes as respiration and photosynthesis. It is known that CO2 reacts with neutral amines at physiological temperatures and pressures to form carbamates. Carbamylation is caused by the nucleophilic attack of an uncharged amine (lysine side chain 3-amino group or N-terminal a-amino group) on CO2 (Fig.1). This post-translational modification has been reported on proteins like RuBisCO and haemoglobin but unexplored at other cellular systems as cyanobacterial CO2 uptake. The objective of this study is to identify carbamates within cyanobacterial metabolism using a chemical trapping technique recently developed and to investigate how this post-translational modification impact in cyanobacteria response to CO2.
Fig. 1. Reversible nucleophiic attack of an uncharged amine on CO2 to from a carbamate
Cyanobacteria CO2 uptake
Environmental oxygen it is due to cyanobacterial CO2 uptake metabolism development. The remarkable decadence in dioxide carbon levels and the increase in oxygen levels that occurred around 350 million years ago may be cause by cyanobacteria adaptation to face the situation with photorespiration and low efficiency carbon gain, for example transport mechanisms for the active uptake of inorganic carbon (Ci) and the resulting rise of ribulose bisphosphate carboxylase-oxygenase (Rubisco) and the division of Rubisco into micro-compartments known as carboxysomes (1).
These micro-organisms are the only photosynthetic prokaryotes able to produce oxygen. They lack membranes around their organelles and their respiratory and photosyntethic complexes share a common membrane; unlike green plants which use chloroplasts for this purpose. However, cyanobacteria have similar photosystem structures as chloroplasts. And instead of using chlorophylls as antenna pigments, they use a different light-harvesting system structured by phycobiliproteins.
Cyanobacteria have evolved an extremely effective single-cell CO2 concentrating mechanism (CCM). And photosynthesis is from where the energy is taken to drive the CCM and carbon fixation reactions. The CCM is a multicomponent system comprised of the carboxysome and a suite of inorganic carbon transporters that collectively enable cyanobacteria to proliferate under relatively low ambient atmospheric CO2 (0.039%) and high O2 (21%) levels (2). CO2 can enter the cell by diffusing over the plasma layer which sometimes makes it hard to accumulate CO2 inside the cell. To keep the loss of CO2 from the cell and to keep up an internal gradient of CO2 over the cell envelope, cyanobacteria contain two NADPH- dependent systems that catalyze the hydration of CO2 to HCO3–, NDH-I4 and NDH-I3 (3).
There are three active transport systems and CO2 hydration components of the CCM that effectively generate a high concentration (up to 40mM) of HCO3– within the cell. HCO3– diffuses into the carboxysome, where it is converted to CO2 by carbonic anhydrase (CA) and then combined with ribulose 1,5-bisphosphate (RuBP) by RuBisCO to produce two molecules of 3-PGA, which after exits the carboxysome and are utilized by the enzymes of the CBB cycle (Fig. 2). Carboxysomes are bacterial micro-compartments composed entirely of protein. They work in the cyanobacterial CO2 concentrating mechanism by encapsulating the ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCO), and carbonic anhydrase (CA) within a protein shell (Fig. 2) (4) .
Fig. 2 Shematic diagram showing the interrelationship of the CCM, Calvin-Benson-Bassham cycle, and the light reaction of photosynthesis. Obtained from “The carboxysome: function, structure and celular dynamics”, by Cameorn et al 2014, The Cell Biology of Cyanobacteria, page 172. Copyright [2014] Caister Academic Press.
By the other hand, photosynthetic organisms not only need atmospheric CO2 to store harvested energy from sunlight in the form of energy rich carbohydrates. But also, in cyanobacteria, algae and plants, CO2 is needed for the regulation of photosynthetic electron transport in Photosystem II (PSII), the enzyme responsible for light-induced primary charge separation and subsequent water oxidation (1). Nevertheless, there is a lack of knowledge how these interactions happen between CO2 and proteins conforming the Photosystem II.
This knowledge of the CCM may clarify the mechanisms by which cyanobacteria handle with limiting CO2/HCO3- but not necessarily how CO2/HCO3- availability is coupled to metabolism, replication, and growth. Interactions between CO2 and neutral amine groups on proteins (to form carbamates) have been proposed as a mechanism for biological regulation. CO2 converts these neutral amines into negatively charged groups (5). The introduction of a negative charge by CO2 binding in this manner (carbamylation) can alter protein function and therefore explain how cyanobacteria detect environmental CO2.
Unfortunately, researchers have lacked experimental tools to broadly assess carbamylation as an underpinning mechanism for organismal responses to CO2 and thus its broad biological relevance is controversial.
Prof. Martin Cann’s group has now developed a new chemical technology to identify protein carbamylation. We now use this technology to investigate the role of carbamylation in the metabolic response of cyanobacteria to environmental CO2.
Previous work in the group has shown potential trapping sites in cyanobacteria proteome. 14CO2 has been trapped on the proteome of Synechocystis and quantified the 14CO2 trapped by scintillation counting. The Synechocystis proteome contained ample trapped 14CO2 even when allowing for 50% of the total protein sample as Rubisco. Synechocystis therefore has carbamylated proteins. We identified a variety of carbamylated proteins supporting the hypothesis that carbamylation is a common and relevant post-translational modification in cyanobacteria that might mediate cellular CO2 responses. We have selected this first year of study one of these targets for initial study in addition to a further analysis of the extent of protein carbamylation in the cyanobacterial proteome described below.
Slr2067. A Photosystem II protein.
The allophycocyanin alpha subunit is a light harvesting phycobiliprotein component mediating energy transfer to phycocyanin and chlorophyll within the cyanobacteria Photosystem II (6).
Cyanobacteria, like algae and plants, perform oxygenic photosynthesis. A process where the energy of light is used to synthesize organic compounds from carbon dioxide and water creating oxygen as a waste product. But unlike plants, cyanobacteria photosynthesis and respiration require electron transport pathways that are catalysed by protein complexes in membranes. In most cases, the photosynthetic machinery is integrated within folds, called thylakoids, of the external cell membrane. The thylakoid membrane is an internal membrane system that separates the cytoplasm from the lumen and that is present in almost all cyanobacteria. It contains both photosynthetic and respiratory electron transport chains. These electron transport chains intersect, and use part of the same components in the membrane. The photosynthetic machinery is constituted by a series of protein complexes in membranes and a series of electron carrier molecules of lipidic nature or protein nature that catalyze the light reactions. There are four major protein complexes in the thylakoid membrane: Photosystem II (PSII), Cytochrome b6f complex, Photosystem I (PSI), and ATP synthase. These four complexes work together to ultimately create the products ATP and NADPH (7, 8, 9, 10).
Phycobilisomes
Cyanobacteria possess a water-soluble antenna located on the top of the thylakoids membrane, the phycobilisomes. They are mainly composed of brilliantly coloured polypeptides, named phycobiliproteins; who exhibit molecular masses in the range 15-22 kDa. The presence of linear tetrapyrroles (phycobilins) cause their strong absorbance in the visible range (11). The pigments are organised in a hierarchic structure where the chromophores absorbing at higher energy (phycocyanin and phycoeritrin) are located at the periphery of the complex, and those absorbing at lower energy (allophycocyanin) build up the phycobilisomes core. Besides these pigment-binding subunits, the phycobilisomes contains a number of proteins called linker proteins, which have a principal structural function. In general, phycobilisomes are considered as an antenna serving principally Photosystem II. Nevertheless, there may be an association of this antenna to the reaction centre of PS I, specifically under conditions that promotes a high level of reduction of the inter-chain electron mediator plastoquinone, or strongly unbalanced excitation between the two photochemical centres (12).
As mentioned before, phycobiliproteins assemble to form an ultra-molecular complex known as phycobilisomes (PBS). Most of the PBSs from cyanobacteria show hemidiscoidal morphology in electron micrographs. The hemidiscoidal PBSs have two discrete substructural domains: the peripheral rods which are stacks of disk-shaped biliproteins, and the core which is seen in front view as either two or three circular objects which arrange side-by-side or stack to form a triangle. For typical hemidiscoidal PBSs, the rod domain is constructed by six or eight cylindrical rods that radiate outwards from the core domain. The rods are made up of disc-shaped phycobiliproteins, phycoerythrin (PE), phycoerythrocyanin (PEC) and phycocyanin (PC), and corresponding rod linker polypeptides (Fig.3). The core domain is more commonly composed of three cylindrical sub-assemblies. Each core cylinder is made up of four disc-shaped phycobiliprotein trimers, allophycocyanin (AP), allophycocyanin B (AP-B) and AP core- membrane linker complex (AP-LCM). It is by the core-membrane linkers that PBSs attach on the stromal side surface of thylakoids and coupled with PSII (12).
Fig 3 Schematic organization of a hemidiscoidal phycobilisomes exhibiting a tricylindrical core domain arrangement. (A) Repartition of the three main phycobiliprotein groups in the phycobilisomes. (B) Distribution of their linker polypeptides in the two domains of the phycobilisomes. Adapted from “Isolation, characterization and electron microscopy analysis of a hemidiscoidal phycobilisomes type from the cyanobacterium Anabaena sp. PCC 7120,” by Ducret et al 1996, Eur. J. Biochem, Volume 263, page 1011.
Electron transport flow
The electron transport chain starts with the absorption of energy from light through the phycobilisomes. This complex is integrated of a number of antenna molecules: phycoerythrin, that is absent in most cyanobacteria, phycocyanin and allophycocyanin. These pigments have different spectral properties, each has a specific absorption and fluorescence emission maximum in the visible range of light (Fig. 4). Thus, their presence and this particular arrangement within the phycobilisomes allows absorption and unidirectional transfer of light to chlorophyll a (Chl a) of the photosystem II. Thereby, in comparison of algae and plants, the cells can take advantage on the available wavelengths of light (in the 500-650 nm range), that is not accessible to chlorophyll.
The general path of excitation energy transfer in most cyanobacteria is:
Phycocyanin →Allophycocyanin →Chlorophyll a
After the photons passes through the antenna molecules, they reach the reaction center Chl a (P680) and then the excited P680 donates its electron to an electron aceptor (A, Figure 4B). The electron missing of the Chl a is donated from an electron donor (D, Figure 4B) (13).
Fig 4 Structural organization of the antenna system of PSII for red algae and cyanobacteria (A) and energy transfer steps including charge separation (photochemical reaction) at the PS II reaction center (B) for cyanobacteria. Adapted from “Adventures with cyanobacteria: a personal perspective,” by Govindjeeand Shevela 2011, Frontiers in plan science, Volume 2, page 3. Copyright [2011] Govindjee and Sheela.
Besides the electron transfer complexes that are specifically involved in photosynthetic electron transfer, Photosystem II (PS II) and Photosystem I (PS I), there are specific specific complexes for respiratory electron flow include type 1 NADPH dehydrogenase (NDH-1), succinate dehydrogenase (SDH), and the terminal oxidase. Plastoquinone (PQ), cytochrome b6f complex (cyt b6f) and plastocyanin (PC) are shared by both pathways (Figure 5).
The electron transfer as we can see in Figure 5, Photosystem II (PS II) uses light energy to split water and to reduce the PQ pool. Electrons are transported from the PQ pool to the cytochrome b6f complex and from there to a soluble electron carrier on the luminal side of the thylakoid membrane. In cyanobacteria, this soluble carrier may be plastocyanin or cytochromec553, depending on the species and on the availability of copper (plastocyanin is a copper-containing enzyme). Either of these soluble one-electron carriers can reduce the oxidized PS I reaction centre chlorophyll, P700. This oxidized form of the reaction centre chlorophyll is formed by a light-induced transfer of an electron from PS I to ferredoxin (Fd) and eventually to NADP. The proton gradient across the thylakoid membrane is used for ATP synthesis by the ATP synthase in the thylakoid; this ATP may be applied for CO2 fixation and for other cell processes (14).
However, in the oxygenic photosynthesizers (cyanobacteria, algae and higher plants), CO2 is not only required as the terminal electron acceptor to synthesize carbohydrates, but also for the regulation of photosynthetic electron transport in Photosystem II (PSII), the enzyme responsible for light-induced primary charge separation and subsequent water oxidation. Although, further experiments are required to deep inside these theme (15).
Cyanobacteria and biofuel production
Energy and environmental concerns have been rising nowadays and the develop of sustainable and renewable energies are needed. As photosynthetic prokaryotes cyanobacteria grow through direct use of sunlight and CO2, which they convert through they metabolism into different products of interest like biofuels. Genetic manipulation and metabolic engineering has been an attractive direction of sustainability. Many studies have been developed to optimize photoautotrophic production of biofuels using cyanobacteria (17). Besides these improvements, cyanobacteria have many interactions with CO2 sources where not all might be beneficial to biofuel production. It is remarkable that we know so little of how cyanobacteria directly detect environmental CO2 and the resulting effects on metabolism. Such knowledge would enhance opportunities for engineering carbon fixation in green biotechnology. Engineering efficient biofuel production requires an understanding of the molecular links between CO2 and metabolism. This project will identify these links.
Hypotheses to be tested.
Direct interactions and modifications between CO2 and cyanobacterial phycobiliproteins explain how cyanobacterial metabolism respond to changes in environmental CO2 levels. Specifically, I hypothesize that CO2 modification of phycocyanins and allophycocyanins enhances energy transfer efficiency enabling the harvesting of light energy to be coupled to carbon availability.
Objectives
Our objective is to investigate carbamylation in the response of cyanobacteria to environmental CO2 levels. Carbamylation is not the only mechanism through which CO2 can interact with proteins nor may it be the only mechanism by which cyanobacteria detect inorganic carbon. However, it is a proven crucial post-translational modification in photosynthetic organisms yet remains wholly underexplored. Our technology therefore gives an original approach to the important question of how cyanobacteria couple carbon availability to normal physiological processes.
I will use recombinant proteins in in vitro assays to prove a specific response to CO2 and examine how CO2 modulates the parameters of the phycobiliproteins. This will be combined with genetic analyses to examine the role of the identified CO2 binding site in the physiology of the intact organism.
MATERIALS AND METHODS
Biological Material
Isolation of phycobilisomes from Synechocystis sp. PCC 6803
For the isolation of phycobilisomes from cyanobacteria methodology from Ducret et al 1966 (18) was modified. The cyanobacterium Synechocystis sp. PCC 6803 was cultivated in a BG-11 liquid medium with a pH of 8.0 at 28ºC. The light was provided by 18W Philips fluorescence lamps arranged in rows and the flasks were placed in between. Cells from six litres of culture were harvested at an OD730nm of 0.2. The cells were pelleted by centrifugation at 4 000 g for 30 minutes at 10ºC. The pellet was resuspended in 100 mL of isolation buffer (0.9 M KH2PO4 pH 7.0, 2mM EDTA, 1mM NaN3). The cell walls were disrupted with a cooled French press at a pressure of 40 PSI four times. The membrane fraction was pelleted at 20 000 rpm for 30 min at 4ºC (Rotor JA 25.50). After discarding the supernatant, the pellet was resuspended in 10 mL of isolation buffer, 1 mM of Pefabloc and 2% v/v of Triton X, and the solution was stirred for 1 hour. The membranes were centrifuged out at 20 000 rpm for 25 minutes (Rotor JA25.50), and a green supernatant was carefully removed. To separate from chlorophyll complexes, phycobilisomes were ultracentrifuged at 42 000 rpm for 1 hour 45 minutes. (Rotor SW60Ti). The supernatant was removed and a deep blue pellet was dissolved and stirred overnight with 5 mL of isolation buffer and 1 mM of Pefabloc.
To remove aggregates from chlorophyll, the solution was centrifuged at 18 000 rpm (Rotor SW60Ti) for 30 at 10ºC.
Finally, a blue supernatant was transferred to a discontinuous sucrose gradient from 2 M to 0.5 M sucrose to isolate intact phycobilisomes. The step gradient was centrifuged at 21 000 rpm for 18 hours at 20ºC and phycobilisomes were recovered from the 1.2 M sucrose zone, dialyzed against PBS (phosphate buffer saline) 50 mM and stored at -80ºC until its use.
Recombinant proteins. Protein expression and purification.
Plasmids used to produce proteins apcAB and cpcAB are described in Table 1. Cyanobacterial apcAB and cpcAB expression vectors were provided by Avjit Biswas from The Pennsylvania State University. Protein expression and purification was performed similar to Biswas et al 2010 method.
For apcA expression, the apcA gene was amplified by PCR from Synechocystis sp. PCC 6803 chromosomal DNA using forward primer (5’-AAGGATCCGATGAGTATCGTCACGAA-3’) and reverse primer (5’-GCGAGCTCCTAGCTCATTTTTCCGAT-3’) and cloned into pCDF Duet plasmid using restriction enzymes BamHI and SacI.
For allophycocyanin trimer expression, plasmids ppcyA, pcpcUS and papcAB were co-transformed by heat-shock into BL21 competent cells.
By the other hand, plasmid ppcyA was transformed into BL21 E3 and made them quimically competent following Inoue methodology. After that, for phycocyanin trimer expression plasmids cpcAB and pcpcUS were co-transformed into ppcyA-BL21 E3 competent cells as well as plasmids papcA and pcpcUS into another cell.
Table 1. Plasmids used to produce recombinant phycobiliproteins
Phycobiliprotein
Plasmids co-transformed
Antibiotic
Reference
αAP, βAP
ppcyA
Cm
(16)
pApcAB
Ap
pcpcUS
Km
αPC, βPC
pcpcBA
Sp
pcpcUS
Km
pPcyA
Cm
αAP
pApcA
Sp
This study
pPcyA
Cm
(16)
pCpcUS
Km
Colonies from LB medium plates with appropriate combinations of antibiotics were selected (Table 1) and tested for expression of each protein.
To produce the phycobiliproteins, a 300 mL overnight starter culture was added to 6 litres of LB medium (50 mL per litre) with the appropriate combination of antibiotics. The culture was shaken at 37ºC for 4 hours until the optical density at 600 nm was 0.6. Production of T7 RNA polymerase was induced with a final concentration of 0.1 mM IPTG at the culture and cooled down to 28ºC to reduce inclusion bodies. The cultures were further incubated for 16h in the dark. The next day, the cells were harvested by centrifugation and washed with a 20 mM sodium phosphate and 20mM imidazole pH 7.4 buffer. Pellets were sonicated six times during 10 seconds each time with 10 seconds on ice intervals. After centrifugation, the lysates were loaded onto a pre-equilibrated 5 mL Histrap Ni2 chelating affinity columns. The column was then washed with 20 mM sodium phosphate and 100mM imidazole pH 7.4, to remove weakly bound host proteins. Finally, the recombinant proteins were then eluted with 20 mM sodium phosphate and 500 mM imidazole, pH 7.4 and then dialyzed overnight with 50 mM PBS pH 8.0.
Carbamates discovery and Ci experiments
Synechocystis sp PCC 6803 intact phycobilisomes were trapped by Linthwaite’s novel trapping CO2 technology to confirm carbamylation sites within the energy antenna (19). Isolated phycobilisomes were dialyzed into phosphate buffer 50 mM pH 7.5 and concentrated to 1 mg/ml. Phycobilisomes were added a HCO3- and TEO system maintaining a stable pH of 7.5 0.5. After the carbamate trapping, the sample was trypsin digested and the samples were sent to analysis by mass spectrometry.
Spectral analysis
Absorption spectra and fluorescence spectra were obtained by a Synergy H4 Plate Reader. The absorption spectra were recorded from 500 nm to 700 nm at room temperature, with a bandwidth of 5 nm, and a scan speed of 240 nm/min.
The fluorescence emission spectra were recorded with excitation at 590 nm. The emission slit widths were set at X nm, and the scan speed was XXX nm/min. Samples concentrations were adjusted to an absorbance between 0.05 – 0.1 at the absorption maximum of each protein.
Spectral response was analysed with and without the presence of inorganic carbon adding sodium bicarbonate in different concentrations to the system. Sodium chloride was added to the system as a control to preserve the same saline concentration in the system.
Photoluminiscence quantum yields (PLQY)
The photoluminiscence quantum yields () were recorded with a Fluorolog-3 spectrofluorometer (HORIBA Scientific) and a Quanta- F-3029 Integrated Sphere at room temperature in 1cmx1cm cuvettes.
The sample is placed in the integrating sphere, and excited with a monochromatic source of wavelength (lambda). The quantum yield, , is, by definition, photons emitted to photons absorbed:
where is the integrated luminescence of the film caused by direct excitation, and is the integrated luminescence form an empty integrating sphere (only a blank). The term is the integrated escitation profile when the sample is directly excited by the incident beam and is the integrated excitation profile from an empty integrating sphere (without the sample, only a blank) (20).
Measurements were determined in PBS (50 mM, pH 8.0) and sample concentrations were adjusted to an absorbance of 0.1 at the maximum fluorescence.
PLQY ( resulted in an average of four scans. The spectral ranges used in this study for the recording of the fluorescence spectra are listed in table 2.
Table 2. Spectral ranges for measurements of photoluminescence quantum yields.
Protein
Excitation (nm)
Fluorescence range(nm)
Phycobilisomes
570
610-720
610
620-720
Allophycocyanin trimer
615
620-750
Phycocyanin trimer
Results and discussion
Phycobilisomes
Intact phycobilisomes of the cyanobacterium Synechocystis sp PCC6803 were obtained by a modified method of Ducret et al. (1996) based on methodology of Gantt et al. (1979). This protein complex was extracted from the 1.2 M zone of a sucrose gradient ultracentrifugation (Fig. 6). The isolation procedure of in this study yielded highly purified phycobilisomes showing an apparently intact energy-transfer pathway as considered from the absorption and fluorescence spectra (Fig. 7). The absorption maximum at 620 nm shows the prevalence of phycocyanins within the phycobilisomes. The fluorescence excitation spectrum showed that these compounds are energetically linked to the terminal fluorescence transmitters. Isolated phycobilisomes showed an emission maximum at 655 nm and excitation maximum at 625 nm, suggesting the energetically linked prevalence of phycocyanins and allophycocyanins (11).
Fig. 6 Phycobilisomes isolated form sucrose gradient
Fig. 7. Absorption and fluorescence spectra of intact phycobilisomes from Synechocystis sp. PCC 6300 in 50 mM Phosphate Buffer pH 8.0. Left (—) absorption spectrum; right, (—) emission fluorescence spectrum, (—) excitation fluorescence spectrum.
Whole phycobilisomes with an OD620nm= 1.5 were analysed by SDS/PAGE with a glycine buffer system achieving optimal separation of phycobiliproteins. The protein bands at 14 kDa and 21 kDa shows the amount of phycoerythrocyanin and rod-core complexes present in the pycobilisome (Fig. 8) (11).
Fig. 8 Phycobiliprotein analysis by SDS/PAGE. L: ladder; PBS: phycobilisomes isolated from sucrose gradient
Phycobilisome Ci Experiments
Trapping carbon dioxide
CO2 and phycobilisomes were trapped using TEO reagent, which transfers an ethyl group to the amino group of the proteins when the carbamate is formed. Intact phycobilisomes were equilibrated with CO2/HCO3- and TEO at pH 7.4. This reaction creates a modification robust enough for downstream analysis by MS. The reaction was dialysed and digested with trypsin. Peptides were fractionated and samples run on a Q-star MS and analysed using X!Tandem. The MS chromatograms were analysed looking for ethyl groups transferred to the carboxyl group of amino acids where the carbamate was expected. We found one trapping site at the K6 of allophycocyanin alpha chain (Fig 9).
Fig. 9 X!Tandem analysis from a CO2 trapping method
Fluorescence intensity
Besides identifying CO2 binging sites, native phycobilisomes experiments were carried with inorganic carbon to see if there was any influence of this PTM at the native antenna structure. Fluorescence emission under different conditions of inorganic carbon were analysed. The emission and excitation spectra was recorded with an excitation steam at 590 nm are shown at Fig. 10. A small increment of the emission and excitation intensity can be observed between phycobilisomes exposed to HCO3- and controls with just NaCl, which suggested to be caused by the increment on the saline concentration.
Fig. 10 Effect of Ci on the fluorescence intensity of Synechocystis sp PCC 6803 phycobilisomes.
PLQY
For the study of energy transfer at the phycobilisomes two excitation wavelengths (Table 3) were used aiming to detect the effect of Ci at the different stages of energy transfer between the phycobiliproteins organized within the phycobilisomes. Integration of the instrument-corrected incident beam’s signal intensities I was performed in first instance from 560 to 580 nm for 0.3 s and then from 600 to 620 nm for 0.3 s. Integration of the emission-signal region of the blank and sample was from 620 to 720 nm for both excitation wavelengths measurements. As
For allophycocyanin and phycocyanin trimers PLQY measurements, integration of the instrument-corrected excitation signal was performed from 505 nm to 525 nm and the integration of the emission signal region from 610 to 720 nm. Normalized PLQY results from these integrations are shown in the Fig. 11. An unpaired t test with Welelch’s correction analysis with a P value <0.05 was performed with Graphpad Prism Software, we could observe a significate difference between the two treated groups: phycobilisomes+Ci excited at 570 nm and phycobilisomes+Ci excited at 610 nm. The PLQY results were normalized to 1.0 to analyse one sample t test with a P value <0.05 and evaluate the difference with and without Ci. Absolute Quantum Yields with a 570 nm excitation wavelength showed a significate difference between the control and samples with Ci. The increment of PLQY at a 610 nm excitation wavelength between the control and the samples suggests a possible influence of CO2 at the energy transfer at phycocyanins and allophycocyanins (pigments of photosystem II which absorbs energy of light in the 630-650 nm range), although the statistical analysis did not show a significant difference(Fig. 11).
Fig. 11. Absolute Quantum Yields of Phycobilisomes. Control samples normalized to 1.0 to show increment of samples with Ci at two different excitation wavelengths.
Recombinant proteins
The recombinant allophycocyanins and phycocyanins were successfully expressed in E. coli cells. The crude and purified protein samples were analysed by SDS-PAGE (Fig. 12). Induced samples and Histag purified proteins showed bands of approximately 14 and 20 kDa, which correspond to the alfa and beta unit of allophycocyanin trimer respectively (Fig. 12. A). And similar size bands corresponding to the alfa and beta unit of the recombinant phycocyanin (Fig.12. B). Protein expression of recombinant alpha allophycocyanin could be observed with a band of 20 kDa (Fig. 12.C).
High yields of allophycocyanins where effectively purified conformed as trimers. To confirm this, a gel filtration assay was performed. After 65 mL of elution only one peak between 66 kDa and 29 kDa size controls was observed (Fig. 13), suggesting the assembled allophycocyanin trimer of 57.4 kDa . Phycobiliproteins are self- assembled, conformed by two and subunits to form the monomers that are further assembled into ()3 trimers. The fractions of the gel filtration were analysed by SDS-PAGE to confirm the presence of both subunits. Two bands could be seen at fraction five and 6 after 65 mL of elution (Fig. 14). The first band 21 kDa corresponds to the subunit and the second band of 14 kDa to the subunit similar to other native and recombinant allophycocyanins reported (21, 22).
Fig. 13 Allophycocyanins gel filtration analysis. Green: allophycocyanin; Orange: BSA cytochromeC of 66 kDa; Blue: aprotinin carbonic anhydrase of 29 kDa
Fig. 14 SDS-PAGE allophycocyanin analysis. L: Ladder; FC5: gel filtration fraction five after 65 mL of elution; FC6: gel filtration fraction six after 66 mL of elution.
The recombinant APC and PC were further analysed for fluorescence spectrum with the presence and absence of inorganic carbon (Fig. 15). Both trimers showed similar spectral response than the phycobilisomes spectrum, increasing the emission and excitation because of a higher saline exposure.
Fig. 15. Effect of Ci on the fluorescence intensity of two recombinant proteins.
By the other hand, a one sample t test un a P value <0.05 of PLQY’s of the recombinant proteins showed a significate difference between allophycocyanin with and without 1 mM Ci (Fig. 16).
Fig.16. Effect of Ci on the energy transfer efficiency of recombinant allophycocyanins and phycocyanins.
CONCLUSIONS AND FURTHER WORK
Successfully isolation of intact native phycobilisomes from cyanobacterium Synechocystis sp. PCC 6803 were used to identify protein carbamates within the pigments conforming it. Carbamylation of Sir2067 and CO2 binding sites were identified. Photoluminescence quantum yields of phycobilisomes showed to increase with the presence of CO2 suggesting possible effect of the carbamylated protein which absorbs energy in a range of 600 to 650 nm.
Recombinant proteins of phycobilisomes component were expressed in E. coli to observe the activity alteration of phycobiliproteins with CO2/HCO3-. Results of this study suggests an increase of the energy transfer efficiency of allophycocyanins in the presence of Ci.
As the identification of lysine 6 CO2 binding site of Sir2067 has been trapped, experiments will be performed in the presence and absence of CO2/HCO3- to examine for a carbon specific influence on the K6 residue of Slr2067.I will determine the energy transfer sensitivity with and without presence of CO2/HCO3-. As well I will determine the sensitivity to total inorganic carbon in appropriate gas sealed assays under a range of conditions (e.g. varying pH, substrate availability, temperature). I will confirm that the proteins are activated by CO2 through low-temperature short-time assays under conditions of inorganic carbon disequilibrium. We will use our trapping reagent and proteins mutant at the CO2 binding site to confirm the mechanism of CO2 binding.
I will investigate model cyanobacteria expressing either wild type CO2 binding phycobiliproteins in which the CO2 binding site has been removed by standard genetic methods. I will examine the response of wild type and mutant strain metabolism to environmental CO2. This will link CO2 binding at a particular site on a particular protein to its metabolic response to CO2 feedstock.
I will look for new CO2 binding sites in Synechocystis metabolism using proteomics technology.