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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.
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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.
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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.
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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
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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,
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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
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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
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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
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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.
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