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3 Biological Background Information 3 Biological Background Information 3.1 Structure of Plant Cells Every higher evolved organism consists of at least more than one cell, necessitating communication and signal transduction between individual cells. A schematic diagram of a plant cell and its different compartments is shown in Figure 3.1 [39]. Every cell is surrounded by a cell membrane (plasma membrane or plasmalemma), which creates and maintains a different electrochemical environment in the inner cell with respect to its outside. Other types of membranes define the boundaries of cell compartments (e.g. the vacuolar or the nuclear membrane) or even form internal compartments (e.g. the endoplasmic reticulum ER). Figure 3.1: Schematic diagram of a plant cell and its different compartments. The plasma–membrane is located close to the cell wall, the cell compartment exhibiting by far the strongest autofluorescence apart from plant chloroplasts [39]. Cell membranes consist of a bilayer of polar lipid molecules and embedded or attached proteins. All proteins investigated in this work are membrane–bound receptor proteins. The role of membrane–located proteins is to overcome the bordering membranes and to render possible an informational and material exchange between individual cells and between the cells and their environment. –23– 3 Biological Background Information All cellular compartments emit a certain amount of autofluorescence when excited with light of the appropriate wavelength, its intensity depending on the respective compartment, on the age of the cell and on several more parameters. In plant cells, the cellular compartment showing by far the strongest autofluorescence – apart from the chloroplasts – is the cell wall. The cell wall is located outside the cell membrane and provides stability, structural support and protection. Cell walls are found in plants, bacteria, fungi and algae but not in animals. The major carbohydrates making up the primary (growing) plant cell wall are cellulose (a polymer of glucose), hemicellulose (a mixture of polysaccharides) and pectin (a heteropolysaccharide). Furthermore, the plant cell wall also contains secreted proteins which, for instance, can regulate its texture and extensibility. Given that the plasma membrane of a plant cell is located close to the autofluorescent cell wall, plasmalemma–located proteins are one of the most challenging protein species for the in planta investigation by fluorescence microscopy. 3.2 Autofluorescent Proteins The first discovered autofluorescent protein, the Green Fluorescent Protein (GFP) from the jellyfish Aequorea victoria is still the most commonly used autofluorescent protein in fluorescence labelling and fluorescence microscopy of living cells. Just recently a whole series of autofluorescent proteins has been developed, covering the whole visible range (Clontech, 2007). This opens up new frontiers in fluorescence microscopy, but does not solve the autofluorescence interference problem, as autofluorescence covers the whole spectral range. - 24 - 3 Biological Background Information Figure 3.2: The Green Fluorescent Protein GFP. (A) Sterical structure of GFP from the jellyfish Aequorea victoria [40]. (B) Fluorescence emission spectrum of purified GFP in aqueous solution [23]. 3.3 Transformation of Plant Cells The basic concept of cell transformation is the transfer of external DNA in living cells. Thus, it is possible to obtain cells expressing a specific protein fused to an autofluorescent protein. Observing the autofluorescent label protein in the living cell via confocal fluorescence microscopy provides detailed information about the localisation and the dynamics of the protein in question. More complex information can be gained by labelling different proteins with diverse emitting autofluorescent proteins to study their localisation and interaction via Foerster Resonance Energy Transfer (FRET). A possible energy transfer from a donor to an acceptor chromophore via FRET can be detected in different ways. One established method is the recording of the donor’s fluorescence lifetime (for a detailed description of the technique see Chapter 2.4: Time Correlated Single Photon Counting) which decreases in the presence of an acceptor molecule. In the last decades many approaches have been established for plant cell transformation. As all plant cells investigated for this thesis have been transformed either by transient or by stable transformation via Agrobacterium tumefaciens, both procedures will be illustrated in the following. –25– 3 Biological Background Information 3.3.1 Stable Transformation via Agrobacterium tumefaciens This technique is frequently used for dicotyledonous plants like the cabbage plant Arabidopsis thaliana (Figure 3.3). The inserted DNA is integrated permanently in the plant’s genome. Stable transformation enables the propagation of the transformed gene to the next generation. All stable transformations in this thesis have been accomplished with Arabidopsis as it is a model organism in plant biology due to its short regeneration time and its small–sized, fully sequenced genome consisting of five chromosomes. Figure 3.3: The cabbage plant Arabidopsis thaliana. Agrobacterium tumefaciens is a Gram negative soil bacterium possessing a native gene transfer system which allows for the exploitation of plant cell metabolism. The strain contains tumor–inducing (Ti) plasmids of about 150 kbp. A schematic diagram of a Ti–plasmid is shown in Figure 3.4. One part of the plasmid, the vir genes, encodes for proteins required for the complete infection and gene transfer process for the transformation of the host plant and catabolic enzymes. Another part of the Ti–plasmid, the transfer–DNA (T–DNA), is transferred to the plant cell and integrated via the left (LB) and right border (RB) into the plant genome. In wild type Agrobacterium strains, the genes on the T–DNA code for - 26 - 3 Biological Background Information proteins which, one the one hand, cause the formation of a fast–growing, tumor–like structure with high biochemical activity at the plant and, on the other hand, the production of specific metabolites in the tumor–like structure, which can only be used by the bacterium. Figure 3.4: Schematic diagram of the Ti–plasmid. The vir genes encode for proteins required for the complete infection and gene transfer process. The T–DNA is inserted into the host cell. The left and right border sequence (LB/RB) serve for the identification of the T–DNA. Agrobacterium and its gene transfer mechanism are widely used in plant biotechnology. Therefore, the wild type genes of the T– DNA are replaced by the gene(s) of interest (e.g. a gene encoding for a GFP–fusion protein) by molecular cloning techniques [41]. For the generation of transgenic plants, the use of so–called binary vector systems have been established in the past decades [42]. A binary system consists of an easy to handle cloning vector, which is able to replicate in E. coli as well as in Agrobacterium tumefaciens, and a helper vector which carries the vir genes. Additionally, the vectors contain antibiotic resistance and/or herbicide resistance genes used to select for the presence of the binary vector in the bacteria and later for the selection of transgenic plants. The genes to be integrated in the plant genome have to be under the control of a promoter – a regulatory sequence in front of the gene(s) which control its activity. For stable transformation the infectious Agrobacterium strain carrying the vectors of interest and the plant cells have to be brought together. This can be achieved by, for instance, –27– 3 Biological Background Information spraying the flowers of the plant with the bacterial suspension, co–cultivation of the bacteria and plant organs such as roots or vacuum infiltration of the bacteria into the plant leaf. Afterwards the transformed cells or seeds have to be selected by applying antibiotics to the growth medium or spraying herbicides and the transgenic plant is regenerated from a single transformed cell. The complete transformation and regeneration process is illustrated in Figure 3.5. Figure 3.5: Stable transformation of plant cells via Agrobac–terium tumefaciens [39]. The gene transfer process can not only be used to insert new genes into cells but also to knock–out or knock–down the activity of a specific gene in the plant host cell. This can be used to gain additional information about the role of the protein encoded by the inactivated gene. 3.3.2 Transient Transformation via Agrobacterium tumefaciens Many scientific problems do not require the time–consuming creation of stable transgenic cell lines. In these cases transient transformation enables a more rapid analysis. In this work, transient transformation was achieved by infiltration of leaf tissue of tobacco plants (Nicotiana benthamiana) with - 28 - 3 Biological Background Information Agrobacterium suspension of the same strain used for the stable transformation. Hereby the Agrobacterium suspension is inserted into the intercellular region of the leaves by means of syringes via open stomatal cells [43]. In the following, the Agrobacteria transform individual cells which can then be investigated via confocal fluorescence microscopy after only few days. An antibiotics or herbicide selection or regeneration of a plant is not required during the transient transformation procedure. 3.4 Plant Chloroplasts 3.4.1 Structure of Plant Chloroplasts Chloroplasts are the location of photosynthesis in plant cells. The most important fluorescent components involved in the photosynthesis process of plants are the photosystems I and II (PSI and PSII) and the light–harvesting complexes LHCI–730, associated with PSI, and LHCII, associated with the PSII. These complexes are located in an internal chloroplast membrane, the thylakoid membrane. Figure 3.6: Diagram of a plant’s chloroplast. The LHCs are located in the tylakoid membrane (green). The thylakoid membrane can either be arranged stacked (granal) or unstacked (agranal) [39]. A diagram of a plant chloroplast is shown in Figure 3.6. The thylakoid membrane can either be arranged stacked (grana) or unstacked (agranal). LHCII and PSII are located mainly in grana stacks, LHCI–730 and PSI are located mainly in agranal thylakoid membranes. The LHCII complex consists of three –29– 3 Biological Background Information equal monomers sized around 3 nm x 5 nm. Each monomer consists of seven molecules chlorophyll a, five molecules chlorophyll b, two molecules Lutein and more than 200 amino acids [44]. 3.4.2 The Function of Light–Harvesting Complexes and Photosystems in Chloroplasts In the LHCs the chlorophyll and carotenoid molecules are specifically arranged, forming a large absorption area as well as an energetic funnel. The delocalised electron system of the chlorophyll molecules absorb light energy, thus being transferred from the ground state to the excited state. Afterwards, the absorbed light energy is transported via non– radiative energy transfer between the pigments to the reaction center of the corresponding photosystem PS. The centers of PSI and PSII consist of two chlorophyll a molecules and are the location of the light–dependent photochemical reaction, the first step in the photosynthesis. In these reactions light energy is converted into chemical energy, in the form of the energy– and redox carriers ATP and NADPH. The subsequent light–independent reactions perform the assimilation of carbon dioxide, meaning that carbon dioxide is converted to carbohydrates like glucose using ATP and NADPH. The chemical reactions occurring during the photosynthesis are very complex and would exceed the scope of this thesis, as their knowledge is not essential for the comprehension of the performed measurements and the obtained results. The complete photosynthesis process can be referred to [39]. Absorption spectra of isolated chlorophyll molecules differ from absorption spectra of chlorophyll molecules bound in light– harvesting complexes due to the high chlorophyll concentration enabling interactions between the chlorophyll molecules. Thus, the energy levels are lowered resulting in a red shift of the absorption maxima. Different kinds of chlorophyll molecules (e.g. chlorophyll a and b) exhibit different absorption maxima due to different functional groups. As LHCI and LHCII exhibit different chlorophyll ratios (Chl a/bLHCII = 1.4; Chl a/bLHCI = 2.5) [44], the absorption maxima and therewith the fluorescence emission of the LHCs - 30 - 3 Biological Background Information are shifted unequally. LHCII exhibits its fluorescence emission maximum at emLHCII = 680 nm whereas LHCI exhibits its fluorescence emission maximum at emLHCI = 730 nm [45]. Figure 3.7 shows fluorescence spectra of isolated LHCI (Figure 3.7.B) [46] and LHCII (Figure 3.7.A) [47] complexes recorded at low temperature (77 K). Figure 3.7: Fluorescence emission of LHC complexes at low temperature (77 K). (A) LHCII exhibits its fluorescence emission maximum at emLHCII = 680 nm (pea) [47]. (B) LHCI exhibits its fluorescence emission maximum at emLHCI = 730 nm (tomato) [46]. At room temperature, most of the fluorescence in both spectral regions is emanated from PSII due to energy transfer between the various photosynthetic complexes. Therefore fluorescence spectroscopy in plant chloroplasts has up to now mainly been performed at low temperature (77 K), accepting the disadvantage of investigating a non–living system. –31–