The role of the ATP synthase in photosynthetic flux control Nádia

The role of the ATP synthase in photosynthetic flux control Nádia

Universidade de Lisboa
Faculdade de Ciências
Departamento de Biologia Vegetal
The role of the ATP synthase in photosynthetic flux control
Nádia Martins
Master Thesis in Celular Biology and Biotechnology
2009
1
This work was done at the Max-Planck-Institute of Molecular Plant Physiology, Potsdam under the supervision of
Dr. Mark Aurel Schöttler from October 2008 until July 2009.
1. External Supervisor: Dr. Mark Aurel Schöttler –Max Planck Institute of Plant Physiology, Golm,
Potsdam, Germany
2. Internal Supervisor: Dr. Jorge Marques da Silva- Departamento de Biologia Vegetal. Faculdade
de Ciências da Universidade de Lisboa
2
“Há um tempo em que é preciso abandonar as roupas usadas, que já têm a forma do nosso corpo, e esquecer os nossos
caminhos, que nos levam sempre aos mesmos lugares. É o tempo da travessia: e, se não ousarmos fazê-la, teremos ficado,
para sempre, à margem de nós mesmos”
Fernando Pessoa
3
Abstract
The role of the ATP synthase in photosynthetic flux control
Higher plants adjust the photosynthetic ATP and NADPH production to the metabolic consumption by
the Calvin cycle, as otherwise, an imbalance could result in the generation of reactive oxygen species,
ultimately leading to cell death. Three components of the electron transport chain (cytochrome-bf
complex, plastocyanin and ATP synthase) are closely adjusted to the metabolic demands (Schöttler et al,
2004; Schöttler et al, 2007). While it is well established that the cytochrome-bf complex exerts
photosynthetic flux control, the close adjustment of ATP synthase contents was surprising. In order to
determine the impact of altered ATP synthase contents on assimilation capacity, transformants with
decreased amounts of ATPase were generated. First, we manipulated the accumulation of the plastomeencoded
subunit. Eight different mutations were introduced into the atpB 5’-UTR, as previous in vitro
translation assays revealed changes in translation rates to between 10 and 200% of the wild-type level.
Surprisingly, none of the transplastomic plants showed clear differences in ATP synthase activity and
growth. This could be due to the existence of specific translation initiation factors in planta, which were
not considered in the in vitro translation assays. Alternatively, the
subunit might not be limiting for
ATP synthase biogenesis.
In a second approach, an antisense repression of the nuclear encoded
subunit resulted in plants with
strongly reduced assimilation and growth rates. This phenotype strictly correlates with the remaining ATP
synthase activity, but is not simply attributable to a reduced ATP availability. Rather, ATP synthase
repression results in an increased PMF above the thylakoid membrane, triggering ―photosynthetic
control‖ already at low light intensities: As plastoquinone reoxidation at the cytochrome-bf complex is
inhibited by strong lumen acidification, both ATP and NADPH production by linear electron flux are
decreased.
5 Key words: photosynthesis, electron transport, ATP synthase, photosynthetic flux control, proton
motive force.
4
Síntese
O papel da ATP sintase no controle do fluxo fotossintético
As plantas superiores ajustam a produção fotossintética de ATP e NADPH ao seu consumo metabólico
pelo ciclo de Calvin, já que um desequilíbrio pode resultar na geração de espécies reactivas de oxigénio,
levando em última instância à morte celular. Três dos componentes da cadeia transportadora de electrões
(complexo citocromo-bf, plastocianina e ATP sintase) ajustam o seu conteúdo às necessidades
metabólicas (Schöttler et al, 2004; Schöttler et al, 2007). Embora esteja bem provado que o complexo
citocromo-bf exerce controle no fluxo fotossintético, o ajuste do conteúdo da ATP sintase foi
surpreendente. De forma a determinar o impacto de alterações no conteúdo da ATP sintase na
assimilação, transformantes com quantidades diminuídas de ATP sintase foram gerados. Numa primeira
abordagem a acumulação da subunidade
da ATP sintase, codificada no cloroplasto, foi manipulada.
Oito mutações diferentes foram introduzidas na regiao 5'-UTR do gene atpB, já que ensaios anteriores,
feitos in vitro, revelaram alterações nas taxas de tradução entre 10 e 200% em relação ao tipo selvagem.
Surpreendentemente nenhuma das plantas transplastómicas mostrou diferenças claras em termos de
crescimento e actividade da ATP sintase. Isto poderá dever-se à existência de factores específicos de
tradução in planta, que não foram considerados nos ensaios in vitro. Alternativamente, a subunidade
pode não ser limitante para a biogénese da ATP sintase em tabaco.
Numa segunda abordagem, uma repressão antisense da subunidade , codificada no núcleo, resultou em
plantas com taxas de crescimento e assimilação extremamemente reduzidas. Este fenótipo correlaciona-se
com a restante actividade da ATP sintase mas não é unicamente atribuída a uma reduzida disponibilidade
de ATP. Aqui a repressão de ATP sintase resulta numa elevada força protomotriz na membrana tilacoidal,
desencadeando um controle fotossintético já em situações de fraca luminosidade: Já que a reoxidação da
plastoquinona ao nível do complexo citocromo-bf é inibida por uma forte acidificação do lúmen, tanto a
produção de ATP como a de NADPH pelo fluxo linear de electrões, decaem.
5 Palavras-chave: fotossíntese, transporte eletrónico, ATP sintase, controle do fluxo fotossintético, força
proto motriz.
5
Resumo
O papel da ATP sintase no controle do fluxo fotossintético
As plantas superiores ajustam a produção cloroplastidial de ATP e NADPH ao seu consumo metabólico
pelas reacções subsequentes (e.g. ciclo de Calvin), já que, de outra forma, o desequilíbrio gerado poderia
resultar na formação de espécies reactivas de oxigénio e, em última instância, em danos oxidativos no
aparelho fotossintético e morte celular. Assim sendo, com o aumento da idade foliar e diminuição da
fixação de carbono a capacidade da cadeia transportadora de electrões decai proporcionalmente (Schöttler
et al, 2004). Este ajustamento é conseguido através de alterações no conteúdo membranar do complexo
citocromo-bf, plastocianina e ATP sintase, que diminuem em paralelo com a capacidade assimilatória
(Schöttler et al, 2004; Schöttler et al, 2007). O complexo citocromo-bf e a plastocianina já mostraram ser
pontos de controlo do fluxo electrónico fotossintético (Price et al, 1998; Schöttler et al, 2004). Assim, a
estreita correlação entre o seu conteúdo e a capacidade assimilatória não foi surpreendente. No entanto, o
ajustamento adicional da ATP sintase foi algo cuja relevância fisiológica não foi totalmente
compreendida (Anderson et al, 1988; Schöttler et al, 2007).
Para resolver esta questão e fazer uma análise mais detalhada do papel da ATP sintase na limitação da
fotossíntese, foram geradas plantas com conteúdo reduzido deste complexo proteico. A alteração do
conteúdo foi feita através da manipulação da acumulação de duas subunidades do sector catalítico da
enzima. Primeiro manipulou-se a acumulação da subunidade , codificada no cloroplasto. A acumulação
desta subunidade é controlada ao nível da tradução e especialmente o seu início é altamente regulado.
Esta subunidade mostrou ser limitante na montagem da ATP sintase em Chlamydomonas reinhardtii
(Drapier et al, 2007). Por esta razão, decidiu-se alterar o início da tradução modificando o sítio de ligação
do ribossoma na região 5' não traduzida do mRNA e, também o codão de iniciação da tradução. Oito
mutações pontuais diferentes foram introduzidas nesta região, já que ensaios prévios feitos in vitro
mostraram alterações nas taxas de tradução que variavam entre 10 % e 200 % em relação ao tipo
selvagem. Numa segunda abordagem, utilizou-se um mutante com uma repressão antisense do gene
nuclear atpC, que codifica para a subunidade . Esta subunidade tem também um papel central na
biogénese da ATP sintase em Chlamydomonas (Drapier et al, 2007).
O objectivo desta tese foi determinar o impacto da alteração do conteúdo da ATP sintase na sua
actividade e também na taxa de assimilação e transporte electrónico. Para além disso, a hipótese de que a
6
ATP sintase limita o transporte electrónico fotossíntético e regula a activação dos mecanismos
fotoprotectores foi também investigada.
Para atingir este fim utilizaram-se métodos espectroscópicos que permitem medir a actividade da ATP
sintase in vivo, através da medição da força protomotriz ao nível da membrana dos tilacóides. Para além
disto, determinou-se a capacidade de assimilação e a eficiência quântica de assimilação de dióxido de
carbono através de medições de trocas gasosas. A actividade da ATP sintase nestes transformantes foi
determinada através de espectroscopia de absorção diferencial. Também se determinou qual a força
protomotriz máxima capaz de ser gerada e a sua partição nos componentes químico e eléctrico ( pH e
). Obtiveram-se ainda curvas de resposta à luz da taxa de transporte electrónico fotossintético, do
coeficiente de amortecimento não fotoquímico (qN), e do estado redox do lado aceitador do fotossistema
II (qL). Outros parâmetros fotoquímicos e.g. Fv/Fm, foram também determinados. Também se mediu,
através de espectroscopia de absorção diferencial, a acumulação dos componentes da cadeia
transportadora de electrões (fotossistema II, plastocianina, complexo citocromo-bf e fotossistema I) para
verificar se a alteração do conteúdo de ATP sintase afecta a sua acumulação. Finalmente, as curvas de
redução do complexo citocromo-bf e do P700, em diferentes intensidades luminosas foram medidas, para
verificar se um aumento da força protomotriz e uma mais rápida acidificação do lúmem tilacoidal, em
plantas com conteúdo reduzido de ATP sintase, resulta na inibição do transporte eletrónico através de
mecanismos de controlo fotossintético (e.g. impedimento da reoxidação do plastoquinol no complexo
citocromo-bf). Adicionalmente, mediu-se a acumulação de amido nestes mutantes, para detectar
diferenças na quantidade de ATP disponível.
Este conjunto de medições deveria permitir testar a hipótese de que o conteúdo de ATP sintase é um
importante ponto de controlo do transporte electrónico fotossintético e que regula o aparecimento de
mecanismos fotoprotectores.
Surpreendentemente, nenhuma das linhas com mutaçoes introduzidas no gene atpB, mostrou diferenças
significativas em termos de fenótipo e actividade da enzima. Esta ausência de fenótipo deve-se,
provavelmente, ao facto da actividade deste complexo não depender exclusivamente do seu conteúdo na
membrana, sendo altamente regulada. Estes mutantes revelaram, contudo, um ligeiro aumento da força
protomotriz em relação à linha selvagem, em particular uma maior acumulação do seu componente
químico, já que este é o responsável pela activação dos mecanismos fotoprotectores. Nestes mutantes a
maior acumulação de força protomotriz deve-se à ligeira repressão do fluxo electrónico linear pela ATP
sintase e fez com que os mecanismos fotoprotectores fossem iniciados mais rapidamente.
Nestes mutantes a composição da cadeia transportadora de electrões não sofreu alterações. Ambos os
fotossistemas mantiveram o seu conteúdo e o fotossistema II não perdeu função. O conteúdo em
citocromo-bf e PC também não sofreu alterações significativas.
7
Apesar da ausência dum fenótipo visível e de efeitos fisiológicos significativos, algumas linhas
apresentavam uma menor acumulação de amido. Isto revela que havia nestas linhas menor
disponibilidade de ATP, o que se deve, provavelmente, a um menor conteúdo de ATP sintase.
Embora quando considerados individualmente os parâmetros fotoquímicos das diversas linhas mutantes
não se diferenciem significativamente do tipo selvagem, quando observamos o quadro global é possível
detectar, em algumas linhas, comportamentos fisiológicos consistentemente distintos.
De um modo geral, podemos considerar que a redução do conteúdo de ATP sintase ficou aquém da
esperada. Isto pode significar que a situação in vitro é distinta da situação in vivo, isto é, que a existencia
de factores de iniciação da tradução in planta não foi adequadamente considerada nos ensaios in vitro.
Alternativamente, a subunidade
pode não ser limitante para a biogénese do complexo ATP sintase em
Nicotiana tabacum, ao contrário de Chlamydomonas.
A linha com uma mutação na subunidade
originou plantas com taxas de crescimento e assimilação
reduzidas. Este fenótipo tem a ver com a restante actividade da enzima, embora não seja unicamente
atribuível à reduzida disponibilidade de ATP (Schöttler et al, 2009, sob revisão). Nestes mutantes, a
repressão da actividade da ATP sintase faz com que o lúmen tilacoidal fique fortemente acidificado,
fazendo com que os mecanismos fotoprotectores, tal como o amortecimento não fotoquímico, sejam
espoletados logo em condições de fraca luminosidade. Enquanto que a acumulação dos outros
componentes da cadeia fotossintética se mantém basicamente inalterada, o fluxo linear de electrões é
fortemente reprimido devido a um controlo fotossintético, isto é, à inibição da reoxidação da
plastoquinona ao nível do citocromo-bf. E como a reoxidação da plastoquinona ao nível do complexo-bf
é inibida pela forte acidificação do lúmen, tanto a produção de ATP como a de NADPH, pela cadeia
transportadora de electrões, diminuiem. Assim, concluí-se que a limitação de ATP resulta numa reduzida
conductividade da membrana tilacoidal, aumentando assim a força protomotriz, reprimindo o crescimento
da planta, devido a uma inibição do fluxo electrónico linear e ao aumento da dissipação da energia na
forma de calor nas antenas.
8
Index
1. Introduction ........................................................................................................................................................... 11
1.1 ATP synthase ................................................................................................................................................... 12
1.1.1 Structure of the ATPase complex ........................................................................................................... 13
1.1.2 Functioning mechanism of ATPase ........................................................................................................ 15
1.1.3 Assembly of ATP synthase ...................................................................................................................... 16
1.1.4 Regulation of ATP synthase .................................................................................................................... 17
1.1.5 Role of ATP synthase ............................................................................................................................... 20
1.2 Objective .......................................................................................................................................................... 21
2. Material and methods............................................................................................................................................ 22
2.1 Plant material and growth conditions ........................................................................................................... 22
2.2 Constructs ........................................................................................................................................................ 22
2.3 Thylakoid membrane isolation....................................................................................................................... 23
2.4 Quantification of photosynthetic complexes ................................................................................................. 23
2.4.1 Quantification of PSII and Cytochrome-b6f complex .......................................................................... 23
2.4.2 Quantification of P700 ............................................................................................................................. 25
2.4.3 Quantification of PC/P700....................................................................................................................... 25
2.5 P700 and PC reduction kinetics ..................................................................................................................... 26
2.6 77K fluorescence measurement ...................................................................................................................... 26
2.7 Fluorescence measurement ............................................................................................................................. 26
2.8 PMF measurements and ATP synthase activity ........................................................................................... 27
2.9 Gas exchange measurements .......................................................................................................................... 28
2.10 Metabolite quantification.............................................................................................................................. 29
2.10.1 Ethanol extraction of soluble sugars .................................................................................................... 29
2.10.2 Starch extraction .................................................................................................................................... 29
2.10.3 Metabolite quantification ...................................................................................................................... 30
3. Results..................................................................................................................................................................... 32
3.1 Growth phenotype ........................................................................................................................................... 32
3.2 Chlorophyll content......................................................................................................................................... 33
3.3 ATPase activity ................................................................................................................................................ 34
3.4 Assimilation capacity ...................................................................................................................................... 35
3.5 Assimilation capacity versus membrane conductivity ................................................................................. 36
3.6 Organization of the electron transport chain................................................................................................ 37
3.7 Maximum PSII efficiency ............................................................................................................................... 38
9
3.8 77K chlorophyll-a fluorescence emission spectra ......................................................................................... 39
3.9 PMF partitioning ............................................................................................................................................. 40
3.10 Light response curves.................................................................................................................................... 41
3.11 Reduction kinetics of Cytochrome-f and P700........................................................................................... 43
3.12 Metabolite quantification.............................................................................................................................. 44
4. Discussion ............................................................................................................................................................... 46
5. Outlook ................................................................................................................................................................... 53
6. Abbreviations ......................................................................................................................................................... 54
7. Literature ............................................................................................................................................................... 56
8. Acknowledgments .................................................................................................................................................. 60
10
1. Introduction
Oxygenic photosynthesis is a highly regulated energy-transducing process. In photosynthetic
eukaryotes it takes place in the chloroplast, in which light energy is captured and converted into
biochemical energy. This multi-step process involves a series of electron transfer reactions
resulting from water oxidation, at the PSII (Photosystem II) lumen side, to NADP+ on the stromal
side of PSI (Photosystem I) (Figure 1).
This energy conversion involves the absorption of photons by antenna pigments in the light
harvesting complexes (LHC’s), which then transfer the excitation energy to the reaction centers
of PSII and PSI, driving the primary photochemical reactions and creating charge separation
(Baker et al, 2007). The light-driven charge separation, at the photosystems, is responsible for
the electron flow between PSII and PSI through plastoquinol (PQ), cytochrome-b6f complex
(Cyt-b6f) and plastocyanin (PC) (Szabó et al, 2005; Baker et al, 2007). Coupled to this linear
electron flux (LEF) is the release of protons during water oxidation at the lumen side of PSII, and
the influx of protons into the thylakoid lumen due to the Q-cycle at the cyt-b6f. These events
allow the establishment of an electrochemical potential difference, or proton motive force
(PMF), across the thylakoid membrane, composed of both electric field (∆Ψ) and concentration
(∆pH) gradients, which is used by ATP synthase (ATPase), to transport protons back into the
stroma, driving ATP synthesis from ADP and Pi (Baker et al, 2007). The products from this
photosynthetic electron transport chain (ETC), ATP and NADPH, are subsequently consumed by
metabolic reactions, like the Calvin cycle and photorespiration (Baker et al, 2007).
This energy conversion has to meet the demands for ATP and NADPH of the downstream
metabolic processes, including not only the reduction of CO2 to the level of sugar phosphates in
the Calvin cycle, but also photorespiration, nitrogen assimilation, etc, each with a different
relative requirement for ATP and NADPH (Avenson et al, 2005; Noctor and Foyer, 2000). These
metabolic reactions must be regulated to prevent the build-up of reactive oxygen species (ROS),
which in sufficiently high concentrations can lead to photoinhibition (photodamage), or even kill
the plant (Kim et al, 2008). Taking this into account, it is clear that the capacity of the
photosynthetic light reactions to provide ATP and NADPH needs to be closely adjusted to their
metabolic consumption (Kim et al, 2008; Schöttler et al, 2004). The photosynthetic apparatus is
sufficiently dynamic and able to respond to environmental changes, allowing the plant to adapt.
11
Although it is well established that the cytochrome-bf complex and PC catalyse the rate-limiting
steps of photosynthetic electron flux (Anderson, 1992; Kirchhoff et al, 2000; Schöttler et al,
2004), until now the role of ATPase in controlling assimilation rates remains elusive. In spite of
this, it is known that the ATPase contents closely correlate with assimilation capacities, e.g. after
changes in light quantity/quality and during leaf aging (Anderson et al, 1988; Schöttler et al,
2007).
stroma
lumen
Figure 1-Composition of the thylakoid membrane and linear electron transport. From Joly and Carpentier
2004.
1.1 ATP synthase
The chloroplast ATPase is a protein complex, located within the chloroplast thylakoid
membranes, which synthesizes ATP from ADP and inorganic phosphate (Pi) through a PMF
formed by the photosynthetic electron transport (Hisabori et al, 2002). This enzyme belongs to a
large family of F0F1 type ATPases and shares a common structure with the ATPases from
mitochondria and bacteria (Evron et al, 2000).
12
1.1.1 Structure of the ATPase complex
The general structure and composition of the complex is highly conserved among different
organisms as observed both in primary sequences of their subunits and their functional
characteristics. In the chloroplast it consists of two sectors: CF1 and CF0 (Ni et al, 2005;
Strotmann et al, 1998). The CF1 moiety is water soluble and extrinsic to the membrane, facing
the stromal side, and contains the nucleotide-binding and catalytic sites for ATP synthesis. The
CF0 is a transmembranal hydrophobic sector that includes a channel for the proton translocation
through the thylakoid membrane, which drives ATP synthesis (Hisabori et al, 2002; Ni et al,
2005). The CF1 sector consists of five different subunits with a stoichiometry
1
and the
CF0 sector is composed of a1b1b’1c14 (Del Riego et al, 2006; Richter et al, 2005).
The structure and subunit arrangement in CF1 and CF0 have been determined by various
techniques (Groth and Strotmann, 1999). The overall shape, dimension and mass distribution of
CF1 was determined by electron microscopy and image analysis (Boekema and Böttcher, 1992).
It was concluded that the CF1 sector is a hexagonal symmetrical ring, consisting of alternating
and
subunits with the central cavity of the hexamer being partially filled by the
subunit
(Groth and Strotmann, 1999) (Figure 2). Furthermore, electron microscopy studies in
subunit-
deficient CF1 mutants showed that this subunit is necessary to maintain the arrangement of the
central core (Akey et al, 1983). Also, the location and orientation of nucleotide binding sites and
regulatory sites in CF1 have been identified by photolabelling and fluorescence resonance energy
transfer and showed that these sites are located on each of the six
interfaces (Groth and Pohl,
2001).
Until now, no atomic resolution of CF1 or individual subunits has been obtained; hence most
detailed structural information about this complex comes from the high resolution structure of
the bovine mitochondrial ATPase (Abrahams et al, 1994). However, a similar structure may be
expected for CF1 due to the homology of subunits between different organisms (Groth and
Strotmann, 1999).
The transmembrane CF0 sector, which mediates proton translocation, consists of four different
subunits in a stoichiometry of a1b1b’1c14 (Hisabori et al, 2002). Low resolution structures giving
the shape, the dimensions and the relative arrangement of the subunits in the CF0 sector have
been obtained by electron microscopy (Fromme et al, 1987). Subunits c and a are directly
13
involved in transmembrane proton transport. Subunit b which can be crosslinked to subunit
(Beckers et al, 1992) and subunit b’ are likely to form a link between CF0 and CF1 (Groth and
Strotmann, 1999). Secondary structure prediction assumes transmembrane
-helical structures
for subunits a, b, b’ and c. Subunit b and b’ probably contain a single transmembrane helix
leaving an extensive polar region exposed to the chloroplast stroma. Subunit a is a very
hydrophobic protein, predicted to contain at least five transmembrane helices and almost no
extramembrane domains. Subunit c consists of two antiparallel transmembrane helices connected
by an extramembrane polar loop. Topological studies obtained by atomic force microscopy,
suggest that the c subunit forms a ring-shaped complex in the membrane, whereas subunits b, b’
and a are located outside the subunit c oligomer (Neff et al, 1997) (Figure 2).
The prevailing model for the structural organization of the CF0F1 complex is shown in Figure 2.
Subunits δ, b and b’ form together a stalk which binds CF1 to CF0. This stalk is considered to
hold the α, β and a subunits still while the γ, ε and c subunits rotate. The ε subunit, and part of
the γ subunit, together form a second, central stalk connected to the ring of c subunits. There are
six nucleotide binding sites, one at each of the αβ subunit interfaces about halfway along the
vertical axis of the hexamer. Three of the sites are located primarily on the β subunits and are
catalytic, the other three are noncatalytic and probably regulatory (Groth and Pohl, 2001).
aa
b‘
b
a
c
Figure 2- Schematic representation of ATP synthase and its subunits. In green tones are the subunits of the CF1
sector and in red tones are the subunits of the CF0 sector. In light green and red are the nucleus encoded subunits and
in dark green and red are the chloroplast encoded subunits. Modified from Groth and Strotmann, 1999.
14
1.1.2 Functioning mechanism of ATPase
The prevailing catalytic mechanism is known as the binding change mechanism and was
suggested by Paul Boyer (Boyer, 1993). In this mechanism three catalytic sites are thought to
interact cooperatively, alternating between tight binding, loose binding and unbound states
during one catalytic cycle. In ATP synthesis, the energy of the PMF is used to elicit structural
changes that cause the tight site to release ATP and become an open site. Coupled to this event,
the loose site that contains bound ADP and Pi becomes a tight site and the open site that was
unoccupied binds ADP and Pi to become a loose site (Figure 3). In a larger scale we have the
rotational catalysis mechanism which proposes that the c subunit oligomers of the ATPase rotate
together with the
and
subunits as protons flow through CF0. This rotation distributes the
asymmetrical interactions among the
subunits and the subunit, allowing the catalytic sites to
switch their properties (McCarty et al, 2000). On rotation of the γ subunit, the conformations of
the three β subunits change sequentially such that each β subunit successively adopts the
different conformations of varying affinity during one rotational cycle. As a result, three
molecules of ATP are synthesized during a full conformational cycle of CF1. In synthesis mode,
the F0 motor converts the electrochemical gradient of protons into torque, to force the F1 motor
to act as an ATP generator. In the hydrolysis mode, F1 converts the chemical energy of ATP
hydrolysis into a torque, causing the membrane-embedded F0 motor to act as an ion pump
(Dimroth et al, 2006).
Figure 3-The binding change mechanism of the ATPase. Three sites interact cooperatively to alternate between
tight (T), loose (L), and open (O) states. The energy (indicated by an asterisk) required for the synthesis of ATP is
used for both substrate binding and product release. From McCarty et al, 2000.
15
1.1.3 Assembly of ATP synthase
In contrast with the major subunits of PSI, PSII and cytochrome bc/b6f complexes which are
integral membrane proteins, the catalytic sector of the proton ATPase, F1, is made of extrinsic
subunits that assemble in a hydrophilic environment (Abrahams et al, 1994). The other sector,
F0, which behaves as a selective proton channel, is embedded in the membrane and assembles
within the lipid bilayer (Deckers-Hebestreit and Altendorf, 1996). One of the special
characteristics of the ATPase biogenesis is the requirement for an uneven stoichiometry between
subunits in both F1 and F0. In F1, 3 copies of the
of the ,
and
and
subunits assemble with only one copy
subunits, whereas 14 copies of subunit c assemble with 1 copy of subunits a, b
and b’ in F0 (Drapier et al, 2007). It is not known if CF0 and CF1 assemble independently or in a
coordinated way. In chloroplasts, both CF0 and CF1 biogenesis requires the assembly of both
nucleus and chloroplast-encoded subunits (Drapier et al, 2007; Rimbault, 2000). The CF0
subunits a, b’ and c are chloroplast-encoded, but b is nucleus-encoded. CF1 subunits ,
and
are expressed from the chloroplast genes atpA, atpB and atpE, respectively, while subunits and
are expressed from the nuclear genes atpC and atpD (Strotmann et al, 1998) (Figure 2).
The nucleus encoded subunits are expressed as soluble precursor proteins that are imported posttranslationally into the chloroplast with their transit sequences being cleaved before they
assemble into the enzyme complex (Strotmann et al, 1998). The chloroplast encoded subunits
and
have been shown to be translated by thylakoid associated polysomes in land plants and
Chlamydomonas reinhardtii (Strotmann et al, 1998).
Defects in the expression of any of the subunits leads to a pleiotropic loss of most polypeptides,
both from CF1 or CF0 (Lemaire and Wollman, 1989). Studies also suggested a tight coupling
between the rates of synthesis of several subunits. In the absence of one subunit, the synthesis of
all other subunits is reduced and the complex remains unassembled; the unassembled subunits
are prone to degradation (Drapier et al, 1992; Lemaire and Wollman, 1989). Recently it was
reported that subunit
activates the synthesis of its assembly partner, subunit
Chlamydomonas. For sustained translation of subunit
in
, the nuclear encoded g subunit is
required. This Control by Epistasy of Synthesis (CES) process comprises intertwining
transactivation and negative feedback loops that ensure the production of
oligomers in the
16
proper stoichiometry, required for their interaction with subunit . The final
3 3
stoichiometry
is required for the functional assembly of CF1 (Drapier et al, 2007).
So far the complete assembly of ATPase is not yet fully understood: It is not known if the first
step is the formation of the
3 3
complex with subsequent binding of
and subunits and later
assembly with the already formed CF0 in the membrane; or if otherwise there is binding of
subunits to the CF0 and later binding of the
3 3
and
complex to the stalk (Drapier et al, 2007;
Strotmann et al, 1998). One thing is certain: The biogenesis of the chloroplast ATPase requires
tight control mechanisms, which regulate both the stoichiometry of the subunits and the cross
talk between two distinct genetic compartments (Lemaire and Wollman, 1989; Drapier et al,
2007), ensuring the coordinated biogenesis of the complex.
1.1.4 Regulation of ATP synthase
Since ATP synthesis is a key reaction for the maintenance of a number of metabolic pathways,
the ATPase must be prone to several regulatory mechanisms, in order to modulate its activity
accordingly with the environmental changes (Konno et al, 2006). In the light, the enzyme
complex is activated in order to achieve a high capacity of ATP synthesis, whereas in the dark, it
is converted to an inactive state, thereby preventing the dissipative cleavage of stromal ATP (Wu
et al, 2007). To optimize photosynthetic productivity, particularly at limiting light availability, it
is necessary to balance and control light harvesting and the consumption of reducing power and
ATP in metabolic reactions. Therefore there must be a tight regulation between efficient energy
absorption and energy conversion, to avoid over reduction and damage of the cell (Oelze et al,
2008). In plant cells, there are at least 4 elements involved in the regulation of the chloroplast
ATPase: nucleotide binding and release to the catalytic and regulatory sites; thiol modulation;
proton transmembrane electrochemical potential (∆µH+) formation across the thylakoid
membrane and phosphorylation followed by 14-3-3 protein binding.
17
1.1.4.1 Nucleotide binding
One of the most common regulatory mechanisms of ATPase activity is the binding of ADP-Mg
to the catalytic sites of the enzyme (Bar-Zvi and Shavit, 1982; Evron et al, 2000; Malyan and
Vitseva, 1990). When tightly bound, this nucleotide is responsible for stabilization (Wang et al,
1993) and regulation of the activity of the enzyme (Bar-Zvi and Shavit, 1982). In the absence of
an electrochemical proton gradient, ADP-Mg binds to the catalytic site of ATPase acting as an
inhibitor and making it almost completely inactive. Upon illumination, and therefore PMF
formation, bound ADP-Mg is released and ATP is synthesized (Evron et al, 2000; Wu et al,
2007).
1.1.4.2 Thioredoxins and thiol modulation
The control of enzyme activity by the ferredoxin–thioredoxin system is the most studied example
of redox regulation in plant cells and was first identified in 1980 in chloroplast enzymes
(Buchanan and Balmer, 2005; Oelze et al, 2008). The subunit of the ATPase is responsible for
the so called thiol modulation or redox modulation in plants and green algae. This subunit
contains a domain of approximately 40 amino acids, in which two cysteines are included (Wu et
al, 2007). These two Cys residues are able to form a reversible intrapeptide disulfide bond,
which enables the regulation of the enzyme via redox thiol modulation (Dal Bosco et al, 2004,
Wu et al, 2007). When exposed to light, this disulphide bond is reduced through thioredoxin-f
thus activating ATPase and synthesizing ATP (Buchanan and Balmer, 2005; Wu et al, 2007).
Under dark conditions, sulfhydryl groups are oxidized to form a disulphide bond, and hence ATP
hydrolysis activity is suppressed. One requisite for the efficient reduction of the disulfide bond is
a transmembrane proton gradient. The energization of the membrane induces conformational
changes that allow the hidden disulfide bridge of the
subunit to become accessible to the
reductant, thereby activating it (Strotmann et al, 1998).
Even though the redox regulation of chloroplast ATPase activity has been extensively studied,
the in planta physiological function remains elusive since redox modulation is not a prerequisite
for either activation or deactivation of the chloroplast ATPase. Both activation and deactivation
18
of the enzyme are rapid processes that normally precede the thioredoxin-dependent
reduction/oxidation of subunit following changes in light conditions (Wu et al, 2007).
1.1.4.3 Thylakoid PMF
As a result of the linear and cyclic electron flux there is proton translocation through the
thylakoid membrane which leads to the formation of a PMF (Oelze et al, 2008; Takizawa et al,
2007). This proton gradient is not only responsible for release of inhibitory, tightly bound MgADP but also for the reduction of the disulfide bridge in the subunit (Evron et al, 2000).
In the presence of light, the generation of ∆µH+ induces conformational changes within the
ATPase complex causing activation of the enzyme. As ∆µH+ dissipates in the dark, these
conformational changes and ADP release are reversed, inactivating the enzyme and avoiding
energy losses (Kleefeld et al, 1990; Strotmann et al, 1998; Wu et al, 2007).
1.1.4.4 Phosphorylation and 14-3-3 protein binding
An additional way of regulation could be executed via 14-3-3 proteins, which are a family of
conserved regulatory proteins expressed in all eukaryotic cells.
14-3-3s were thought to be present only in the cytoplasm and on the plasma membrane, but now
it has been shown that they are also present in the stroma of the chloroplast (Sehnke et al, 2000).
In plants, consumers of energy and reducing power produced by the chloroplasts and
mitochondria e.g., the plasma membrane H1 ATPase and nitrate reductase (NR), as well as
protein import processes into plastids and mitochondria are all regulated by 14-3-3 proteins.
In 2001 Bunney and coworkers showed that the activity of F0F1 synthases from both plant
mitochondria and chloroplasts is controlled by interaction with 14-3-3 proteins. They suggested
that 14-3-3 proteins bind to phosphorylated
subunits, thereby preventing the rotation and
catalytic action of the ATPase complex (Bunney et al, 2001). The most probable targets are the
subunits since they undergo phosphorylation in vivo (Del Riego et al, 2006)
and are
phosphorylated by casein kinase II located in the chloroplast stroma (Reiland et al, 2009), but the
process by which 14-3-3 proteins regulate the ATPase still remains unknown.
19
1.1.1.5 Role of ATP synthase
In plants the synthesis of ATP is driven by a flux of protons through the ATPase, across the
membrane. This efflux of protons from the lumen is determined by the rate of their translocation
through the ATPase (Kramer et al, 2003). The enzyme gates the flow of protons out of the
thylakoid lumen and hence regulates the PMF, together with LEF/proton influx into the lumen.
The thylakoid PMF has a central role in plants cells as it can both drive ATP synthesis, as well as
trigger the initiation of photoprotective mechanisms such as non photochemical quenching
(NPQ) (Szabó et al, 2005) and photosynthetic control.
In excessive light conditions, the rate of carbon fixation is surpassed by the amount of available
light and the lumen becomes extremely acidic, leading to protonation of an antenna protein,
PsbS (Li et al, 2004) and also activation of violaxanthin deepoxidase, which converts
violaxanthin to antheraxanthin and finally zeaxanthin, also known as xanthophyll cycle (Oelze et
al, 2008). The formation of zeaxanthin together with the protonation of PsbS activates
photoprotective mechanisms, allowing the dissipation of excessive energy in the form of heat
(Müller et al, 2001; Szabó et al, 2005).
The regulation of PMF can occur by modulating either the light driven proton influx (into the
lumen) or the proton efflux (from the lumen) through the ATPase e.g. when the ATPase is
limited by ADP or Pi availability due to slow ATP consumption by the Calvin Cycle (Kramer et
al, 2004).
In the chloroplast the ATPase has a critical role in photosynthetic control, which is the inhibition
of plastoquinol reoxidation at the Cyt-bf due to lumen acidification. Since lumen acidification
counteracts the proton pumping reactions at the plastoquinone shuffle, there is a control of the
rate of electron donation to PSI (Laasch and Weis, 1989) (Figure 4).
The disturbance of ATPase can be harmful to the cell and hence must be tightly regulated in
order to prevent cellular death (Johnson, 2008).
20
Figure 4-Schematic representation of the electron (orange arrows) and proton transfers (blue arrows), and
associated processes that can occur as a result of light absorption by the thylakoid photosystems. From Baker
et al 2007.
1.2 Objective
Higher plants adjust the photosynthetic ATP and NADPH production to the metabolic
consumption by the Calvin cycle, as otherwise, an imbalance could result in the generation of
reactive oxygen species, ultimately leading to cell death. The content of certain electron transport
chain components, as cytochrome-bf complex, plastocyanin and ATPase, are closely adjusted to
the metabolic demands. Although it is well established that the cytochrome-bf complex controls
photosynthetic flux, an additional contribution of ATPase contents to flux control could also be
possible (Schöttler et al, 2007).
The present study aimed to determine the impact of altered ATPase contents on ATPase activity
and on assimilation capacity. Furthermore, the hypothesis that ATPase limits the photosynthetic
electron transport, and regulates the onset of photoprotective mechanisms should also be
investigated. Therefore transplastomic plants with decreased amounts of ATPase, through
manipulation of the accumulation of the plastome-encoded
subunit and nuclear encoded
subunit were generated. Both types of transformants were chosen due to their central role in
ATPase biogenesis as shown in Chlamydomonas (Drapier et al, 2007).
21
2. Material and methods
2.1 Plant material and growth conditions
Transgenic tobacco plants were generated within the PhD thesis of Markus Rott (AG Schöttler,
MPIMPP) and were available from the beginning of this master thesis. Both wild-type tobacco
(Nicotiana tabacum) plants from the cultivar Petit Havana (PH) and transformed plants were
grown in a controlled environment chamber. The actinic light intensity was 800 µE m-2 s-1, with
plants being illuminated daily for a period of 16h. During the day the relative humidity was 70%
and the temperature 22°C. During the night period the temperature was reduced to 18°C and the
relative humidity was 70%. An additional transgenic line, atpC1, (Lein et al, 2008), from the
cultivar Sansun (SNN), was grown under the same conditions.
2.2 Constructs
Eight mutations were introduced into the atpB 5’-untranslated region (UTR). Lines MJR1 and
MJR2 had a point mutation introduced into the atpB translation initiation codon. Lines MJR3,
MJR4, MJR5 and MJR6 had a point mutation in the 5´-Untranslated (UTR) region of the mRNA
of atpB. Line MJR7 had a deletion between base pair 25 and 53, and line MJR10 contained only
the selectable marker gene aadA, which in all transplastomic plants was introduced as additional
transcript, comprising both aadA and atpB, due to incomplete transcript accumulation in
chloroplasts in higher plants (Figure 5).
amp
5‘part of atpB
aadA
rbcL
WT
5‘UTR
MJR1
MJR2
ATT ATT ATG
atpB
5‘UTR
ATT ATT TTG
atpB
5‘UTR
ATT ATT GTG
atpB
MJR3
5‘UTR
ATT ACT ATG
atpB
MJR4
5‘UTR
ATT TTT ATG
atpB
MJR5
5‘UTR
ATT AAT ATG
atpB
MJR6
5‘UTR
ATT ATC ATG
atpB
MJR7
5‘UTR deleted ATT ATT ATG
atpB
MJR10
aadA control
Figure 5-Schematic figure of the constructs with the inserted point mutations in red (MJR1 to MJR6) and
the deletion (MJR7). amp- Ampicillin resistance gene; rbcL- The sequence of the large subunit of the ribulosebisphosphate carboxylase gene; aadA- Aminoglycose resistance protein; WT- Wild type; ATG- Start codon.
Modified from Hirose and Sugiura, 2004.
22
2.3 Thylakoid membrane isolation
In order to measure the photosynthetic complexes (PSII, PSI, Cyt-bf) in vitro, thylakoids were
isolated from intact leaves number 3, 4 and 5 (from the top). All isolations and centrifugations
were made at 4°C according to Schöttler et al, (2004). In Table 1, the composition of the
extraction media is described.
The chlorophyll content was determined according to Porra et al, (1989) in 80% (v/v) acetone.
Table 1-Composition of thylakoid extraction media
Medium
Extraction medium
Shock medium
Double medium
Storage medium
Substances
Concentration [g/l]
Concentration [mmol/l]
HEPES
MgCl2·6 H2O
KCl
Sorbitol
KOH- Solution
HEPES
MgCl2·6 H2O
KCl
KOH- Solution
HEPES
MgCl2·6 H2O
KCl
Sorbitol
KOH- Solution
HEPES
MgCl2·6 H2O
KCl
MnCl2
Sorbitol
KOH- Solution
9,60
1,02
2,24
63,80
11,92
1,02
2,24
11,92
1,02
2,24
127,44
11,90
1,00
6,00
0,10
60,10
-
50,0
5,0
30,0
350,0
pH=6,1
50,0
5,0
30,0
pH=7,6
50,0
5,0
30,0
700,0
pH=7,6
50,0
5,0
40,0
0,5
330,0
pH=7,8
2.4 Quantification of photosynthetic complexes
2.4.1 Quantification of PSII and Cytochrome-b6f complex
The contents of PSII and the Cyt-bf were determined from difference absorption signals of Cytb559 (PSII) and Cyt-f and b6 (Cyt-bf). Isolated thylakoids equivalent to 50 µg chlorophyll ml -1
were unstacked in a low salt medium containing 0,03
ß-DM (ß-Dodecylmaltosid) (Table
23
2), to improve the optical properties of the probe (Kirchhoff et al, 2002). Addition of 1 mM
Ferricyanide (+III) leads to oxidation of all cytochromes. The subsequent addition of ascorbate
results in the reduction of Cyt-f and the high-potential form of Cyt-b559 (Ascorbate-Ferricyanide
difference spectrum) and the addition of dithionit in the reduction of cyt-b6 and the low-potential
form of cyt-b559 (Dithionit-Ascorbate difference spectrum) (Table 3). At each redox potential,
absorption spectra were measured between 575 and 540 nm wavelength with a V-550
spectrophotometer (Jasco GmbH, Groß-Umstadt, Germany) equipped with a head-on
photomultiplier. The spectral bandwidth was 1 nm and the scanning speed was 100 nm min-1.
Difference absorption spectra were deconvoluted using reference spectra and difference
extinction coefficients as in Kirchhoff et al, (2002). The concentration of the cytochrome bf
complex was determined from chemically induced absorbance changes (Kirchhoff et al, 2002)
and the content in PSII was calculated from the sum of the difference absorption signals of the
high potential and low potential forms of Cyt-b559 (Lamkemeyer et al, 2006).
Table 2-Cytochrome medium composition
Medium
Cytochrom-Medium
Substance
Na-HEPES
KCl
Na2EDTA·2H2O
Cytochrom-Medium +
ß-DM
Concentration(g/l)
7,15
2,30
0,04
Concentration(mmol/l)
30
30
0,01
50ml Cyt-Medium + 15 mg ß-DM (0,03% (w/v))
Table 3-Redox potential of cytochromes and effect on components of ETC at pH 7,6
Chemical substance
Em [mV]:
Effect
Ferricyanide K4[Fe(CN)6]·H2O
+ 400
Oxidation of Cytochromes
Na-Ascorbate
+ 90
Reduction of Cyt-f and Cyt b559 HP
Dithionit Na2S2O4
- 800
Reduction of Cyt b6, Cyt b559 LP
and C550
24
2.4.2 Quantification of P700
The P700 contents were determined from light-induced difference absorption changes of P700.
Isolated thylakoids equivalent to 50 µg chlorophyll ml-1 were solubilised in 0.2% (w/v) ß-DM
medium (Table 4) and then an electron acceptor (100µM methylviologen) and electron donor (10
mM ascorbate) where added. P700 was oxidised by the application of a saturating light pulse
(2000 µE m-2 s-1 red light, 200 ms duration). The measurements were done using the Dual-PAM
instrument (Heinz Walz GmbH, Effeltrich, Germany). The amplitude of the DAS (Difference
Absorption Spectrum) was proportional to P700 content, which can be calculated using the
difference extinction coefficient = 6,0 cm2/µmol-1 according to Lambert-Beer at 830-870-nm
wavelength.
Table 4-PSI medium composition
Medium
PSI-Medium
Substance
Na-HEPES
MgCl2·6H2O
KCl
PSI-Medium
+ ß-DM
Concentration(g/l)
7,15
1,00
2,30
Concentration(mmol/l)
30
5
30
50ml PSI -Medium + 100 mg ß-DM (0,2% (w/v))
2.4.3 Quantification of PC/P700
The PC content was determined by in vivo difference absorption spectroscopy in the far-red
range of the spectrum, and then recalculated based on the absolute P700 quantification in
isolated thylakoids (2.4.2) as described in Schöttler et al, (2004). Light-induced absorption
changes at 830-870 nm wavelength (predominant contribution of P700) and at 870-950 nm
wavelength (predominant contribution of PC) were done on pre-illuminated leaves with fully
activated Calvin cycle, to avoid an acceptor-side limitation of PSI.
25
2.5 P700 and PC reduction kinetics
To determine the maximum difference absorption signals of PC and PSI, pre-illuminated leaves
were put in darkness for some seconds, to fully reduce both components, followed by 10 s
illumination with far-red light (715 nm wavelength), to selectively excite PSI. After 10 s, a
saturating pulse of red light was applied (5000 µE m-2 s-1, 200 ms duration), to completely
oxidize PC and PSI and reduce the PSII side of the electron transport chain. At the end of the
actinic light pulse, all light sources were switched off, and PC and PSI reduction kinetics were
determined (Schöttler et al, 2007).
2.6 77K fluorescence measurement
The 77K chlorophyll-a fluorescence emission spectrum was determined on isolated thylakoids
equivalent to 10 µg chlorophyll ml-1, using PSI medium without -DM, in a F6500 fluorometer
(Jasco GmbH, Groß-Umstadt, Germany). The sample was excited at 430 nm wavelength (10 nm
bandwidth). The emission spectra between 655 and 800 nm was recorded with a bandwidth of 1
nm. The scanning speed was of 200 nm min-1. Depending on the quality of the measured signals,
these were repeated up to ten times and averaged to reduce noise.
2.7 Fluorescence measurement
Chlorophyll-a fluorescence of intact leaves was determined at room temperature using a DualPAM-100 instrument (Heinz Walz GmbH, Effeltrich, Germany). After measurement of the
dark-adapted F0 and FV/FM, the actinic light was switched on, and light response curves of linear
electron flux (LEF) and of non-photochemical quenching (qN), as well as of the redox state of
the PSII-acceptor side (qL) were recorded. The light intensity varied between 0 and 2000 µE m2 -1
s , taking 60 seconds per light intensity in a total of 20 steps. Fluorescence parameters were
calculated according to Baker et al, (2007).
26
2.8 PMF measurements and ATP synthase activity
The electrochromic absorption shift (ECS) of carotenoids is proportional to the electrochemical
component (∆H) of the PMF across the thylakoid membrane and allows the in vivo measurement
of changes in the PMF across the thylakoid membrane (Kramer et al, 2003; Baker et al, 2007).
The
difference
absorption
signal
was
measured
using
a
KLAS-100
LED-array
spectrophotometer (Heinz Walz GmbH, Effeltrich, Germany), allowing the simultaneous
measurement of light-induced difference absorption signals at six pairs of wavelengths in the
visible range of the spectrum between 500 and 570 nm. The ECS was deconvoluted from signals
arising from zeaxanthin formation, scattering effects, the ―C550‖ pheophytin signal, and from
redox changes of the cytochromes f, b6, and b559, as described in Klughammer et al, 1991. The
deconvoluted signals were then normalized to the chlorophyll content of the measured leaf
section.
The maximum amplitude of the ECS (ECST) was used as a measure for the light-induced PMF
above the thylakoid membrane. Leaves were illuminated for 5 min prior to each measurement, to
allow photosynthesis to reach steady state. ECST was determined after illuminating the leaves
with saturating light (2100 µmol m-2 s-1), which then was interrupted by a short interval of
darkness (15 seconds), and the dark-interval relaxation kinetics of the ECS were measured. 15
seconds dark intervals were sufficient for complete relaxation of the PMF. PMF partitioning into
pH and
was determined by analysing the slow relaxation phase of the ECS between 1
seconds and 15 seconds of darkness as described by Takizawa et al, (2007). A typical ECS curve
is shown in Figure 6.
To determine the ATPase activity, the fast phase of the dark-interval relaxation kinetics (between
0 and 500 milliseconds after the end of actinic illumination) was fitted with a single exponential
decay function (Figure 6A). The reciprocal value of the halftime of the ECS decay, which
corresponds to the thylakoid conductivity (gH+), was used as a measure of ATPase activity, since
the fast ECS decay is exclusively attributable to proton efflux through the ATPase complex
(Baker et al, 2007; Takizawa et al, 2007).
27
Light
off
Light off
ECS ( A)
A
t ECS1/gH+
ECSt PMF
Light off
B Light
ECS(∆A)
ECS ( A)
ECSss 
ECSinv  pH
Figure 6-Typical dark-interval relaxation kinetics of the electrochromic shift (ECS). A rapid light–dark
transition from steady-state conditions leads to several phases of ECS decay that can be analyzed to give information
about the total PMF and its partitioning into pH and
. A) The total PMF above the thylakoid membrane is
determined from the maximum amplitude of the ECS signal after a light to dark transition (ECS t). The inverse of the
lifetime for ECS decay, ECS, reflects the conductivity of the thylakoid membrane to protons e.g. ATPase activity. B)
Over a longer time-scale, the quasi-stable state decays, reflecting the movements of counter-ions, which relax the
portion of the ECS signal caused by the pH component of PMF, termed ECSinv. ECS is measured as the absorption
changes occurring around 520 nm (∆A). Modified from Baker et al, 2007.
2.9 Gas exchange measurements
The leaf assimilation capacity was determined in a closed-cuvette system with a Clark-type
electrode (LD2, Hansatech Instruments, Norfolk, England). Leaves were dark-adapted for 15
min, to determine respiration rates, and subsequently illuminated until photosynthetic oxygen
evolution reached its steady state. The leaf discs, with area of 10 cm2, were measured in a CO2saturated gas mixture (5% CO2), to totally repress photorespiration. Saturating actinic light was
provided by a FL-460 (Heinz Walz GmbH). The chlorophyll content of the leaf discs was
28
determined according to Porra et al, (1989), and assimilation capacity was calculated on leaf area
and chlorophyll basis.
The assimilation rates were corrected for the dark respiration, assuming that respiratory
activities in the light and in darkness are comparable (Fernie et al, 2004).
2.10 Metabolite quantification
2.10.1 Ethanol extraction of soluble sugars
For the extraction of the soluble sugars the leaf material was harvested always at the same time
(12 am). After pulverizing the leaf material with a Retsch mill (Retsch MM301), 50 mg of frozen
leaf material were put in 2 ml tubes. Afterwards 500
L of 80% EtOH were added and the
sample was incubated for 10 minutes at 78 ºC, at 440 rpm in a heating-shaker (Eppendorf
thermomixer comfort), followed by 5 minutes of centrifugation (14000 rpm). The supernatant
was transferred to a new 2ml tube and put on ice. The remaining sediment was resuspended with
500 L of 50% EtOH and again incubated in 78 ºC for 10 minutes, followed by centrifugation
and transfer of the supernatant to a 2 ml tube. The resuspension with 50% EtOH was repeated
one more time, and the supernatants were united. These samples were kept at -20 ºC and used for
soluble sugars quantification the following day.
2.10.2 Starch extraction
Since starch is insoluble in water, it must be broken into glucose monomers by heating and
enzymatic digestion for quantification. In order to do this, the sediment resultant from the
ethanol extraction was dried overnight at 20ºC.
The next day, it was dissolved with 1 ml of H2O. The sample was then put in a 2 ml tube (with
screwing lid) and incubated for 3 hours at 120 ºC. Next the sample was put on ice and cooled
down, finally adding 200 µL of Enzyme mix (Table 5). The sample was incubated overnight at
37 ºC. The following day the sample was centrifuged for 30 minutes (20.000 g). The supernatant
was put in a new 2 mL tube and put on ice. The remaining pellet was resuspended with 500 µL
H2O, and then incubated for 3 hours at 120ºC and then 300 mL of Enzyme mix were added.
29
Then the sample incubated overnight at 37 ºC. The next day, the starch was hydrolyzed and
quantification was done through determination of glucose units. The starch quantification was
done according to Smith and Zeeman, (2006).
Table 5-Enzyme mix composition
Medium
Enzyme-mix
Substance
5 ml Sodium Acetate (100 mM; pH = 5,5)
200 µl Amyloglucosidase of Aspergilus niger, 140
U/ml, Roche, Mannheim)
4 µl α-Amylase (of Pig pancreas, 10.000 U/ml, Roche,
Mannheim)
80 µl G-6-P-DH: centrifuge (13400 rcf, 2 min, 4ºC) and
dissolve the pellet in the glucose mix
2.10.3 Metabolite quantification
The samples were put in a 96 wells microplate and measured in a Photometer (SpectraMAX,
Molecular Devices, Sunnyvale, USA). The method is based in the enzymatic oxidation of
activated glucose via G-6-P-DH (Glucose-6-Phospate Dehydrogenase) which is coupled to the
reduction of NADP+ (Jones, 1977). NADPH has an absorption maximum at 340 nm. Therefore
the metabolite content can be quantified by photometric endpoint determination through the law
of Lambert-Beer. Each of the wells contained 160 µL of Glucose mix (Table 6) and in parallel 4,
6, 8 and 10µL of sample were added in each well. First the baseline was measured for 3 minutes
until it reached steady-state. After this, this step was repeated by addition of Hexokinase,
Phosphoglucose-Isomerase (PGI) and Invertase (Inv) (Table 7), keeping in mind that the addition
of a new enzyme could be done only when the absorption was constant (steady-state). After
glucose quantification, Fructose-6-Phosphate was converted into Glucose-6-Phosphate, by
addition of PGI, and fully oxidized. After complete consumption of glucose and fructose, sucrose
was cleaved by invertase. A general overview of the reactions involved in the metabolite
quantification is presented in Figure 7.
30
Table 6-Glucose mix composition
Medium
Substance
15,5 ml HEPES-MgCl2 (0,1 M, 3mM MgCl2)
Glucose-mix
480 µl ATP (60mg/ml) (Roche Diagnostics,
Mannheim, Germany)
480 µl NADP (36mg/ml) (Roche Diagnostics,
Mannheim, Germany)
80 µl G6PDH: centrifuge(13400 rcf, 2 min, 4ºC) and
dissolve the pellet in the Glucose mix
Table 7- Enzymes preparation
Hexokinase- Centrifuge 120 µl and dissolve pellet in
Enzymes
200 µl HEPES (0,1M)
Phosphoglucose-Isomerase- Centrifuge 60 µl and
dissolve pellet in 200 µl HEPES (0,1M)
Invertase- Dissolve a bit in 200 µl HEPES (0,1M).
Solution must turn yellow
ATP
Glucose
Sucrose
Inverta se
Hexokina se
ATP
Fructose
ADP
ADP
NADP+
Glucose-6-Phosphate
NADPH
G-6-P-DH
a bsorption
6-Phosphogluconate
Phosphoglucose-isomera se
Fructose-6-Phosphate
Figure 7- Overview of the enzymatic reactions occurring in the metabolite quantification.
31
3. Results
3.1 Growth phenotype
Figure 8- Growth phenotype in WT tobacco (PH) and ATPase transformant lines. A-WT; B-MJR1; C-MJR2;
D-MJR3; E-MJR4; F-MJR5; G-MJR6; H-MJR7; I-MJR10; J-atpC1. The atpB lines, except for line MJR2, showed
no differences in the growth phenotype. The atpC line showed impaired growth, reduced leaf size, and massive
retardation of flowering.
In this work transformants with point mutations introduced into the 5’-UTR of the plastid
encoded atpB gene (2.2) were characterized. Previous in vitro translation assays revealed
changes in translation rates between 10 and 200% of the wild-type level (Hirose and Sugiura,
2004). The lines showed no unequivocal growth phenotype (Figure 8), except for the line MJR2
whose growth was extremely retarded (Figure 8C). Therefore seeds from this line could not be
obtained in time to measure it in this master thesis. For this reason this line will not be mentioned
any further in this work. Additionally to MJR2, also an atpC antisense line (atpC1), with reduced
atpC mRNA accumulation was analysed as a positive control, for which a strong repression of
32
the ATPase had already been established (Schöttler et al, 2009, under revision). It showed a
retarded growth and severely delayed flowering (Figure 8J).
3.2 Chlorophyll content
700
B
5
4.5
600
4
500
Chlorophyll a/b
Chlorophyll content [mg m-2]
A
400
300
200
3.5
3
2.5
2
1.5
1
100
0.5
0
0
1
WT
2
MJR1
3
4
MJR4
MJR3
5
MJR5
6
MJR6
7
8
9
atpC1
MJR7
MJR10
1
WT
2
MJR1
3
4
MJR4
MJR3
5
MJR5
6
MJR6
7
8
9
MJR7
atpC1
MJR10
Figure 9- Chlorophyll content (A) and chlorophyll a/b ratio (B) in WT tobacco (PH) and ATPase
transformant lines. The chlorophyll content was determined on a leaf area basis. Some atpB lines had slightly
increased chlorophyll content, although with no significant differences. The antisense line showed the highest
chlorophyll content. It is important to say that the atpC line belonged to a different cultivar, SNN, than the atpB
lines, PH. The SNN variety has a higher chlorophyll concentration than PH. The value of the chlorophyll a/b ratio
showed no differences between the WT and atpB mutants. The antisense line had the lowest chlorophyll a/b ratio.
The chlorophyll content was determined on a leaf area basis, according to Porra et al, (1989).
The obtained data showed that the chlorophyll content was slightly higher in lines MJR1 (440
mg m-2), MJR5 (413 mg m-2) and MJR10 (416 mg m-2) than the WT (369 mg m-2) (
Figure 9A). All other atpB lines did not show changes in the chlorophyll content compared to the
WT. The antisense line showed a higher chlorophyll content (519 mg m-2) in comparison with all
other lines, including WT, (Figure 9A). This is due to the fact that this line belonged to a
different tobacco cultivar-SNN. This variety is known for having higher chlorophyll content than
the Petit Havana variety (Schöttler et al, 2009, under revision).
The chlorophyll a/b ratio showed no differences between the WT and atpB mutants (Figure 9B),
with a constant value of about 4,2. Only the antisense line showed a significantly lower
chlorophyll a/b ratio (Figure 9B).
33
3.3 ATPase activity
B
ECST /chl [∆I/I.(mg chl)-1]
Half life time of PMF [ms]
A 70
60
50
40
30
20
10
90
80
70
60
50
40
30
20
10
0
0
WT 1 MJR12 MJR33
4
MJR4
MJR6
5
6 MJR7
7
MJR5
8
MJR10
atpC1
9
WT1
2
3
MJR3
MJR1
4
MJR4
5
MJR5
6
MJR6
7
MJR7
8
9
atpC1
MJR10
Figure 10- Relaxation halftime of PMF (A) and maximum proton motive force (B) in WT tobacco (PH) and
ATPase transformant lines. Except for the antisense line none of the ATPase mutants showed an alteration in the
ATPase activity in comparison with WT (A). The PMF reached a maximum in the antisense mutant and lines MJR4
and MJR6 had the highest PMF of the atpB mutants (B).
To check the impact of atpB mutations and atpC antisense repression on ATPase, its activity
was determined through the fast dark-relaxation kinetics of the maximum electrochromic
absorption shift (ECST), which is a measure of the PMF above the thylakoid membrane (Kramer
et al, 2003; Takizawa et al, 2007). During the short dark interval the relaxation of the PMF is
determined by the rate of proton efflux through the ATPase, e.g. thylakoid membrane
conductivity, and therefore represents a measure of the ATPase activity (Takizawa et al, 2007)
(2.8). The average relaxation halftime of the PMF did not vary between the WT and the atpB
transformants (Figure 10A). In the WT the fastest relaxation time was observed, 14 ms, and line
MJR1 showed the slowest relaxation time of all atpB transformant with 17 ms, although it was
not a significant difference (Figure 10A). In the antisense line the halftime was increased to 50
ms (Figure 10A), which is attributable to its 80% reduction in ATPase content (Schöttler et al,
2009, under revision).
The maximum PMF accumulation was determined and the results showed a significant
difference in comparison with WT. Lines MJR4 and MJR6 showed the higher PMF formation
above the thylakoid membrane. Strangely, although line MJR1 showed a slightly lower
conductivity in comparison with the other atpB transformants, it did not have the highest value of
PMF (Figure 10B). In the antisense line the thylakoid conductivity was strongly repressed
34
(Figure 10A) and as a consequence the PMF above the thylakoid membrane was highly
increased (Figure 10B).
Assimilation [µmol EP / mg *h]
A 1000
900
800
700
600
500
400
300
200
B
50
Assimilation [µmol O2 m-2 s-1]
3.4 Assimilation capacity
45
40
35
30
25
20
15
10
5
100
0
0
WT
1
2
MJR1
MJR3
3
MJR4
MJR5
4
5
MJR6
6
MJR7
7
MJR10
atpC1
8
9
WT
1
MJR1
MJR3
2
3
MJR4
4
MJR5
MJR6
MJR7
5
6
7
atpC1
MJR10
8
9
Figure 11-Assimilation capacity on a chlorophyll basis (A) and on leaf area (B) in WT tobacco (PH) and
ATPase transformant lines. The assimilation capacity was measured on both chlorophyll and leaf area basis. Both
measurements revealed a decrease in assimilation in some atpB lines, but the assimilation was most repressed in the
atpC antisense line.
To check the impact of atpB mutations and atpC antisense repression on assimilation capacity,
the assimilation rate was determined on chlorophyll basis and on leaf area basis. On a
chlorophyll basis the obtained data showed significant differences between the WT and some of
the atpB lines. Lines MJR1 (583 µmol EP/mg*h), MJR4 (587 µmol EP/mg*h), MJR5 (578 µmol
EP/mg*h) and MJR6 (620 µmol EP/mg*h) showed the lowest assimilation capacities of the atpB
transformants (Figure 11A), having a decrease of about 25% in comparison with the WT (750
µmol EP/mg *h). The atpC1 line showed the lowest assimilation rate (240 µmol EP/mg *h) with
a decrease of about 70% in comparison with the WT (Figure 11A). The results of assimilation
capacity per leaf area were not exactly comparable with the results per chlorophyll. Only line
MJR4 showed a significant assimilation reduction (about 23%), while in all other lines the
reduction was less apparent. The atpC1 line was the line that showed the most reduced
35
assimilation per leaf area, 15 µmol O2 m-2 s-1. The SNN WT has comparable assimilation
capacities as PH (Schöttler et al, 2009, under revision).
3.5 Assimilation capacity versus membrane conductivity
100
80
gH+
60
WT
MJR1
MJR3
MJR4
MJR5
MJR6
MJR7
MJR10
atpC1
40
20
0
0
200
400
600
800
1000
Assimilation [µmoles electron pairs (mg chl.h)-1 ]
Figure 12- Assimilation capacity versus conductivity of the membrane in WT tobacco (PH) and ATPase
transformant lines. The atpB lines did not show a linear correlation between the conductivity and the assimilation
capacity. In the antisense line the assimilation declined in parallel with ATP synthase activity repression, indicating
a limiting role of ATPase.
A plot of the reciprocal values of the conductivity of the thylakoid membrane against
assimilation capacities per chlorophyll was made (Figure 12). Unfortunately as all data points for
the atpB mutants scattered closely together, due to minor differences in assimilation and gH+, no
conclusions on a correlation between ATPase activity and assimilation capacity could be drawn.
However, taking the atpC antisense line into account, the overall data showed that a lower
conductivity closely correlates with reduced assimilation capacity (Schöttler et al 2009, under
revision), which might suggest that ATPase activity, can limit leaf assimilation.
36
3.6 Organization of the electron transport chain
A
4
B
1.6
Cyt-bf [mmol/mol chl.]
PSII [mmol/mol chl.]
3.5
3
2.5
2
1.5
1
0.5
1.2
1
0.8
0.6
0.4
0
1
WT
2
MJR1
3
MJR3
4
MJR4
5
MJR5
6
MJR6
7
MJR7
8
MJR10
1
WT
9
atpC1
12
D
2
MJR1
3
MJR3
4
MJR4
5
MJR5
6
MJR6
7
M JR7
8
MJR10
9
atpC1
3
2.5
PSI [mmol/mol chl.]
10
PC [mmol/mol chl.]
1.4
0.2
0
C
1.8
8
6
4
2
1.5
1
0.5
2
0
0
WT1
MJR1
2
MJR3
3
MJR4
4
MJR5
5
MJR6
6
MJR7
7
MJR10
8
atpC1
9
WT
1
MJR1
2
3
MJR3
MJR4
4
MJR5
5
6
MJR6
MJR7
7
MJR10
8
atpC1
9
Figure 13- Concentration of the components of the ETC in WT tobacco (PH) and ATPase transformant lines.
A-PSII; B-Cyt-bf; C-PC; D-PSI. The photosystems composition was unaltered between WT and ATPase
transformant lines. The Cyt-bf content also did not vary between WT and mutants. The PC content did not change
between the WT and the mutants except for the antisense line who had a higher PC content.
To see how the other components of the electron transport chain are affected by the atpB
mutations, the composition of the ETC was determined by difference absorption spectroscopy
(Figure 13). It was observed that neither PSII nor PSI contents differed significantly between
wild type and the atpB transformants (Figure 13A and 13D). The PSII content reached values of
3 mmol/mol chl, and its function seemed not to be affected in the atpB transformant lines, as
indicated by the unaltered maximum quantum efficiency of PSII (FV/FM) in comparison with the
WT (Figure 14). The PSI contents varied around 2,4 mmol/mol chl in WT and all mutants, which
is in good agreement with the literature (Schöttler et al, 2007). The atpC line showed a clear
reduction in PSI content, which concurs with its reduced chlorophyll a/b ratio (Figure 9B). The
Cyt-bf content was overall unchanged in the atpB mutants as well as the antisense mutant, with
37
an average value of 1,4 mmol/mol chl (Figure 13B). The PC contents showed little differences,
which were not significant, except for the atpC line (Figure 13C). In this line the PC content was
20% higher than in the WT (Figure 13C).
3.7 Maximum PSII efficiency
0.82
0.8
0.78
Fv / Fm
0.76
0.74
0.72
0.7
0.68
0.66
0.64
0.62
WT1
MJR1
2
MJR3
3 MJR4
4 MJR55
MJR6
6 MJR7
7
MJR10
8
atpC1
9
Figure 14- Maximum quantum efficiency in WT tobacco (PH) and ATPase transformant lines. The Fv/Fm ratio
values of dark-adapted leaves showed no alterations between the WT and all atpB lines. Only the atpC line showed
a reduction in this ratio.
The Fv/Fm ratio is a measure of the intrinsic (or maximum) efficiency of PSII e.g. the quantum
efficiency if all PSII centers were open (Baker, 2008). The Fv/Fm ratio values of dark-adapted
leaves showed no alterations between the WT and all atpB lines (Figure 14). In all these lines the
average value was 0,78. The unalteration of this value indicates that the PSII function was not
affected in the atpB lines. In the atpC line the ratio was of 0,72 which was in accordance with
previous data (Schöttler et al, 2009, under revision).
38
3.8 77K chlorophyll-a fluorescence emission spectra
1.8
WT
MJR1
MJR3
MJR4
MJR5
MJR6
MJR7
MJR10
atpC1
Chlorophyll-a fluorescence
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
660
680
700
720
740
760
780
800
Wavelenght [nm]
Figure 15- 77K chlorophyll-a fluorescence emission spectra in WT tobacco (PH) and ATPase transformant
lines. For better comparability the PSII peak was normalized to 1 in all lines. The 77K chlorophyll-a fluorescence
emission spectra showed no differences between the wild type and all mutant lines.
The two peaks at 686 nm and 733 nm reflect the fluorescence of PSII and PSI, respectively. The
absence of a shift in the wavelength in all mutants, in comparison with WT, indicates that there
were no free light harvesting complexes. The ratio of the maximum emission of PSII and PSI
gives information about the ratio of the two photosystems. The fact that the 77K chlorophyll-a
fluorescence emission spectra did not change between the WT and all atpB mutants (Figure 15)
indicates and is in accordance with the unaltered contents of both photosystems (Figure 13). In
the antisense line, although there was a small reduction in PSI content (Figure 13) the difference
in the chl-a fluorescence emission spectra was not significant (Figure 15).
39
3.9 PMF partitioning
A
1
B
0.7
0.9
0.6
0.8
0.5
0.7
∆Ψ
∆pH
0.6
0.5
0.4
0.3
0.4
0.3
0.2
0.2
0.1
0.1
0
0
1
WT
2
MJR1
3
4
MJR4
MJR3
MJR5
5
6
MJR6
7
MJR7
M JR10
8
9
atpC1
1
WT
2
MJR1
3
MJR3
4
MJR4
5
MJR5
6
7
MJR6
MJR7
9
M 8JR10 atpC1
Figure 16- PMF partitioning into ∆pH (A) and ∆Ψ (B) in WT tobacco (PH) and ATPase transformant lines.
All lines, except MJR1, showed a tendency to favour the accumulation of the osmotic component ( p ) of the PMF.
The partitioning of PMF into its ∆pH and ∆Ψ components was determined from the proportion of
the slow relaxation kinetic of the ECS attributable to counter ion movements above the thylakoid
membrane (ECSinv), relative to ECST (Baker et al, 2007). In all atpB lines, except for line MJR1,
on average 65% of the PMF was stored as ∆pH (Figure 16A and 16B), although the total PMF
was increased in the atpC antisense line, the fraction of PMF stored as ∆pH was almost the same
as in all atpB lines. As only the ∆pH component is relevant for triggering the photoprotective
mechanisms and initiating photosynthetic control (Takizawa et al, 2007), we next checked for
changes in photoprotection.
40
3.10 Light response curves
A
ETR PSII [µmol electrons m-2 s-1]
200
150
100
50
0
0
1000
2000
3000
4000
Light intensity [µE m-2 s-1]
B
1.0
0.8
qN
0.6
0.4
WT
MJR1
MJR3
MJR4
MJR5
MJR6
MRJ7
MJR10
atpC1
0.2
0.0
0
1000
2000
3000
C
WT
MJR1
MJR3
MJR4
MJR5
MJR6
MRJ7
MJR10
atpC1
1.0
0.8
qL
4000
Light intensity [µE m-2 s-1]
0.6
0.4
0.2
0.0
1000
2000
3000
4000
Light intensity [µE m-2s-1]
Figure 17- Light response curves of WT tobacco and transformant lines. A-ETR of PSII; B- Thermal
dissipation of excitation energy-qN; C-Redox state of the PSII acceptor side-qL. These light response curves
were calculated as described in 2.7. The ETR of the atpB lines was reduced in comparison with the WT but not as
much as the atpC line. The qN curve showed a rapid induction of NPQ in the antisense line, being much more
delayed in the atpB mutants. Also in the atpC line the PSII acceptor side was more reduced than the atpB mutants
and the WT.
41
The acidification of the lumen alters both the efficiency of linear electron flux and PSII antenna
function. Therefore the light saturation curve of linear electron flux and the light response curves
of the chlorophyll fluorescence parameters qN and qL, which are measures for the thermal
dissipation of excitation energy and for the redox state of the PSII acceptor side, respectively
(Krause and Weis, 1991; Baker et al, 2007), were checked. The saturation curve of linear
electron flux, as calculated from the PSII quantum yield, showed that in agreement with the
slightly lower assimilation rates (Figure 11A), lines MJR1, MJR4 and MJR5 also had a slightly
reduced linear electron transport rates per leaf area under almost all light intensities, when
compared with WT (Figure 17A). Both WT and atpB mutants reached their maximum electron
transport rate (ETR) at 600 µE. While at this light intensity the WT had an ETR of 80 µmol
electrons m-2 s-1 the atpB lines had an average ETR of 65 µmol electrons m-2 s-1. In the antisense
line, in agreement with the lowest assimilation rate determined by means of gas exchange
measurements (Figure 11A), also linear electron transport rates per leaf area was severely
repressed being saturated at already 300 µE with only 25 µmol electrons m-2 s-1 (Figure 17A).
The qN curves showed that in lines MJR1, MJR4 and MJR5, there was a slight shift in the onset
of non-photochemical quenching qN (Figure 17B), is slightly more rapid than in WT. In the
antisense line the rise of qN was strongly accelerated, reaching its maximum already at 300 µE
light intensity, resulting in increased thermal dissipation of excitation energy and reduced
quantum efficiency of photosynthesis (Figure 17B).
Additionally also the qL curve confirmed an impairment of the LEF. The results showed that at
low light intensities the PSII acceptor side of lines MJR1, MJR4 and MJR5 was slightly more
reduced in comparison with the WT (Fig.16C). In the antisense line, as indicated by the much
lower qL values (Figure 17C), the PSII acceptor side was much more reduced, which suggested a
strong impairment of LEF.
42
3.11 Reduction kinetics of Cytochrome-f and P700
B
1.0
1.0
0.8
0.8
P700 (normalized)
Cytochrome-f (normalized)
A
0.6
0.4
0.6
0.4
0.2
0.2
0.0
0.0
0
20
40
60
80
100
120
140
WT
MJR1
MJR3
MJR4
MJR5
MJR6
MJR7
MJR10
atpC1
0
20
40
Time [ms]
60
80
100
120
140
Time [ms]
Figure 18- Reduction kinetics of Cytochrome-f (A) and P700 (B) in WT tobacco and ATPase transformants.
For better comparability, the fully oxidised states of Cyt-f and P700 were normalized to one, and the completely
reduced states were normalized to zero. The reduction kinetics of the atpB lines and WT were indistinguishable for
both Cyt-bf and P700. In the antisense line the reduction kinetics were much slower in both cases.
To confirm that the LEF between the photosystems was restricted, the reduction kinetics of Cyt-f
and P700 after the end of a saturating light pulse were measured.
The data showed that there were no significant changes in the relaxation time of P700 reduction
between the MJR transformants, with the highest value corresponding to 8 ms (MJR1) and the
lowest to 6 ms (WT) (Figure 18B). The relaxation time of the antisense line was increased to 15
ms which concurs with its decreased assimilation rate (Figure 11A).
The halftime of Cyt-f reduction kinetics was also measured and the results showed that the atpB
transformants had no significant differences among them, with the relaxation times varying
between 11 ms and 13 ms. Once again the antisense line had the slowest relaxation time, 40 ms,
Figure 18A) which is in line with the observed decrease of leaf assimilation rate (Figure 11A).
The increase in the halftime of Cyt-f reduction together with the more reduced state of the PSII
acceptor side (Figure 17C) and slower reduction kinetics of PSI (Figure 18B) suggests a
restriction of LEF.
43
3.12 Metabolite quantification
450
140
B
400
120
Starch (µmol GLU/g FW)
Total soluble Sugars (µg Sugar /g FW)
A
100
80
60
40
350
300
250
200
150
100
20
50
0
0
WT1
2
MJR1
3
MJR3
MJR4
4
MJR5
5
MJR6
6
MJR7
7
MJR10
8
atpC1
9
1
WT
2
MJR1
3
MJR3
4
MJR4
5
MJR5
6
MJR6
7
MJR7
8
MJR10
9
atpC1
Figure 19- Total soluble sugars concentration (glucose+fructose+sucrose) (A) and starch concentration (B) in
WT tobacco (PH) and ATPase transformant lines. The accumulation of total soluble sugars was significantly
different between WT and mutants. Lines MJR1, MJR4, MJR5 and the atpC line had the highest accumulation of
total soluble sugars (A). The starch quantification also showed significant differences between WT and
transformants. Lines MJR1, MJR4, MJR5 presented the lowest starch content of all atpB mutants. The atpC line
showed the lowest starch accumulation of all transformant lines (B).
Starch is the main form in which plants store carbohydrates, and is of importance in the carbon
economy of many organs, tissues and cell types in the plant. In leaves the flux of carbon into the
starch-biosynthetic pathway is thought to be controlled by modulation of ADP-Glucose
Pyrophosphorylase (AGPase) activity (Smith and Zeeman, 2006). The reaction catalyzed by
AGPase is very sensitive to ATP availability and inhibited by phosphate (Zeeman et al, 2007).
Therefore the starch content can reveal very subtle changes in ATP availability. Since the
production of ATP is dependent on ATPase activity, the starch quantification indirectly shows
the accumulation of ATPase.
The quantification of total soluble sugars showed that lines MJR1, MJR5 and MJR4 had the
highest amounts of total soluble sugars of the atpB mutants, with an average of 90 µmol sugar/g
FW (Figure19A). The value of total sugar accumulation in atpC1 was comparable to the lines
mentioned previously (Figure 19A). The WT and line MJR6 had the lowest sugar accumulation,
with an average of 60 µmol sugar/g FW.
The starch content was maximal in the WT and line MJR7, with an average of 320 µmol GLU/g
FW (Figure 19B). Lines MJR1, MJR4 and MJR5 had a reduction of 20% in starch accumulation
44
compared with WT. The antisense line had the lowest starch concentration (Figure 19B) with a
reduction of 66% compared with WT, well in line with the strongly reduced assimilation in this
line (Figure 11A).
It was observed a tendency for the lines that had highest total sugar contents to accumulate less
starch (Figure 19).
45
4. Discussion
The components of the photosynthetic electron transport chain closely adjust to changes in the
metabolic demand for ATP and NADPH, so that their production exactly matches their
consumption by the Calvin cycle, as otherwise ROS might be generated leading eventually to
cell death (Kim et al, 2008; Laloi et al, 2007). It was previously shown that the content of Cytbf, PC and ATPase decreases linearly with the leaf development and assimilation capacity
(Anderson et al, 1988; Schöttler et al, 2004; Schöttler et al, 2007).
The fact that the Cyt-bf complex catalyzes the rate-limiting reaction of linear electron flux, and
therefore adjusts the photosynthetic light reactions to the metabolic demand is well established
(Anderson, 1992; Price et al, 1995; Price et al, 1998). PC has also been suggested to contribute
to electron flux control (Kirchhoff et al, 2004; Schöttler et al, 2004). Therefore the close
correlation between the contents of these components and the assimilation capacity was
unsurprising. However, the physiological relevance of the additional adjustment of ATPase
(Anderson et al, 1988; Schöttler et al, 2007) was not fully understood. Therefore, the objective
of this work was to determine if the ATPase might also control photosynthetic activity and
function as an additional point of flux control.
In order to do this, the accumulation of the essential plastome-encoded
manipulated, since together with the nuclear encoded
subunit was
subunit, it has been shown to be rate-
limiting for ATPase assembly in Chlamydomonas (Drapier et al, 2007). As a positive control
previously generated tobacco plants with an antisense construct directed against the atpC gene
encoding the subunit were used.
No significant differences in the chlorophyll accumulation, chlorophyll a/b ratio and in the
accumulation of any component of the ETC were seen. Therefore any changes in physiological
parameters in atpC (as well as atpB) should be attributable to alterations in ATPase function, as
will be discussed in detail later.
46
ATPase content requires a threshold reduction to obtain growth phenotype
The atpB transformants (with exception for line MJR2) showed no visible phenotype (Figure 8),
but the atpC mutant had a retarded growth which was due to the more than 80% reduction in
ATPase content (Schöttler et al, 2009, under revision). The absence of a phenotype in all atpB
lines (except MJR2) might be explained if we consider that previous work, on subunit mutants,
showed that changes in ATPase contents of less than 40% had no clear effects on growth,
assimilation and ATPase activity (Schöttler et al, 2009, under revision). Taking this into account
we can assume that the ATPase contents in the atpB mutants might be above a certain threshold
necessary for a mutant phenotype to be visible.
ATPase activity is subjected to post-translational regulation
Since there was no visible phenotype and the assimilation rates were not as repressed as in the
atpC line (Figure 11A), the in vivo activity of the ATPase was determined. For this purpose, the
kinetics of PMF relaxation was measured during short intervals of darkness. The reciprocal value
of the halftime of PMF relaxation directly reflects ATPase activity, as in the dark the proton
efflux rates from the lumen are exclusively determined by the rate of ATP synthesis (Baker et al,
2007; Takizawa et al, 2007). The results showed that, except for the atpC line, the conductivity
of the membrane, e.g. ATPase activity, was not significantly altered in any other line in
comparison with WT (Figure 10A). It is known that the ATPase activity is not exclusively
controlled by the enzyme content but is rather highly regulated (1.1.4). Therefore the fact that in
these lines there was no change in the membrane conductivity might be explained by two
scenarios: Either ATPase contents might be unaltered (despite of the introduced mutation in the
5’-UTR); or the ATPase from the mutants was working more efficiently than the one from WT.
It is known that in the WT, not all ATPases operate with maximum activity, either because a
subpopulation of enzymes is not active at all, or because the activity of a large number of
ATPases is partially down-regulated (Schöttler et al, 2009, under revision). As other
transplastomic plants bearing similar mutations in the 5’-UTR as the atpB mutants showed
reductions in the contents of the targeted proteins we favor a scenario where relatively small
47
changes in ATPase amounts in the atpB transformants are compensated by some kind of posttranslational regulation. However, to get to a definitive conclusion, the accumulation of ATPase
needs to be ultimately determined by immunoblot analysis (See Outlook).
ATPase activity limits leaf assimilation
In the atpC line the lower membrane conductivity resulted in a higher PMF accumulation above
the membrane, suggesting that ATPase was restricting proton flow out of the lumen and leading
to a stronger lumen acidification. Additionally, in this line the lower assimilation capacity
correlated with a much slower relaxation of the PMF. As the accumulation of the other
photosynthetic components was unaltered, the reduced ATPase accumulation should cause the
reduced assimilation. A plot of the conductivity of the membrane versus assimilation rates
showed no clear correlation between the conductivity of the membrane and the assimilation
capacity in the atpB lines (Figure 12), probably because the level of ATPase reduction was not
sufficient to reduce the membrane’s conductivity. On the other hand, the atpC line had already
shown a correlation between these two factors (Schöttler et al, 2009, under revision) (Figure 12).
This confirmed the suggestion that ATPase activity was limiting leaf assimilation. Taking this
into account, the activity of the ATPase might be considered as a limiting factor of plant growth.
This observation is in agreement with data from Wu et al, (2007), who characterized an
Arabidopsis mutant with a point mutation within the atpC gene, resulting in a shift of the redox
potential of the cysteine bridge of the
subunit. This led to an impairment of the reductive
activation of the enzyme and resulted in reduced growth rates. However, Wu et al, (2007) did not
determine the effect of reduced ATPase activation on linear flux rates and assimilation; therefore
they did not establish a strict correlation between these factors. But in principle, the strict
correlation between enzyme activity on the one hand and assimilation capacity and growth on the
other hand, indicates a limiting effect of ATPase repression on photosynthetic electron transport.
48
Osmotic component of the PMF is preferentially accumulated
The partitioning of the PMF revealed that in all plants (WT, atpB lines and antisense line) there
was a tendency to favour the osmotic component ( p ) (Figure 16A). This occurs because both
∆pH and ∆Ψ can drive ATP synthesis but only the osmotic component is relevant for triggering
the photoprotective mechanisms and initiating photosynthetic control (Takizawa et al, 2007).
Although it is well established that the partitioning of the PMF can vary strongly between
different plant species (Takizawa et al, 2007), the mechanisms underlying this different
partitioning of PMF are still not well understood.
In principle PMF partitioning into
pH and ∆Ψ is determined by the extent of counter-ion
movements. Therefore the PMF partitioning could be regulated by the expression of putative ion
channels that are yet to identify (Marmagne et al, 2007).
ATP synthase restriction leads to faster lumen acidification and repression of PQ reoxidation
The reduced assimilation capacity of the atpC antisense line might be either explained by a
limited ATP availability for the Calvin Cycle, or by more complex feedback effects of the
reduced ATPase activity on LET, so that both ATP and NADPH production are reduced. We
favour the second scenario: As a consequence of the much strongly reduced ATPase activity in
the atpC line, the linear electron transport (Figure 17A) is strongly repressed. This is due to the
increased PMF above the thylakoid membrane (Figure 10B), resulting in the extremely rapid
activation of the NPQ already at low light intensities (Figure 17B). As a consequence, the
quantum efficiency of CO2 fixation should be strongly reduced, which might contribute to the
retarded growth of this line (Figure 8J). The more reduced steady-state redox poise of the PSII
acceptor side and the PQ pool, as deviated from the light response curve of qL (Figure 17C) is
also in line with a restricted linear electron flux. As also the reduction of Cyt-f and P700 (Figure
18), after the end of a saturating light pulse, is delayed, this indicates a repression of plastoquinol
(PQ) reoxidation. Ultimately this inhibition of linear electron flux by the increase of lumen
acidification, due to restricted PQ reoxidation, e.g. ―photosynthetic control‖ is probably the
major contributing factor to the growth phenotype of the atpC antisense line. In wild-type plants,
49
PQ reoxidation is the rate-limiting step of LEF (Anderson, 1992; Price et al, 1998), but when the
thylakoid lumen pH-value drops below 6,5, it is further slowed down, as PQ oxidation works
against a much higher PMF (Kramer et al, 1999). Therefore, the Calvin cycle is being limited by
the provision of ATP and NADPH. Some atpB transformants showed a slightly reduced electron
transport rate in addition to a slightly faster NPQ activation (Figure 17A), in comparison with the
WT. This is in line with slightly higher PMF and might indicate that ATPase activity in these
mutants was somewhat reduced. It is known that below a lumen acidification of pH = 6,5,
―photosynthetic control‖ increasingly slows down proton translocation into the lumen, so that
proton influx by linear electron flow and efflux through the ATPase are re-balanced to prevent
destruction due to lumen over-acidification (Takizawa et al, 2007). The induction of
photoprotective mechanisms such as PsbS-protein and xanthophyll-cycle is also believed to be
initiated at pH-values of 6,5 (Takizawa et al, 2007). Taking this into consideration, we may
conclude that minor changes in the steady state lumen acidification may exist in some atpB
mutants as well, so that down regulation of the LEF and induction of NPQ occurs at slightly
lower light intensities than in the WT. The atpB mutants also show a slightly more reduced redox
state of the PSII acceptor side, which indicate that the more restricted LEF (Figure 17A) is
probably due to a down regulation of the Cyt-bf activity (Anderson, 1992; Price et al, 1998). The
reduction kinetics of Cyt-f and P700 (Figure 18) was not significantly different from the WT and
the slightly more reduced steady-state redox poise of the PSII acceptor side and the PQ pool, in
lines MJR1, MJR4, and MJR5, indicates only a slight limitation of Calvin cycle due to ATP and
NADPH deprivation.
ATP synthase reduction does not affect the other components of the photosynthetic apparatus
Despite the increased steady-state PMF and a much stronger thylakoid lumen acidification in the
atpC antisense mutant, no detrimental effects were seen on the photosynthetic apparatus itself. In
vitro, both PC and the oxygen-evolving complex of PSII have been shown to suffer from great
damage at pH-values below 5.5 (Kramer et al, 1999). In the atpC mutants neither PSII nor PC
contents were decreased (Figure 13A and 13C) and also PSII function was not impaired, as
indicated by the basically unaltered FV/FM values (Figure 14) indicating that the pH probably did
50
not drop to a critical level. For this reason it is not surprising that, among the atpB transformants,
the composition of ETC was also unaltered (Figure 13) since the reduction of ATPase was
probably much less pronounced in these lines compared with the antisense line. In the atpB lines
the PSII function was also not impaired, as indicated by unaltered FV/FM values (Figure 14).
Starch accumulation is impaired by ATP restriction
In spite of an absence of a growth phenotype and strong overall physiological effects in the atpB
lines some subtle differences were seen in terms of ATP accumulation. Starch is the main form
in which plants store carbohydrate, and is of importance in the carbon economy of many organs,
tissues and cell types in the plant. In leaves the flux of carbon into the starch-biosynthetic
pathway is thought to be controlled by modulation of AGPase activity and is very sensitive to
ATP and free Pi availability (Zeeman et al, 2007). The atpB lines had a lower accumulation of
starch than the WT plants, although not as low as the antisense line (Figure 19B), suggesting that
the ATP availability is in fact lower.
The AGPase activity is allosterically regulated by the levels of Pi, an inhibitor, and 3phosphoglycerate (PGA), an activator. The ratio of these two metabolites changes according to
supply of photo-assimilates and the demand for them (Zeeman et al, 2007). Under conditions
where supply exceeds demand, high levels of photo-assimilates translate into a high 3-PGA/Pi
ratio in the chloroplast stroma, the activation of AGPase and the synthesis of starch. When the
demand is higher than the supply, the 3-PGA/Pi ratio is lower and the AGPase is inhibited which
translates in a lower starch synthesis. If atpB lines with lower starch concentration produced less
ATP resulting in a higher level of free phosphate and higher inhibition of AGPase this could
explain the different starch accumulation (Zeeman et al, 2007). In case there is an inhibition of
AGPase, less starch will form and possibly the soluble sugars, which are formed previously in
the metabolic pathway (Zeeman et al, 2007) will accumulate, explaining why the lines that had
lower starch content had a higher total sugar concentration (Figure 19A).
51
In vitro data are not transferable to in vivo situation
If we consider all the obtained data, some atpB lines (MJR1, MJR4 and MJR5) showed small but
significant differences from the WT (total PMF,ETR,NPQ), although the differences were not as
obvious as in the atpC antisense line. The expected alteration of ATPase content was based on
previous in vitro data that revealed changes in translation rates between 10 and 200% of the WT
level (Hirose and Sugiura, 2004). Although the content of ATPase in these atpB transformants
must still be verified, one thing is certain: the obtained changes are probably not as pronounced
as suggested in the work by Hirose and Sugiura, (2004). This is probably due to the fact that in
vitro data are not always transferable to the in planta situation, e.g. due to the existence of
specific translation initiation factors, which might bind to other parts of the 5’-UTR.
Alternatively, the processes of CFl assembly in Nicotiana tabacum might be different from the
complex assembly in Chlamydomonas (Drapier et al, 2007), in which translation of the
subunit, although essential, may not be limiting for assembly unless it is dramatically repressed
(as in MJR2). Instead the ATPase biogenesis might be controlled only by the availability of the
nuclear-encoded atpC.
ATP synthase repression restricts plant growth by ―photosynthetic control‖
Although the atpB mutants do not allow the extraction of many conclusions about the effect of
reduced ATPase content on photosynthetic electron transport, the data obtained with the
antisense line indicate that, in addition to the Cyt-bf and PC, ATPase is a major control point of
the photosynthetic electron flux. The regulation of electron flux via down-regulation of ATPase
content, could be especially important during leaf ageing (Schöttler et al, 2007) and in response
to drought stress (Kohzuma et al, 2009), which both result in a strong repression of ATPase in
parallel with the Cyt-bf. The best advantage of an ATPase repression is that the initiation of
photoprotective mechanisms is shifted to lower light intensities, since a lower proton influx rate
is sufficient to acidify the lumen and to trigger qN. In case of a selective reduction of the Cyt-bf,
without similar down-regulation of the ATPase, this might not be the case, since only the
capacity for proton influx into the lumen would be reduced, while proton efflux through the
ATPase would stay high, resulting in a lower steady-state PMF and probably a compromised
52
induction of the photoprotective mechanisms. For this reason, the ATPase seems to be a very
important factor for the adjustment of the photosynthetic light reactions to the metabolic
demands.
5. Outlook
Since the atpB mutants did not show the expected massive repression of ATPase contents, we
can imagine two scenarios: Either the in vitro data are not transferrable to the in vivo situation
e.g. due to the existence of specific translation initiation factors, which might bind to other parts
of the 5’-UTR, or the
subunit is not limiting for the assembly of ATPase unless it is
dramatically repressed.
To determine the obtained changes in translation rates of the atpB mutants polysome analysis
must be performed. Furthermore, the actual ATPase contents need to be determined by
immunoblots. In case the immunoblots reveal altered ATPase content, we must try to unveil
which kind of mechanism alters ATPase activity on a post translational level: If they are due to
phosphorylation (1.1.4.4), which will be checked by using anti-phosphothreonine and antiphosphoserine antibodies and also through proteomics; or if there is dimerization of the enzyme,
as suggested by Schwaßman et al, (2007). In this case Blue native-Electrophoresis should be
performed.
53
6. Abbreviations
ADP- Adenosine Diphosphate
AGPase- ADP-Glucose Pyrophosphorylase
ATP- Adenosine Triphosphate
ATPase- ATP synthase
ß-DM- ß Dodecylmaltosid
CES- Control by Epistasy of Synthesis
CF0- chloroplastidial F0- complex
CF1- chloroplastidial F1- complex
CO2- Carbon Dioxide
Cyt-b6f- Cytochrome b6f Complex
Cyt-f- Cytochrome f
DAS-Difference absorption spectrum
EP- Electron pairs
ETC- Electron Transport Chain
EtOH- Ethanol
ETR- Electron Transport Rate
FW- Fresh weight
G-6-P-DH- Glucose-6-Phospate Dehydrogenase
GLU- Glucose
H20- Water
Inv-Invertase
KCl- Potassium Chloride
LEF- Linear electron flux
LHC- Light harvesting complex
MgCl2- Magnesium Chloride
54
Ms- Milliseconds
NaAc- Sodium Acetate
Na-Ascorbate- Sodium Ascorbate
NADP+- Reduced Nicotine Adenine Dinucleotide Phosphate
NPQ-Non photochemical quenching
P700- chlorophyll a dimer from photosystem I reaction-center.
PC- Plastocyanin
PGA- Phosphoglycerate
PGI- Phosphoglucose-Isomerase
PH- Petit Havana (tobacco cultivar)
Pi- Inorganic Phosphate
PMF- Proton motive force
PQ- Plastoquinol
PSI- Photosystem I
PSII- Photosystem II
RCF- relative centrifugal force
ROS- Reactive oxygen species
RPM- Rotations per minute
SNN- Sansun (tobacco cultivar)
UTR- Untranslated region
WT- Wild Type
55
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8. Acknowledgments
Firstly, I would like to thank Dr. Mark Aurel Schöttler for the opportunity to do this work in his
group, for his help, his advice and discussions.
I would also like to thank Dr. Jorge Marques da Silva for his supervision, for his help, friendly
support and advice.
A very special thanks to Wolfram Thiele for all his help in the lab and for making the basement a
much brighter place.
A very big thanks to Markus Rott for generating the transplastomic plants.
I would like to thank Dr. Claudia Flügel and Dr. Yinghong Lu for reviewing this thesis and all
their advice.
A special thanks to Martin Ballaschk and Henrik Zauber, for making lunch time the most wanted
time of the day.
I thank Sebastian Hasdorf for his help in the lab and with computers.
I am very grateful to the whole AG Schöttler and AG Bock for making me overcome any kind of
problem in the lab and also for the nice working atmosphere.
I would also like to thank my parents and my brother for their unconditional love and support.
60