Novel Applications of Fluorescence Decay Analysis in Living Tissue at Single Cell Level

Neuartige Anwendungen der Fluoreszenz–Zerfalls–Analyse in lebendem Gewebe auf Einzel–Zell–Niveau

Kirstin Elgass

Kurzübersicht

Neuartige Anwendungen der Fluoreszenz–Zerfalls–Analyse in lebendem Gewebe auf Einzel–Zell–Niveau
ISBN: 978-3-941216-24-2
Veröffentlicht: Oktober 2010, 1. Auflage, Einband: Hardcover, Seiten 200, Format 170x240 mm, Gewicht 0.55 kg
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Novel Applications of Fluorescence Decay Analysis in Living Tissue at Single Cell Level

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Fluorescence microscopy is a powerful tool to study the subcellular localisation and intracellular dynamics of fluorophore tagged proteins in living cells. Optical and spectroscopic technologies working at subcellular resolution with quantitative output are required for a deeper understanding of molecular processes in living cells and for the realization of predictive biology at a cellular and subcellular level. However, although established in the physical sciences, these techniques are rarely applied to cell biology especially in the plant sciences. This is due to several problems inherent in live-cell imaging, which will be specified in the first part of this volume. In the following, several approaches to circumvent these problems will be presented. This includes a novel method to suppress autofluorescence interference in fluorescence microscopy images of living cells. Additionally, the power of combining these approaches is demonstrated in several impressive examples.

 

Dr. rer. nat. Kirstin Elgass is member of the group "Plant high resolution imaging/spectroscopy
Eberhard Karls Universität Tübingen: Institut für Physikalische und Theoretische Chemie
Contact:
Institute for Physical and Theoretical Chemistry 
Universität Tübingen
Auf der Morgenstelle 8
D-72076 Tübingen

.

Introduction

Scope of this Thesis
Fluorescence microscopy is a powerful tool to study the subcellular localisation, interaction and intracellular dynamics of fluorophore tagged proteins in living cells [1–3]. The use of green fluorescent protein (GFP) and its derivates to create fluorescent fusion proteins as well as of the labelling of compartments with organic dyes facilitates the in vivo analysis of cell biological processes [1–3].

In the past years, many efforts have been done to improve the spatial resolution of confocal fluorescence microscopes below diffraction limit allowing for the investigation of subcellular compartments and responses on a nanometer scale. This has become possible by combining fluorescence microscopy with new time–resolved laser spectroscopic methods like higher–order pulsed laser irradiation (Stimulated Emission Depletion microscopy, STED) [4, 5]. Different approaches use single molecule blinking statistics (Photo Activated Localisation Microscopy, PALM) [6] and high resolution colocalisation of single molecules (STORM) [7] to obtain ultra–high spatial resolution. For observing dynamic processes associated with single molecules, techniques such as fluorescence correlation spectroscopy (FCS) are available [8–13].

However, live–cell imaging is still hampered due to several inherent problems, which will be described shortly in the following and will get even more obvious in the main text of this thesis.

The major problem in live–cell imaging using fluorescent labels is the interference of unspecific autofluorescence exhibited to a certain degree by every living organism. Therefore, high resolution and sensitive fluorescence microscopy is difficult to carry out in all living systems ranging from single cell organisms like bacteria to cells of higher organisms (mammalian cells). Extreme conditions are given in living plants, due to the additional autofluorescence of cellular compartments typical for plants such as the cell wall. This problem is usually circumvented by studying proteins of higher plants in mammalian cells [14], plant protoplasts [15, 16] or plant cells after plasmolysis [17]. However, protoplast preparation and plasmolysis induce stress responses and may modify the subcellular partitioning, intracellular dynamics and activity of the receptors. Furthermore, expression of plant receptors in heterologous systems may lead to artificial results. Due to these limitations, the knowledge of cell biological mechanisms involving for instance, membrane–associated receptors, which act at the plant cell surface, traffic inside the plant cell and induction of early cellular responses, is restricted.

Additionally, even if the autofluorescence interference is circumvented in any way, fluorescence intensity measurements provide live–cell images with spatio–temporal resolution but usually no information about the physico–chemical parameters in the very close environment of the fluorescence tagged fusion protein. Thus, it is difficult to observe effects of internal or external effects on a GFP fusion protein, which do not result in a change of its localisation and stability but in a change of its nearby environment. Recent progress in fluorescence microscopy, using autofluorescent proteins which show a spectral shift upon changes in the physico–chemical environment such as pH or redox state [18–21], has opened new possibilities in live–cell imaging. Although these mutants have extended the field for intracellular environmental studies, their use has some intrinsic limitations. For instance, it is often not feasible to perform spectral mapping, a method which records fluorescence spectra spatially resolved for every image spot, due to extraordinary long measuring times. In addition, spectral filtering using band pass filters always suffer from spectral “bleed–through” (leakage of a particular signal into a different detection channel), which causes interference of the spectral channels, hampering a quantitative data analysis [22]. Background fluorescence with a broad spectral emission of differential wavelength–specific intensity such as that of the plant cell walls [23] also interferes with the quantitative analysis of environmentally induced spectral fluorescence shifts.

A complex system such as a living organism contains a manifold of components often coupled with each other e.g. in signal transduction or energy transfer pathways. Due to this interplay of several components, it often is highly complicated to investigate these components separately, particularly with regard to their exact amount and relative concentration. This can be done by extracting the components in question from their native environment but this obviously results in the destruction of the intact organism. For the investigation of variations in the constitution of a coupled system due to changes in external parameters, it is essential to keep the system alive and therefore, one has to develop methods to monitor the constitution of such a coupled system in vivo.

This thesis deals with several approaches to circumvent all specified problems inherent in fluorescence microscopy of living cells.

 

Outcome of this Thesis

Optical and spectroscopic technologies working at subcellular resolution with quantitative output are required for a deeper understanding of molecular processes and mechanisms in living cells. Such technologies are a prerequisite for the realization of predictive biology at a cellular and subcellular level. However, although established in the physical sciences, these techniques are rarely applied to cell biology in the plant sciences.

This thesis deals with several approaches to circumvent all specified problems inherent in fluorescence microscopy of living cells. A combined application of one–chromophore fluorescence lifetime imaging microscopy (ocFLIM) and wavelength–selective fluorescence microscopy is presented to analyse the subcellular localisation and intracellular dynamics of fluorophore tagged proteins in living plant cells resulting in the revelation of a ligand induced response pathway in living plants.

The experiments also show the potential of one–chromophore fluorescence lifetime imaging microscopy (ocFLIM) for the in vivo monitoring of the biochemical and biophysical subcellular environment using GFP fusion proteins as probes. The excited state lifetime of fluorescent proteins varies in response to alterations in the physico–chemical environment such as the pH–value, the refractive index and the electric field of the surrounding medium. Thus, alterations in the fluorescence lifetime of GFP–fusion proteins have been used to measure these physico–chemical parameters in living plant cells. One–chromophore fluorescence lifetime microscopy, combined with wavelength–specific fluorescence microscopy, opens up new frontiers for dynamic and quantitative in vivo analysis of cellular processes at high resolution which are not addressable by standard imaging technologies or transmission electron microscopy.

Furthermore, a novel method, fluorescence intensity decay shape analysis microscopy (FIDSAM), to suppress autofluorescence interference in common fluorescence intensity images is presented. The method benefits from the fact, that the fluorescence decay shape of a fluorescent label differs significantly from the autofluorescence decay shape. The applicability of this method is demonstrated impressively in several biological samples including some of the most challenging ones like AFP–fusion proteins in living plants expressed under their native promotor resulting in a very low label–to–autofluorescence ratio.

The photosystems of plants consists of a multitude of interacting components transferring energy from one component to the other. Thus, only the last component of the energy transfer chain can be observed by fluorescence microscopy at room temperature. By applying fluorescence microscopy at low temperature or by extracting the different components from their native environment, this energy transfer can be eliminated. However, due to this energy transfer, it is very difficult to investigate the components separately without destroying the intact, living system. This thesis presents two independent approaches for the investigation of the very complex photosystems and changes in their constitution in vivo.

 

Summary of the Thesis

In this thesis, novel approaches to circumvent serious problems inherent in live–cell imaging are presented. The major problem in live–cell imaging using fluorescent labels is the interference of unspecific autofluorescence exhibited to a certain degree by every living organism. Therefore, high resolution and sensitive fluorescence microscopy is difficult to carry out in all living systems ranging from single cell organisms like bacteria to cells of higher organisms (mammalian cells). Extreme conditions are given in living plants, due to the additional autofluorescence of cellular compartments typically for plants such as the cell wall. Therefore, the methods presented here have been applied to living plants representing the most challenging samples for fluorescence microscopy.

Additionally, even if the autofluorescence interference is circumvented in any way, fluorescence intensity measurements provide live–cell images with spatio–temporal resolution but usually no information about the physico–chemical parameters in the very close environment of the fluorescence tagged fusion protein. Thus, it is difficult to observe effects of internal or external parameters on a GFP–fusion protein, which do not result in a change of its localisation and stability but in a change of its nearby environment.

A complex system such as a living organism contains a manifold of components often coupled with each other e.g. in signal transduction or energy transfer pathways. Due to this interplay of several components, it often is highly complicated to investigate these components separately, particularly with regard to their exact amount and relative concentration. This can be done by extracting the components in question from their native environment but obviously results in the destruction of the intact organism. For the investigation of variations in the constitution of a coupled system due to changes in external parameters, it is essential to keep the system alive and, therefore, one has to develop methods to monitor the constitution of such a coupled system in vivo.

After an introduction into the topic and the applied experimental techniques given in Chapter 1 and 2, the biological background knowledge for the studies described in this work are presented in Chapter 3.

PART I: ocFL(I)M
In Part I, one–chromophore fluorescence lifetime (imaging) microscopy (ocFL(I)M) and its application to use a label protein as probe for its physico–chemical environment are presented. Using the example of BRI1–GFP, it was demonstrated that ocFLIM can be used to monitor cell–specific responses to a corresponding ligand, which do not result in a change of localisation and stability but in a change of the nearby environment of the fusion protein and enables the revelation of a ligand induced response pathway in living plants. Furthermore, a novel application of BRI1–GFP as a probe for the non–invasive determination of the membrane potential and the local P–ATPase activity is introduced. The experiments also show the potential of one–chromophore fluorescence lifetime imaging microscopy (ocFLIM) for the in vivo monitoring of not only the membrane potential but of other biochemical and biophysical parameters in the subcellular environment (pH–value and refractive index) using GFP fusion proteins as probes.

PART II: FIDSAM
Part II presents a novel method to suppress the autofluorescence in common fluorescence intensity images and therefore to enhance the signal–to–background ratio. The method is based on the well–known Fluorescence Lifetime Imaging Microscopy (FLIM) technique and is performed by decay shape analysis of the recorded fluorescence decays. The applicability of this method could be demonstrated impressively in several biological samples like human fibroblasts and plants, including some of the most challenging ones like AFP–fusion proteins in living plants expressed under their native promotor resulting in a very low label–to–autofluorescence ratio.

PART III: PSI/PSII Ratio
In Part III fluorescence spectroscopy is applied to the photosystems of living plants at room temperature. The photosystems are located in the chloroplasts, the energy factories of plants. Their composition is expected to change under extreme external conditions like a deficit of light or carbon, to optimize chloroplast efficiency. However, energy transfer between the various components results in the main detection of photons emitted by PSII, the last component of the energy migration chain and consequently, changes in the composition of the chloroplasts due to changed external parameters will result only in slight changes of the fluorescence spectra. By applying fluorescence microscopy at low temperature or by extracting the different components from their native environment, this energy transfer can be eliminated. However, due to this energy transfer, it is very difficult to investigate the components separately without destroying the intact, living system. By accomplishing statistical analysis to fluorescence spectra recorded at room temperature, it is possible to single out marginal differences in the fluorescence spectra and to determine a relative PSI/PSII ratio in living cells.

The possibility to determine the relative PSI/PSII ratio by fluorescence spectroscopy is demonstrated in living maize plants, which are known to exhibit mainly grana stacking and therewith mainly PSII in mesophyll cells. On the other hand, bundle sheath cells exhibit agranal thylakoid membranes and therewith mainly PSI. Due to these characteristics, maize plants are an appropriate system for the demonstration of the applicability of this method.

Afterwards, the method is transferred to several carbon deficient mutants to analyse the effects of light or carbon deficit on the PSI/PSII ratio. As a carbon or light deficit has to be compensated by the plant in some way, an adequate response would be e.g. changes in the composition of the photosystems to increase their efficiency.

PART IV: FExS
Based on the observations in carbon deficient mutants reported in PART III, fluorescence excitation spectroscopy (FexS) was performed in the same carbon deficient mutant plants. The photosystems of plants consist of a multitude of components coupled via energy transfer. Therefore these components cannot be investigated separately without destruction of the native environment. One approach to overcome this problem is Fluorescence Excitation (FEx) microscopy. Each component of an energy transfer chain such as the photosystems can be efficiently excited with different wavelengths. Thus, scanning the excitation wavelength continuously allows not only for the determination of the amount of components but even for their relative concentration with high spatial resolution. This makes FExS a more informative and therefore superior method compared to common fluorescence spectroscopy to determine differences in the composition of the photosystems.

PART V: smSERS on AFPs
In the last part of this thesis single molecule Surface Enhanced Raman Scattering (smSERS) on autofluorescent proteins (AFPs) is performed. SERS spectroscopy provides insight into the chemical structure of the investigated molecules and therefore is a competing method to fluorescence spectroscopy. However, SERS spectra of biological samples are often very complex and thus, the correct interpretation of the obtained data is very difficult. On the track to SERS in living cells it therefore is essential to initially focus on individual biological molecules instead of using a great variety of biological molecules as existent in living cells. Consequently, it is suggestive to investigate the AFPs used for labelling as this will later on enable the differentiation between label Raman bands and Raman bands deriving from native cellular components. Furthermore, the SERS spectra presented in this thesis confirm findings about the chemical structure of AFPs obtained by x–ray structure analysis and offer possible future applications of AFPs.
 

Zusammenfassung

In dieser Arbeit werden neuartige Verfahren zur Umgehung schwerwiegender Probleme vorgestellt, die der Untersuchung biologischer Proben mittels Fluoreszenz–Mikroskopie innewohnen. Das Hauptproblem besteht in der unspezifischen Autofluoreszenz, die das Signal des Fluoreszenz–Markers überstrahlen kann. Jede biologische Probe enthält einen gewissen Grad an Autofluoreszenz. Daher ist es in jeglichen lebenden Systemen, von einzelligen Organismen (z. B. Bak¬terien) bis hin zu Zellen höherer Organismen (z. B. Säugetieren) schwierig, hochauflösende und sensitive Fluoreszenz–Mikroskopie durchzuführen. Außergewöhnlich schwierige Bedingungen sind in lebenden Pflanzenzellen gegeben aufgrund einiger für Pflanzenzellen typischen, stark autofluoreszierenden Zellkompartimente (Zellwand, Chloro¬plasten).

Auch für den Fall, dass die störenden Einflüsse durch Autofluoreszenz in irgendeiner Weise umgangen werden können, erhält man aus Fluoreszenz–Intensitäts–Messungen hochaufgelöste Abbildungen der untersuchten Zellen, aber für gewöhnlich keine Informationen über physikalisch–chemische Parameter in der direkten Umgebung des markierten Fusionsproteins. Daher ist es schwierig, interne oder externe Auswirkungen auf ein GFP–Fusionsprotein zu untersuchen, die keine Veränderung der Lokalisation oder der Stabilität sondern eine Veränderung der direkten Umgebung zur Folge haben.

Ein komplexes System, wie es ein lebender Organismus ist, beinhaltet eine Vielzahl an Komponenten, die oftmals untereinander gekoppelt sind, z. B. zur Signalweiterleitung oder zum Energietransfer. Aufgrund dieses Zusammenspiels mehre¬rer Komponenten, ist es oft schwierig, diese Kom-ponenten getrennt voneinander zu untersuchen, besonders im Hinblick auf ihre genaue Anzahl und ihre relative Konzentra¬tion. Man kann die Komponenten aus ihrer natürlichen Umgebung entfernen und isolieren, jedoch nicht, ohne den Organismus zu zerstören. Will man Veränderungen in der Zusammensetzung eines solchen gekoppelten Systems in Abhängigkeit von externen Parametern untersuchen, ist es notwendig, das zu untersuchende System am Leben zu erhalten. Daher müssen Methoden entwickelt werden, die Zusammensetzung solcher gekoppelten Systeme in vivo zu untersuchen.
Nach einer allgemeinen Einführung in das Themengebiet und der Vorstellung der verwendeten experimentellen Methoden in Kapitel 1 und 2, wird in Kapitel 3 das biologische Hinter-grundwissen vorgestellt, das zum Verständnis der Experimente in dieser Arbeit notwendig ist.

TEIL I: ocFL(I)M
In Teil I wird one–chromophore Fluorescence Lifetime (Imaging) Microscopy (ocFL(I)M) vorgestellt anhand der Anwendung, ein Marker–Protein als Sensor für die direkte physikalisch–chemische Umgebung zu verwenden. Am Beispiel von BRI1–GFP wird gezeigt, dass ocFLIM verwendet werden kann, um zell–spezifische Reaktionen auf Ligandzugabe zu untersuchen, die keine Veränderung der Lokalisation oder der Stabilität sondern eine Veränderung der direkten Umgebung des Fusionsproteins zur Folge haben. Dies ermöglicht die Aufklärung Ligand–induzierter Signalwege in lebenden Pflanzen. Aus den gezeigten Experimenten ergibt sich, dass BRI1–GFP als Sensor zur nichtinvasiven Bestimmung des Membranpotentials und der lokalen P–ATPase Aktivität verwendet werden kann. Weiterhin ist es möglich, ocFLIM zur in vivo Bestimmung von anderen biochemischen und biophysikalischen Parametern (pH–Wert und Brechungsindex) auf subzellulärer Ebene zu verwenden, durch Einsetzen von GFP–Fusionsproteinen als Sensoren.

TEIL II: FIDSAM
Teil II beschreibt eine neue Methode, Autofluoreszenz in herkömmlichen Fluoreszenz–Intensitäts–Bildern zu unterdrücken und damit das Signal–Hintergrund–Verhältnis zu verbessern. Die Methode basiert auf der bekannten Fluorescence Lifetime (Imaging) Microscopy (FLIM) Technik und bedient sich der Zerfallsformanalyse der aufgenommenen Fluoreszenzzerfälle. Die Eignung dieser Methode wird eindrucksvoll an verschiedenen biologischen Proben wie z. B. an menschlichen Fibroblasten gezeigt. Zu den untersuchten Proben gehört eine der größten Herausforderungen für die Fluoreszenz–Mikroskopie: AFP–Fusionsproteine in lebenden Pflanzen, die unter nativem Promotor exprimiert werden und daher ein sehr niedriges Signal–Hintergrund–Verhältnis aufweisen.

TEIL III: PSI/PSII Verhältnis
In Teil III wird Fluoreszenz–Spektroskopie auf die Photosysteme lebender Pflanzen bei Raumtemperatur angewendet. Die Photosysteme befinden sich in den Chloroplasten, den Energiefabriken der Pflanzen. Seine Zusammensetzung sollte sich unter extremen äußeren Bedingungen wie Licht– oder Kohlenstoff–Mangel verändern, aufgrund der Notwendigkeit, die Effizienz der Chloroplasten zu erhöhen. In Folge von Energietransfer zwischen den Komponenten dieses Systems werden bei Fluoreszenz–Spektroskopie hauptsächlich Photonen detektiert, die von PSII emittiert wurden. Daher haben Veränderungen in der Zusammensetzung der Chloroplasten aufgrund äußerer Bedingungen nur geringe Veränderungen im Fluoreszenz–spektrum zur Folge. Mit Hilfe von Fluoreszenz–Spektroskopie niedrigen Temperaturen oder durch Isolierung der einzelnen Komponenten kann der Energietransfer unterbunden werden. Dennoch ist es aufgrund des Energie¬transfers sehr schwierig, die Komponenten getrennt voneinander zu untersuchen, ohne das lebende System zu zerstören. Durch statistische Analyse von bei Raumtemperatur aufgenommenen Spektren ist es möglich, geringfügige Unter-schiede der Spektren herauszuheben und so das relative PSI/PSII Verhältnis in lebenden Zellen zu bestimmen.

Die Möglichkeit, das relative PSI/PSII Verhältnis mittels Fluoreszenz–Spektroskopie zu bestimmen, wird hier anhand von lebenden Mais–Zellen gezeigt, die in Mesophyll–Zellen hauptsächlich Grana Stacks und damit PSII aufweisen, in Leitbündellzellen dagegen hauptsächlich PSI. Diese Eigenschaft macht Mais–Zellen zu einem angemessenen System um die Eignung der Methode zu beweisen.

Anschließend wird die Methode auf verschiedene Kohlenstoff–Mangel–Mutanten übertragen und die Einflüsse von Licht– oder Kohlenstoff–Mangel auf das PSI/PSII Verhältnis studiert. Da ein Kohlenstoff–Mangel auf irgendeine Weise von der Pflanze kompensiert werden muss, wäre eine angemessene Reaktion die Veränderung der Zusammensetzung der Chloroplasten zur Erhöhung ihrer Effizienz.

TEIL IV: FExS
Basierend auf den Beobachtungen in Kohlenstoff–Mangel–Mutanten in TEIL III wurde an den gleichen Mutanten Fluoreszenz–Anregungs–Spektroskopie (Fluorescence Excitation Spectroscopy: FExS) durchgeführt. Die Photosysteme von Pflanzen bestehen aus einer Vielzahl von Komponenten, die über Energietransfer miteinander gekoppelt sind. Daher können diese Komponenten nicht getrennt untersucht werden, ohne ihre natürliche Umgebung zu zerstören. Eine Möglichkeit, dieses Problem zu umgehen ist die Fluoreszenz–Anregungs–Mikroskopie. Jede Komponente einer Energietransfer–Kette wie die Photosysteme kann bei einer anderen Wellenlänge effizient angeregt werden. Daher kann durch kontinuierliches Verändern der Anregungs–Wellenlänge nicht nur die genaue Anzahl der Komponenten sondern auch ihre relative Konzentration mit hoher räumlicher Auflösung bestimmt werden. Diese Eigenschaften machen FExS zu einer konkurrenzfähigen und aufschluss–reicheren Methode als Fluoreszenz–Spektroskopie, um unterschiedliche Zusammen-setzungen der Photosysteme zu untersuchen.

TEIL V: smSERS an AFPs
Im letzten Teil dieser Arbeit werden Surface Enhanced Raman Scattering (SERS) Experimente an autofluoreszierenden Proteinen (AFPs) auf Einzelmolekül–Ebene (single molecule: sm) durchgeführt. SERS–Spektroskopie bietet Einblick in die chemische Struktur der untersuchten Moleküle und ist daher eine aufschlussreichere Methode als Fluoreszenz–Spektroskopie. Leider sind SERS–Spektren von biologischen Molekülen oft sehr komplex und daher sehr schwer zu interpretieren. Auf dem Weg zu SERS–Anwendungen in lebenden Zellen ist es daher notwendig, sich zu Anfang auf einzelne biologische Moleküle zu konzentrieren anstatt eine große Vielzahl an Molekülen zu untersuchen, wie sie in lebenden Zellen herrscht. Daher ist es naheliegend, AFPs zu untersuchen, die als Fluoreszenz–Marker verwendet werden, da diese Untersuchungen später die Unterscheidung zwischen Marker–Raman–Banden und Raman–Banden von natürlichen Zellmaterial ermöglichen werden. Weiterhin bestätigen die in dieser Arbeit gezeigten SERS–Spektren Ergebnisse über die chemische Struktur von AFPs, wie sie über Röntgen-strukturanalyse erhalten wurden und zeigen mögliche zukünftige Anwendungen für AFPs auf.

 

Table of Contents

1    Introduction    1
1.1. Scope of this Thesis    1
1.2. Outcome of this Thesis    4
1.3. Outline of this Thesis    6

2    Instrumentation and Applied Techniques    9
2.1. Confocal Microscopy    9
2.2. Fluorescence Spectroscopy    11
2.3. Raman Spectroscopy    14
2.3.1. Basics of Raman Spectroscopy    14
2.3.2. Surface Enhanced Raman Spectroscopy    16
2.4. Time Correlated Single Photon Counting    18
2.5. Least–Square Analysis    20

3    Biological Background Information    23
3.1. Structure of Plant Cells    23
3.2. Autofluorescent Proteins    24
3.3. Transformation of Plant Cells    25
3.3.1. Stable Transformation via Agrobacterium tumefaciens    26
3.3.2. Transient Transformation via Agrobacterium tumefaciens    28
3.4. Plant Chloroplasts    29
3.4.1. Structure of Plant Chloroplasts    29
3.4.2. The Function of Light–Harvesting Complexes and Photosystems in Chloroplasts    30

PART I: ocFL(I)M    33

4    Rapid BL–Induced Cell Wall Expansion and Lifetime Decrease of BRI1–GFP    33
4.1    Differentiation of Signal and Autofluorescence Background    34
4.2    Rapid BL–Induced Cell Wall Expansion in vivo    38
4.3    The Degree of BL–Induced Cell Wall Expansion Depends on the in vivo Level of BRI1–GFP    43
4.4    Monitoring the Environment of Plasma Membrane–Located GFP Fusion Proteins in vivo    45
4.5    BL–Induced Responses are Cell Physiological Processes    48
4.6    Measurement of Cell Wall Extension at Single Cell Level at Subcellular Resolution    49
4.7    Summary    50

5    BRI1–GFP as Non–Invasive Membrane Potential Sensor    51
5.1    Physico–Chemical Parameters Influencing the Fluorescence Lifetime of BRI1–GFP    52
5.2    Dependence of the Fluorescence Lifetime of GFP on the Applied Electric Field    54
5.3    Confirmation by Electrophysiological Experiments    56
5.4    Summary    57

6    A Short Brassinolide–Regulated Response Pathway in the Plasma Membrane of Arabidopsis thaliana    59
6.1    Inhibition of the BL–Induced Responses of
BRI1–GFP by Sodium Acetate    60
6.2    Effect of Auxin on BRI1–GFP Expressing Plant Cells    61
6.3    BL–Induced Responses of BRI1–GFP Depend on P–ATPase Activity    62
6.4    Quantitative Determination of Cell Wall Expansion    66
6.5    FRET Studies Reveal Interaction of BRI1–GFP with the P–ATPase    67
6.7    Regulation of BL–Induced Cell Wall Expansion is a Complex Process    69
6.6    BRI1 may Regulate the P–ATPase Activity by Protein–Protein Interaction    70
6.7    Summary    71

PART II: FIDSAM    73

7    FIDSAM* for Quantitative and Sensitive Live–Cell Imaging    73
7.1    Conventional Techniques for Resolution and Contrast Enhancement in Fluorescence Microscopy are Corrupted by Autofluorescence of Living Plant Cells    75
7.2    Physical and Optical Principle of FIDSAM    77
7.3    Application of FIDSAM in Living Plant Cells    80
7.4    Apparent Membrane Drifting    83
7.5    Validation of FIDSAM by Several Control Tests    85
7.6    CILA (Cell Imaging by Lifetime Analysis)    87
7.7    FIDSAM is Capable of Suppressing Chloroplast Autofluorescence    89
7.8    FIDSAM in Endogenously Expressing Plant Cells    91
7.9    FIDSAM on Fluorescent Labels with Biexponential Fluorescence Intensity Decays    93
7.10    Summary    96

8    Combining ocFLIM and FIDSAM Reveals Fast and Dynamic Physiological Responses at Subcellular Resolution in Living Plant Cells    97
8.1    Standard Confocal Microscopy Reveals the Occurrence of Green Fluorescent Cellular Compartments in eGFP–LTI6b Expressing Cells after Induction by Salt Stress    98
8.2    3D–Fluorescence Intensity Image Obtained by Spinning Disc Microscopy    101
8.3    Effect of different eGFP expression levels on the signal–to–background ratio in FIDSAM    101
8.4    FIDSAM reveals two different kinds of response on salt stress    105
8.5    ocFLIM Reveals Salt Stress Induced Functional Dynamics in the Cell Membranes    106
8.6    LTI6b may prevent hyperpolarization of the plasma membrane    108
8.7    Summary    110


PART III: PSI/PSII Ratio    111

9    Determination of the PSI/II Ratio in Living Plant Cells at Room Temperature by Fluorescence Spectroscopy    111
9.1    Determination of PSI/PSII Ratio in Living Maize Plants    112
9.2    Differentiation between Wild Type and Carbon Deficient Chloroplasts via Frequency Resolved Fluorescence Spectroscopy    114
9.3    Summary    117

PART IV: FExS    119

10    Fluorescence Excitation Spectroscopy (FExS) in Chloroplasts of Living Plant Cells    119
10.1    Basic Idea and Principle of Fluorescence Excitation Spectroscopy    120
10.2    Fluorescence Excitation Spectroscopy in Carbon Deficient Plant Chloroplasts    122
10.3    Summary    124

PART V: smSERS on AFPs    126

11    New Insights into the Molecular Structure of mRFP and eYFP by Single Molecule SERS Spectroscopy    127
11.1    The Aim: SERS in Living Cell    128
11.2    Averaged Spectra of Single AFP Molecules    129
11.3    eYFP–Specific Low Vibrational Energy Bands    132
11.4    Single Molecule Raman Spectra of AFPs    133
11.5    Summary    140

12    Summary of the Thesis    143

13    Zusammenfassung    147

Appendix 1:    Materials and Methods    151
A.1.1    Plant Material and Growth Conditions (BRI1, LTI6b, PMA4, BAK1, Col–0)    151
A.1.2    Plant Material and Growth Conditions (Carbon Deficient Mutants, Wild Type)    151
A.1.3    Optical and Spectroscopic Measurements    151
A.1.4    Sample Preparation for TEM    152
A.1.5    Sample Preparation for Electophysiological Measurements    152
A.1.6    Transient transformation of Nicotiana benthamiana leaves    153
A.1.6    Sample Preparation for smSERS    153
A.1.7    Data Analysis    154

Appendix 2:    FWHM Calculation    155
A.2.1    Apparent FWHM of a Plasmalemmata–Cell Wall Section in a 2D–Fluorescence–Intensity Image of a GFP–Labelled Membrane–Located Protein in Plant Cells    155
A.2.2    Apparent FWHM of a Plasmalemmata–Cell Wall Section in the Case of Tilted Membranes    158

Appendix 3: Error Calculation for Monoexponential Lifetime Fits    159
A.4.1    Fitting Error According to Gauss    159
A.4.2    Experimental Error    161

Appendix 4:    Scientific Contributions    163
A.4.1    List of Publications    163
A.4.2    Manuscripts under Review    164
A.4.3    Manuscripts in Preparation    165
A.4.4    Oral Presentations    165
A.4.5    Poster Presentations    166
A.4.6    Patent Application    166

Appendix 5: Curriculum Vitae    167

Appendix 6: Academic Teachers    168

References    169


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