CHARLES UNIVERSITY IN PRAGUE, FACULTY OF SCIENCE
Department of Biochemistry
REGULATION OF CYTOKININ METABOLISM IN TOBACCO
PLANTS AND CHLOROPLASTS
Ph.D. THESIS
MARIE HAVLOVÁ
Supervisor: RNDr. Radomíra Vaňková, CSc.
Prague 2011
Declaration
I hereby declare that I completed this PhD. thesis independently, except where explicitly
indicated otherwise. It documents my own work, carried out under the supervision of RNDr.
Radomíra Vaňková, CSc. Throughout, I have properly acknowledged and cited all sources used.
Neither this thesis nor its substantial part have been submitted to obtain this or other academic
degree.
Prague................................
...........................................
Marie Havlová
Declaration of co-authors
I hereby declare that Marie Havlová/Nováková has substantially contributed to all 4 articles
which represent an integral part of this Ph.D. thesis. She performed most of the experiments,
especially in the papers where she is the first author and she actively participated in the set-up of the
experiments, in the interpretation of the results and in the preparation of the manuscripts.
Prague................................
...........................................
RNDr. Radomíra Vaňková, CSc.
Aknowledgement:
I am heartily thankful to my supervisor, RNDr. Radomíra Vaňková, CSc., without whose
encouragement, guidance and support from the initial to the final level I would not have been able
to successfully complete my studies.
I would like to thank all my dear collegues who helped me with particular experiments and
stimulated to my further work, namely RNDr. Alena Gaudinová and Eva Kobzová.
Special thanks belongs to ing. Petre I. Dobrev, CSc. for HPLC and to ing. Jiří Malbeck, CSc. for
mass spectroscopy analysis.
I also would like to make a special reference to Prof. RNDr. Danuše Sofrová, CSc., who was
always so kind and prepared to give me an advice.
Finally, and most importantly, I would like to thank my husband Jan and children Jindřich and
Alžběta, who had to be very, very patient with me during last few months. And last but not least, I
thank my parents for their assistance during my whole study.
The work was supported by Grant Agency of CR, grant no: 522/04/0549 and 206/06/1306.
ABSTRACT
Cytokinins (CKs) are one of the most important group of phytohormones influencing many
processes throughout the whole plant.
As many processes are regulated both by the light and phytohormones, the first part of this work
has been focused on evaluation of diurnal rhytmicity in levels of cytokinins and other cooperating
hormones like auxin (indol-3-acetic acid, IAA), abscisic acid (ABA) and polyamines (PA). The
changes in activity of selected enzymes participating in metabolism of the above mentioned
phytohormones were followed as well. Diurnal variation of phytohormones was tested in tobacco
leaves (Nicotiana tabacum L. cv. Wisconsin 38) grown under a 16/8 h (light/dark) period.
The main peak of the physiologically active forms of CKs, found after the middle of the light
period, coincided well with the maximum of IAA and PA levels and with activity of the
corresponding enzymes. The achieved data indicate that metabolism of CKs, IAA and PAs is tightly
regulated by the circadian clock.
The other part of the study has been focused on changes in the contents of CKs, IAA and ABA in
transgenic tobacco plants with altered cytokinin metabolism, achieved via the over-expression of
particular enzymes participating in CK metabolism (biosynthesis, degradation and reversible
conjugation). As CKs are known to be involved in plastid differentiation and function as well as
part of CK biosynthetic pathway is localized in plastids, the impact of modulation of CK
metabolism in cytoplasm on CK levels in the whole leaves and isolated chloroplasts was compared.
The achieved results suggest that plant hormone compartmentation plays an important role in
hormone homeostasis and that the chloroplasts are relatively autonomous organelles with respect to
regulation of CK metabolism.
Finally, we studied the impact of water deficit progression on CK, IAA and ABA levels in upper,
middle and lower leaves and roots of wild type tobacco plants and two transformants overexpressing trans-zeatin O-glucosyltransferase gene (ZOG1) either under constitutive 35S or
senescence-inducible SAG12 promoters. During drought stress, ABA content strongly increased,
especially in upper leaves. A delay in ABA elevation was observed in plants with constitutive overexpression of ZOG1, which indicates that high CK levels might postpone stress sensing. As drought
progressed, content of bioactive CKs in leaves gradually decreased, being maintained longer in the
upper leaves of all tested genotypes. Establishment of active CK gradient in the favour of upper
leaves suggests preferential protection of these leaves. During drought, significant accumulation
both of CKs and IAA occured in roots, indicating the role of these hormones in root response to
severe drought, which involves the stimulation of primary root growth and inhibition of lateral root
initiation and formation.
ABSTRAKT
Cytokininy (CK) jsou jednou z nejvýznamnějších skupin rostlinných hormonů, které ovlivňují
mnoho fyziologických dějů probíhajících v rostlinách.
Jelikož je známo, že mnoho procesů je regulováno jak fytohormony, tak světlem, byla první část
této práce zaměřena na diurnální rytmy regulace hladin jednotlivých cytokininů a dalších
fytohormonů - auxinu (kyseliny indol-3-octové, IAA), kyseliny abscisové (ABA) a polyaminů
(PA). Současně byly sledovány i změny v aktivitách enzymů účastnících se metabolismu CK a PA.
Diurnální změny byly měřeny v listech rostlin tabáku (Nicotiana tabacum L. cv. Wisconsin 38),
které byly pěstovány v režimu osvětlení 16/8 h světlo/tma.
Maximální hladina fyziologicky aktivních CK, naměřená po 9 h světla (tj., uprostřed světelné fáze),
velmi dobře odpovídá maximům IAA a PA a je rovněž v souladu s aktivitami odpovídajících
enzymů. Tyto údaje nasvědčují tomu, že metabolismus výše uvedených
hormonů vykazuje
diurnální rytmus.
Další část práce byla zaměřena na změny hladin CK, IAA a ABA v transgenních rostlinách tabáku,
které měly pozměněný metabolismus cytokininů způsobený zvýšenou expresí vybraných enzymů
zapojených do jejich metabolismu (biosyntézy, degradace a reverzibilní konjugace).Vzhledem
k tomu, že CK hrají důležitou roli ve funkci a diferenciaci plastidů a zároveň velká část biosyntézy
CK probíhá právě v plastidech, porovnávali jsme dopad modulace metabolismu CK v cytoplazmě
na hladiny CK na úrovni listů a izolovaných chloroplastů. Výsledky svědčí o tom, že
kompartmentalizace CK hraje velmi významnou roli v jejich homeostázi a zároveň, že chloroplasty
jsou relativně autonomními organelami vzhledem k regulaci metabolismu cytokininů.
Poslední část této práce se zabývala dopadem stresu z nedostatku vody na hladiny CK, IAA a ABA
na horní, střední a dolní listy a kořeny kontrolních rostlin tabáku a dvou transformantů s
nadprodukcí genu pro trans-zeatin O-glukosyltransferasu, a to buď pod konstitutivním 35S nebo
indukovatelným SAG12 promotorem, který spouští expresi genu až během postupující senescence.
Během sucha hladiny ABA velmi výrazně vzrostly, zvláště v horních listech. U rostlin s
konstitutivní expresí ZOG1, a tedy se zvýšeným „turn-over“ CK již před začátkem stresu, byl tento
nástup pozorován se zpožděním. S prohlubujícím se stresem obsah bioaktivních CK v listech
postupně klesal, nejpomaleji v horních listech, a to u všech studovaných genotypů. Ustavení
gradientu aktivních CK ve prospěch nejmladších listů naznačuje jejich prioritní ochranu. Během
sucha byla dále pozorována významná akumulace CK a IAA v kořenech, což poukazuje na jejich
roli při změně morfologie kořenového systému v reakci na silný stres suchem, zejména stimulaci
růstu primárních kořenů a inhibici zakládání a růstu laterálních kořenů.
ABBREVIATIONS
ABA
ADC
ADP
AHK
AHK4/CRE1/WOL
AHP
AMP
APRR
ARR
AtCKX
AtIPT
ATP
cAMP
Cad
Cdc2
CDK
CK
CKX
CYCD3
cZ
DAO
DHZ
DMAPP
DNA
dSAM
GARP domain
HPt
IAA
i6Ade
i6Ado
IPT
iP
iP7G
iPMP
iPR
iPRMP
iPRDP
iPRTP
LOG
MEP
MVA
mRNA
NADPH
ODC
PA
Put
PCA
abscisic acid
arginine decarboxylase
adenosine 5´-diphosphate
Arabidopsis thaliana histidine kinases
cytokinin receptor (histidine kinase 4/cytokinin response/wooden leg)
Arabidopsis thaliana HPt
adenosine 5´-monophosphate
Arabidopsis thaliana pseudo response regulator
Arabidopsis thaliana response regulator
Arabidopsis thaliana cytokinin oxidase/dehydrogenase
Arabidopsis thaliana isopentenyl transferase
adenosine 5´-triphosphate
cyclic adenosine 5´-monophosphate
cadaverine
gene encoding the major CDK
cyclin-dependent protein kinase
cytokinin
cytokinin oxidase/dehydrogenase
D-type cyclin
cis-zeatin
diamine oxidase
dihydrozeatin
dimethylallyl diphosphate
deoxyribonucleic acid
decarboxylated S-adenosylmethionine
a plant-specific, conserved DNA binding domain
protein with a histidine-containing phosphotransmitter domain
indol-3-acetic acid
N6-(2-isopentenyl)-adenine
N6-(2-isopentenyl)-adenosine
isopentenyl transferase
N6-(2-isopentenyl)-adenine
N6-(2-isopentenyl)-adenine 7-glucoside
isopentenyladenosine-5´-monophosphate
iP riboside
iP riboside 5´-monophosphate
iP riboside 5´-diphosphate
iP riboside 5´-triphosphate
cytokinin nucleoside 5´-monophosphate phosphorylase (called “lonely
guy”)
methylerythritol phosphate
mevalonate
messenger ribonucleic acid
nicotinamide adenine dinucleotide phosphate (reduced form)
ornithine decarboxylase
polyamine
putrescine
perchloric acid
Pssu-ipt
RT-PCR
RuBisCO
SAG
Sho
Spm
Spd
T-DNA
Ti-plasmid
tRNA
tRNA-IPT
tZ
UDPG
UDPX
ZOG1
Zm-p60.1
non-rooting transgenic tobacco with ipt under the control of the
promoter of small subunit of RuBisCO
reverse transcription polymerase chain reaction
ribulose-1,5-bisphosphate carboxylase/oxygenase
senescence-associated gene
(„shooting“) IPT homologue gene from Petunia hybrida
spermine
spermidine
transfer deoxyribonucleic acid
from „tumor inducing“ plasmid
transfer ribonucleic acid
tRNA-isopentenyl transferase
trans-zeatin
uridine diphosphoglucose
uridine diphosphoxylose
zeatin O-glucosyltransferase
gene for β-glucosidase
CONTENT
1. INTRODUCTION.............................................................................................1
1. 1. Discovery of cytokinins..........................................................................................1
1. 2. Structure of cytokinins.........................................................................................2
1. 3. Cytokinin metabolism............................................................................................2
1. 4. Degradation of cytokinins.....................................................................................6
1. 5. Biosynthesis of cytokinins.....................................................................................7
1. 6. Cytokinins and biological functions.....................................................................11
1. 6. 1. Cytokinins and cell cycle regulation...............................................................11
1. 6. 2. Cytokinins and leaf senescence.......................................................................11
1. 6. 3. Cytokinins and chloroplast development........................................................12
1. 6. 4. Cytokinins and movement of nutrients...........................................................12
1. 6. 5. Cytokinins and signalling................................................................................13
1. 7. Polyamines – plant growth regulators.................................................................15
1. 8. Polyamine metabolism...........................................................................................16
2. AIMS AND SCOPES.........................................................................................17
3. PLANT MATERIALS AND METHODS........................................................18
3.1. Growing conditions and collecting samples..........................................................18
3. 1. 1. Isolation of CKs and study of diurnal rhythms...............................................18
3. 1. 2. Isolation of PAs and study of diurnal rhythms................................................18
3. 1. 3. Isolation of CKs and study of altered CK metabolism....................................18
3. 1. 4. Isolation of CKs and study of water deficit.....................................................19
3. 1. 5. Incubation of samples with tritiated trans-zeatin............................................19
3. 1. 6. Chloroplast isolation........................................................................................19
3. 2. Plant hormone extraction and purification.........................................................19
3. 2. 1. Extraction and purification of CKs.................................................................19
3. 2. 2. Extraction and purification of PAs..................................................................19
3. 3. Chromatographic methods...................................................................................19
3. 3. 1. HPLC of radiolabelled trans-zeatin and its metabolic products.....................19
3. 3. 2. HPLC of auxin and abscisic acid....................................................................19
3. 3. 3. HPLC/MS........................................................................................................19
3. 3. 4. PA analysis.......................................................................................................19
3. 3. 5. Chlorophyl determination................................................................................19
3. 3. 6. Chloroplast integrity........................................................................................19
3. 4. Determination of enzymes activities.....................................................................19
3. 4. 1. Determination of CKX activity.......................................................................19
3. 4. 2. Determination of β-glucosidase activity.........................................................19
3. 4. 3. Determination of ODC, ADC activity.............................................................19
3. 4. 4. Determination of DAO activity.......................................................................20
3. 5. Microscopic methods.............................................................................................20
3. 5. 1. Transmission electron microscopy..................................................................20
3. 6. Molecular methods.................................................................................................20
3. 6. 1. RNA extraction and reverse transcription.......................................................20
3. 6. 2. qRT-PCR..........................................................................................................20
3. 7. Determination of relative water content..............................................................20
4. RESULTS AND DISCUSSION........................................................................21
4. 1. Diurnal rhythms of cytokinins, auxin and abscisic acid.....................................21
4. 2. Diurnal rhythms of polyamines............................................................................23
4. 3. The study of chloroplast autonomy in regulation of CK levels..........................25
4. 4. The role of CK in responses to water deficit........................................................27
5. SUMMARY........................................................................................................31
LITERATURE.......................................................................................................32
SUPPLEMENT 1...................................................................................................48
SUPPLEMENT 2...................................................................................................55
SUPPLEMENT 3...................................................................................................64
SUPPLEMENT 4...................................................................................................77
1. Introduction
At the end of the 19th century, German botanist Julius von Sachs proposed a theory
concerning the existence of chemical signals, which would play an important role in
communication among plant organs. Since that time, a lot of information has been collected
on substances, which could influence and regulate growth and development in plants. These
substances are generally called plant growth regulators and can be separated into two groups:
plant hormones (phytohormones) and other substances with growth regulatory activity.
Phytohormones are naturally occuring substances which belong into nine major groups:
auxins, cytokinins, abscisic acid, ethylene, gibberellins, brassinosteroids, jasmonic acid,
salicylic acid and strigolactones. Recently, a group of peptide hormones has emerged. Other
molecules with regulatory activity, recognized as polyamines or phenolics, are not classified
as phytohormones because they are effective in relatively high concentrations.
Due to the sessile character of plants and their cell totipotency, mechanisms of plant hormone
functions differ from those in animals and humans. Phytohormones are not produced in
special glands, but in a wide range of tissues. In contrast to animal hormones, each
phytohormone influences several physiological processes and on the other hand, one process
is influenced (positively or negatively) by more than one phytohormone or growth regulator.
Plant growth and development is thus controlled by a very subtile balance among levels of
individual phytohormones.
1. 1. Discovery of cytokinins
Cytokinins have been defined as a class of plant hormones crucial to cell division
(“cytokinesis”). Discovery of the first cytokinin, kinetin, by Skoog, Miller and associates in
1955 in herring sperm DNA (Miller et al. 1955) triggered an avalanche of experiments with
the aim to isolate and identify other compounds with CK activity. This effort led to discovery
of diphenylurea as a synthetic compound with CK activity (Shantz et al. 1955), trans-zeatin,
the most active naturally occuring CK in plants (Letham 1963), and CKs with aromatic side
chain (Horgan et al. 1973).
1
1. 2. Structure of cytokinins
Naturally occuring CKs are N6-substituated adenine derivatives with either isoprenoid or
aromatic side chain (Strnad 1997, Mok and Mok 2001).
The abundance of individual CK derivatives varies significantly in the dependence on plant
species, tissue or developmental stage. Isoprenoid CKs include derivatives of N 6-(Δ2isopentenyl)-adenine (iP), trans-zeatin (tZ), cis-zeatin (cZ) and dihydrozeatin (DHZ) (Fig. 1).
CK bases and, to a lesser extent, also ribosides are considered to be physiologically active CK
forms, as they are recognised by CK receptors (Inoue et al. 2001, Yamada et al. 2001, Spichal
et al. 2004). There are nevertheless big differences in biological activity of individual bases
and ribosides due to the structure and conformation of the side chain, trans-zeatin being the
most active naturally occurring CK.
Fig. 1. The structure of bioactive cytokinins.
1. 3. Cytokinin metabolism
Regulation of the level of bioactive CKs involves their metabolic conversions. CK
modifications include changes of either purine ring or of the side chain. Those reactions may
significantly affect physiological activity of the respective CK. Interconversion of CK
2
phosphates to ribosides and CK bases is probably a major feature of CK metabolism (Chen
1981). While CK ribosides (especially trans-zeatin riboside) have been considered as CK
transport forms, CK phosphates were identified as biosynthetic intermediates (Kakimoto
2001a, Takei et al. 2001). However, they may be also the product of metabolic conversion of
CK bases and ribosides by adenosinekinase (EC 2. 7. 1. 20), as was shown in Physcomitrella
patens (von Swartzenberg et al. 1998) or tobacco cells (Kwade et al. 2005), or adenine
phosphoribosyltransferase (EC 2. 4. 2. 7; Chen et al. 1982, Shukla and Sawhney 1997). Due
to the structure similarity with other, highly abundant, purine derivatives, CKs may share
some metabolic steps with the purine metabolic pathway, although amount and type of
metabolites can be influenced by incubation conditions and the type of cells used in
experiment (Chen et al. 1985, Letham 1994). These enzymes, participating in CK
interconversions have higher affinity for adenine, adenosine and AMP than for corresponding
CK.
5´-nucleotidase (EC 3. 1. 3. 5), isolated from wheat germ and tomato (Chen and Kristopeit
1981a, Burch and Stuchbury 1987) can convert cytokinin nucleotide to its corresponding
nucleoside, which can be further metabolized to CK free base by for example adenosine
nucleosidase (EC 3. 2. 2. 7), which is responsible for deribosylation of CK nucleoside (i6Ado)
to CK free base (i6Ade) (Chen and Kristopeit 1981b).
Similarly, there exist enzymes catalyzing conversion of free bases to corresponding
nucleosides or nucleotides. For example, adenosine phosphorylase (EC 2. 4. 2. 1), an enzyme
requiring ribose-1-phosphate for ribosylation of CK base strongly favoured CK nucleoside
formation (Chen and Petschow 1978b).
Nucleotides can be formed either directly from CK bases due to the enzyme adenine
phosphoribosyltranferase (EC 2. 4. 2. 8) in presence of 5´-phosphoribose-1´-pyrophosphate
(Chen et al. 1982) or from corresponding nucleoside by adenosine kinase (EC 2. 7. 1. 20).
Modifications of the purine ring of CK include N-glycosylation at the 3, 7, or 9 position (Fox
et al. 1971, 1973, McGaw et al. 1984) and conjugation of alanine at the 9 position of zeatin,
forming lupinic acid.
Although N-glucosylation in 3 position has been reported in some plant species (Letham
1978), specific N-glucosyltransferase has not been identified yet.
N-glucosylation in 7 and 9 position leads to metabolically stable compounds with very low
biological activity, which are resistant to glucosidases (Brzobohatý et al. 1993), although
recent data show that N-glucoside in position 9 could be cleaved by some isoforms of CKX
(Galuszka et al. 2007, Kowalska et al., 2010). For these reasons N-glucosides are considered
3
to be involved in deactivation and detoxification (Letham and Palni 1983), especially in case
of sudden excess of bioactive CKs (e.g. by exogenous application). Two Nglucosyltransferases from A. thaliana were identified in 2004 by Hou et al.. Both of them
recognized all classical CKs and form N-glucoside in 7 and 9 position in in vitro assay.
Fig. 2. Enzymes (E) and genes (G) involved in the modifications of N6-isoprenoid side chain.
According to Mok and Mok (2001).
4
Modifications of the isoprenoid side chain of CKs have been also very often observed and
were detected in almost all plant tissues (Fig. 2). The most common modification is
glycosylation of the hydroxyl group of the side chain of t-Z, DHZ and cis-Z, resulting in CK
O-glucosides or O-xylosides. O-glycosylation is a reversible process, the deglycosylation,
forming the active aglycone, is catalyzed by β-glucosidase (EC 3. 2. 1. 21; Brzobohatý et al.
1993). Thus O-glucosylation may function as an important mechanism in the regulation of
levels of active CKs. The gene for β-glucosidase (Zm-p60.1) was isolated from maize
coleoptiles (Campos et al. 1992; Esen 1992). Recently, it was shown that overexpression of
Zm-p60.1 can disturb trans-zeatin metabolism (Kiran et al. 2006).
Trans-zeatin O-glucosyltransferase (EC 2. 4. 1. 203) was isolated from Phaseolus lunatus
(Dixon et al. 1989) and trans-zeatin O-xylosyltransferase (EC 2. 4. 1. 204) was found in P.
vulgaris (Turner et al. 1987). The former enzyme utilizes as donor substrates UDPG or
UDPX, but the affinity to UDPG and UDPX differs and that to UDPG is much higher. The
latter enzyme accepts only UDPX.
The only CK substrate which is recognized by both enzymes is DHZ and it forms
dihydrozeatin O-xyloside in presence of UDPX (Dixon et al. 1989, Martin et al. 2000, Mok et
al. 2000). As DHZ O-glucoside was found in P. vulgaris (Wang et al. 1977), but reported
trans-zeatin O-glucosyltransferase from beans does not catalyze the formation of this
compound in the presence of UDPG, it seems that other O-glucosyltransferases converting
DHZ to DHZ O-glucoside exist.
In 2001, an O-glucosyltransferase (EC 2. 4. 1. 215) with 41% identity to the Phaseolus transzeatin O-glucosyltransferase, but highly specific to cis-zeatin, was discovered in maize
(Martin et al. 2001b). Substrates of this enzyme are cis-zeatin (but not trans-zeatin, DHZ or
cis-zeatin riboside) and UDPG (not UDPX). The discovery of the enzyme with function
parallel to those for trans-zeatin led to a new view on the importance and relevance of cis-CK
derivatives in plants. Existence of cis-specific regulatory mechanisms in plants suggests that
cis-zeatin and its derivatives may play a more important role in CK homeostasis than it was
thought. Till now, cis-CK derivatives were isolated from a number of plant species, including
rice (Takagi et al. 1989), potato (Suttle and Banowetz 2000), wheat and oats (Parker et al.
1989), lupins (Emery et al. 2000), tobacco (Dobrev et al. 2002).
Among other side chain modifications of CKs pertain reduction of trans-zeatin to form DHZ
and cis-trans isomeration. Reduction of the trans-zeatin side chain is catalyzed by a zeatin
reductase (EC 1.3.1.69) isolated from immature seeds of P. vulgaris (Martin et al. 1989) and
pea (Gaudinová et al. 2005) forming dihydrozeatin. The enzyme shows high specificity for
5
trans-zeatin and does not recognize other CK derivatives (cis-zeatin, iP) as substrates and
requires NADPH as a cofactor. As cytokinin oxidase/dehydrogenase does not use DHZ as a
substrate, reduction of trans-zeatin to DHZ may preserve a cytokinin activity (Gaudinová et
al. 2005).
A cis-trans zeatin isomerase was partially purified from Phaseolus (Bassil et al. 1993). As the
enzyme favours conversion from cis- to trans- isomer, isomeration may be a way to increase
amount of highly active trans-zeatin in plant tissues.
Also another enzyme, hydroxylase, catalyzing a conversion of iP to zeatin and iPR to transzeatin riboside was found in a microsomal fraction of cauliflower (Chen and Leisner 1984).
1. 4. Degradation of cytokinins
The last but definitely not least side chain modification is CK degradation mediated by
cytokinin oxidase/dehydrogenase (CKX), which irreversibly inactivates free CK bases and
nucleosides with an unsaturated N6-side chain in a single enzymatic step (Fig. 3). Changes in
CKX activity alter the concentrations of bioactive cytokinins in tissues, therefore CKX
enzymes play an important role in controlling CK levels and thus influence cytokinin
dependent processes.
The first report on degradation of CKs by an enzyme system came in 1968 by Chen et al.
(1968). They observed degradation of iPR to adenosine by an enzyme preparation from
tobacco callus culture. A few years later, conversion of labelled i6Ade to adenine in crude
tobacco cell culture extracts was reported (Pačes et al. 1971). In 1974, Whitty and Hall
described similar activity in Zea mays kernels and named the enzyme cytokinin oxidase.
Since then, CKX activity was observed in many other plant species like beans, poplar, wheat
(Jones and Schreiber 1997). The various CKX could differ in their biochemical properties like
substrate affinity, etc. CKX from majority of plants are glycoproteins (Armstrong 1994),
however, non-glycosylated CKX or CKX with low degree of glycosylation were observed in
several plants including tobacco and Phaseolus (Motyka et al. 1996, 2003, Kamínek and
Armstrong 1990). CKX glycosylation affects enzyme localization, the apoplastic forms being
glucosylated. The occurrence of several CKX isoenzyme (e.g. 7 members in Arabidopsis),
which differ in their localization as well as in their regulation, indicate the involvement of
CKX in regulation of different CK controlled processes (Schmülling et al. 2003).
For a long time, CKX was supposed to require copper in its active center and molecular
oxygen as an electron acceptor for its activity. For this reason the enzyme was named
6
cytokinin oxidase and classified as copper amine oxidase (EC 1. 4. 3. 6) (Whitty and Hall
1974, Hare and Van Staden 1994a). After cloning of maize gene, the view was corrected,
because a flavin-binding site was observed (Morris et al. 1999) and a few years later atomic
absorption analysis did not prove the presence of copper (Bilyeu et al. 2001). Also, the
opinion that molecular oxygen is the electron acceptor was changed after finding that CKX
can use a variety of artificial electron acceptors under anaerobic conditions (Galuszka et al.
2001, Frébort et al. 2002). Regarding these evidences that CKX are flavine enzymes not using
oxygen as electron acceptors, enzyme was reclassified as cytokinin dehydrogenase (EC 1. 5.
99. 12).
Degradation of aromatic CKs still remains to be solved, as CKX exhibit only very low
reaction rate towards aromatic CKs, e.g. benzyladenine (Frébortová et al. 2004).
Fig. 3. CKX function scheme
1. 5. Biosynthesis of cytokinins
CK homestasis is regulated not only by CK interconversion, transport and degradation, but
also by the control of the rate of their biosynthesis. The occurence of modified bases,
including the isopentenylated ones, in plant tRNAs led to the hypothesis that the breakdown
of tRNA (which results in the release of iP or zeatin derivatives) could be a potential
mechanism for biosynthesis of CKs as free hormones (Hall 1970, Murai 1994, Morris 1995).
CK moieties in plant tRNA include iPR, cis-zeatin riboside, trans-zeatin riboside, 2methylthio-iPR, 2-methylthio-zeatin riboside (Horgan 1984). It is generally thought that
purine moieties in tRNAs are first isopentenylated and then can be further modified by side
chain hydroxylation and/or ring methylation to give CK derivatives (Cherayil and Lipsett
1977). An enzyme that catalyzes the reaction between Δ2-isopentenyl diphosphate and
unmodified tRNA is Δ2-isopentenyl diphosphate:tRNA- Δ2-isopentenyltransferase (EC 2. 5. 1.
7
8) and has been partially purified from various sources (Hall 1971). Recently, the
corresponding gene was identified in Arabidopsis thaliana (Golovko et al. 2002). Also the
theory that tRNA is a source especially of cis-type CKs was supported by finding that
isoprenoide moiety of cis-zeatin is a product of mevalonate pathway localized in cytoplasm as
well as tRNA-IPT2 (Kasahara et al. 2004). The role of tRNA-IPT in synthesis of cis-zeatin
was demonstrated recently (Miyawaki et al. 2006). However, the slow turnover rate of tRNAs
does not seem sufficient to be a sole source of CKs, especially when the necessity of a precise
control of the level of bioactive CKs is taken into consideration.
The first demonstration of an enzyme catalyzing transfer of the isopentenyl moiety from
dimethylallyl diphosphate (DMAPP) to AMP was reported in 1978 in Dictyostelium
discoideum (Taya et al. 1978). The enzyme was named DMAPP:AMP isopentenyltransferase.
ATP, ADP or cAMP did not function as isopentenyl acceptors. The reaction product,
isopentenyladenosine-5´-monophosphate
(iPMP)
was
converted
to
iP.
The
first
characterization of ipt (tmr) gene came from Agrobacterium tumefaciens, a crown gallforming bacterium (Akiyoshi et al. 1984, Barry et al. 1984). The tmr is encoded on the TDNA region of the Ti-plasmid and during the infection is introduced into the host plant
chromosome. The tmr (ipt) gene was cloned and expressed in E. coli and its extracts were
shown to catalyze production of iPMP from DMAPP and AMP. It has been assumed that
similar biosynthetic pathway functions also in higher plants. Many attempts have been
performed to isolate and characterize plant ipt genes, but due to their low expression levels
and enzyme instability, only little biochemical information was reported. In 1979, Chen and
Melitz partially purified an enzyme from cytokinin-autotrophic cultured cells of tobacco and a
few years later, Blackwell and Horgan (1994) did the same from kernels of Zea mays.
Recently, ipt genes have been identified in Arabidopsis thaliana (Kakimoto 2001a, Takei et
al. 2001a), petunia (Zubko et al. 2002), hop (Sakano et al. 2004), soybean (Ye et al. 2006),
Chinese cabbage (Ando et al. 2005), rice (Sakamoto et al. 2006), pingyitiancha (Malus
hupehensis, Peng et al. 2008), maize (Brugiere et al. 2008) and finally, a prokaryote-type
tRNA-IPT gene from the moss Physcomitrella patens was characterised (Yevdakova and von
Schwartzenberg 2007). The completion of Arabidopsis genomic sequence showed a total of
nine ipt-homologues described as AtIPT1 to AtIPT9. A phylogenetic analysis revealed that
AtIPT2 (Golovko et al. 2002) and AtIPT9 (Kakimoto 2001a) encode a putative tRNAisopentenyltransferases, while the other seven AtIPTs participate in de novo CK synthesis
(Kakimoto 2001a, Takei et al. 2001a). It had been assumed, according to the studies with
prokaryotes, that the first step in CK biosynthesis in plants could be isopentenylation of AMP.
8
But biochemical studies showed that plant IPTs use ADP or ATP rather than AMP as prenyl
acceptors, resulting in production of iP riboside 5´-diphosphate (iPRDP) or iP riboside 5´triphosphate (iPRTP) (Kakimoto 2001a, Sakano et al. 2004) (Fig. 4).
Plants have two possible biosynthetic pathways for the prenyl group of CKs – the
methylerythritol phosphate (MEP) pathway in plastids (Lichtenhaler 1999) and the
mevalonate (MVA) pathway in the cytosol (Rohmer 2003). Both pathways produce the
isoprenoid precursors isopentenyl diphosphate and DMAPP.
In spite of the fact that MEP and MVA pathways exist in different subcellular compartments,
exchange of precursors between these two pathways was found (Rohmer 1999, Kasahara et
al. 2002). It has been observed that isoprene units from both pathways were incorporated into
a single downstream isoprenoid (Kasahara et al. 2002).
Using a 13C labeled precursor specific to the MEP and MVA pathways, it was demonstrated
that both pathways are able to supply DMAPP to CKs in Arabidopsis (Kasahara et al. 2004).
They proved the dominant role of the MEP pathway in the biosynthesis of iP- and tZ- type
CK. Kasahara et al. (2004) postulated that the majority of the prenyl group of cZ-type CK
comes from MVA pathway. They also used fusion of AtIPTs with green fluorescent protein
and showed that a large number of the AtIPTs (AtIPT1, 3, 5, 8) are located in plastids
(suggesting plastids to be a major subcellular compartment for iP- and tZ-type CK
biosynthesis in higher plants), AtIPT4 and 7 are localized in the cytosol and mitochondria.
Recently, using in vivo isotope labeling, an iPRMP-independent pathway for the biosynthesis
of tZ-type CK was proposed, which involves binding of hydroxylated derivate of DMAPP
directly to AMP (Åstot et al. 2000).
In order to obtain a physiological activity, CK nucleotides must be converted to bases. It was
assumed that this process has two steps operated by nucleosidases and nucleotidases
participating in general metabolism of purine (Chen and Kristopeid 1981a,b). Recently, a new
metabolic pathway releasing a free base directly from nucleotide was identified. The reaction
is catalyzed by cytokinin nucleoside 5´-monophosphate phosphorylase (called LOG - “lonely
guy”), which utilizes only mononucleotides (Kurakawa et al. 2007, Kuroha et al. 2009).
In spite of the fact that aromatic CK exhibit strong physiological activity, their biosynthesis
and degradation pathways remain still unclear.
Regulation of CK biosynthesis can be studied at different levels. Using constructs of AtIPT
promoter:reporter genes, tissue- and organ-specific patterns of CK biosynthesis were revealed
(Miyawaki et al. 2004, Takei et al. 2004). CK biosynthesis is significantly affected by other
phytohormones, namely auxin. In Arabidopsis, accumulation of AtIPT5 and 7 is promoted by
9
auxin in roots, while high CK levels affect negatively expression of AtIPT1, 3, 5 and 7
(Miyawaki et al. 2004). In shoots, auxin down-regulate IPT expression (Nordström et al.
2004, Tanaka et al. 2006). Genes for CKX in maize are upregulated by CK and abscisic acid
(ABA) (Brugiere et al. 2003). This kind of hormone cross-talk may play a role in regulation
of plant development and in the plant responses to environmental conditions.
CKs are also pivotal signalling substances delivering information about nitrogen availibility
from root to shoot via the xylem vessels (Takei et al. 2001, 2002). Recently, AtIPT3 and 5
were revealed to be regulated differently, depending on the nitrogen sources available.
Whereas AtIPT3 specifically responses to NO3- under nitrogen-limited conditions, AtIPT5
reacts to both NO3- and NH4+ under long-term treatment (Miyawaki et al. 2004, Takei et al.
2004).
Fig. 4. Scheme of cytokinin biosynthesis (according to Kamada-Nobusada and Sakakibara,
2009)
10
1. 6. Cytokinins and biological functions
1. 6. 1. Cytokinins and cell cycle regulation
Regulation of cell division and growth involves tight control of cell cycle progression. The
eukaryotic cell cycle consists of four phases, which lead to the reproduction of genetic
material and its transport to the next cell generation.
CKs were discovered due to their ability to promote cell division in presence of optimal level
of auxin and it seems that CKs affect and control various activities through the cell cycle.
There is an evidence that CKs play an important role especially in G2/M and G1/S transition
(Hare and Van Staden 1997, Riou-Khamlichi et al. 1999, Dobrev et al. 2002).
It is suggested that both auxin and CK regulate the cell cycle by controlling the activity of
cyclin-dependent protein kinases (CDKs), enzymes regulating eukaryotic cell cycle.
Expression of the gene encoding the major CDK (Cdc2) is regulated by auxin, but this CDK
is inactive until an inhibitory phosphate group from the Cdc2 is removed. And this is the role
for CKs, which activate phosphatase catalyzing removal of the phosphate group from Cdc2
(Zhang et al. 1996) and enable the cell transition from the G2 to M-phase. CKs also increase
expression of the CYCD3 gene encoding a D-type cyclin that leads to elevation of cell
proliferation and enables transition from G1 to S-phase (Riou-Khamlichi et al. 1999, Soni et
al. 1995).
1. 6. 2. Cytokinins and leaf senescence
Leaf senescence is a form of programmed cell death and the final phase of leaf development
during which the nutrients contained in the macromolecules (e.g. nucleic acids, proteins) are
metabolized to the basic components (e.g. amino acids) and transported from old, senescing
leaves to the young productive leaves and developing seeds. This strategy is very
advantageous, especially under adverse conditions where nutrients are limiting factors
(Noodén 1988).
Senescence is a highly regulated process that requires de novo synthesis of many nuclear
genes, referred as senescence-associated genes (SAGs) (Gan and Amasino 1997, BuchananWollaston 1997, Weaver et al. 1998). Senescence can be triggered by different environmental
11
and endogenous stimuli such as drought, nutrient deficiency, temperature, pathogen attack,
age, concentration of plant hormones etc.
CKs have been showed to delay and, in some cases, reverse the leaf senescence process (Gan
and Amasino 1996), as revealed by re-greening of tissues after CK elevation.
To study leaf senescence and overcome problems with CK overproduction in transgenic
plants, Gan and Amasino (1995) developed a gene construct using highly senescence specific
promoter from SAG12. This gene is thought to be the most specific for natural senescence,
showing no detectable expression in young leaves and not accumulating until the leaf is 20 %
chlorotic (Weaver et al. 1998). When SAG12 promoter is used to drive IPT expression,
transcription of IPT is triggered and production od functional enzyme is activated only when
the leaf senescence is stimulated.
In our work, we used transgenic plants in which the SAG12 promoter drove expression of
ZOG1, gene for another key enzyme participating in CK metabolism, as described earlier. We
used also plants with 35S promoter driven expression of ZOG1 to compare drought stress
response of plants with senescence-induced CK elevation and those with constitutive
stimulation of CK turn-over.
1. 6. 3. Cytokinins and chloroplast development
Already 10 years after discovery of kinetin as a plant hormone (Miller et al. 1955), it was
shown that kinetin has been able to promote chloroplast differentiation from proplastids
(Stetler and Laetsch 1965). It has been proved that exogenously applied CK can induce partial
development of chloroplasts from proplastids, amyloplasts and etioplasts (Chory et al. 1994,
Kusnetsov et al. 1994). The importance of CKs for plastid development can be deduced from
partial localization of the CK biosynthetic pathway to this compartment (Kasahara et al.
2004). CKs are also known to function in the protection of photosynthetic machinery
(Chernyadev 2009).
1. 6. 4. Cytokinins and movement of nutrients
In general, CKs have been shown to play a major role in the regulation of many processes
associated with the supply of nutrients to growing tissues and with regulation of the sink
strength.
12
Several reports show the effect of CKs on the expression of genes for nitrate reductase (Lu et
al. 1990, Hänsch et al. 2001), a hexose transporter (Ehneß and Roitsch 1997) and an
extracellular invertase, catalyzing the hydrolytic cleavage of the transport sugar sucrose
released into the apoplast, which seems to be an essential component of the molecular
mechanism of the delay of senescence by CKs (Ehneß and Roitsch 1997, Lara et al. 2004). It
is suggested that de novo synthesis and recycling of storage forms (O-glucosides) might
contribute to CK accumulation in the roots, followed by translocation to the leaves via the
xylem during recovery from nitrogen deficiency in maize (Takei et al. 2001, 2002). Nitrogendependent accumulation of CKs was also observed in roots of Arabidopsis thaliana (Takei et
al. 2002) and barley (Samuelson and Larsson 1993). The main CKs accumulated in response
to nitrate supply in roots are generally isopentenyladenine-type CKs, which are transformed
into trans-zeatin riboside, when entering the xylem, and into free base trans-zeatin, which
accumulates in the leaves (Takei et al. 2001).
1. 6. 5. Cytokinins and signalling
As plants cannot escape from adverse environmental conditions, their lives depend on their
ability to react quickly and efficiently. Therefore they have developed different signal
perception and transduction systems.
In higher plants, the major role in CK perception and signalling is played by the twocomponent system. Until recently, signal transduction via the two-component system was
thought to be only a prokaryotic concern. But now it was proved that this mechanism
participates in higher plants, too (Lohrmann and Harter 2002, Hass et al. 2004a). The twocomponent system serves as a sensing/responding mechanism. It is composed of hybrid
histidine kinases, histidine-containing phosphotransfer domain proteins and response
regulators that are biochemically linked by His-to-Asp phosphorelay (Fig. 5). As a reaction to
endogenous or exogenous stimuli, autophosphorylation of the histidine kinase at a conserved
histidine residue is induced and signalling is initiated, followed by a multistep system with
additional phosphorylation steps. The hybrid histidine kinase carries an additional receiver
domain, usually at the COOH-terminal end. Then, the phosphoryl group is transferred to a
conserved Asp residue of its own receiver domain instead of transferring the phosphoryl
group directly to the response regulator.
13
Additional proteins with a histidine-containing phosphotransmitter (HPt) domain then
percieve the phosphoryl group from the receiver domain of the hybrid histidine kinase and
transfer it to the receiver domain of the response regulator, what induces a conformational
change in output domain that alters its biological activity (West and Stock 2001, Hass et al.
2004a).
The plant species intensively studied in most detail is Arabidopsis thaliana. Sequence analysis
of Arabidopsis genome has revelead 8 histidine kinases (AHKs), 5 HPt proteins (AHPs), 23
response regulators (ARRs) and 9 pseudo response regulators (APRRs) (Urao et al. 2000a,
Hwang et al. 2002).
Arabidopsis AHKs can be divided into several subfamilies, where one subfamily functions as
CK receptors. It is composed of AHK2, AHK3 and AHK4/CRE1/WOL (histidine kinase
4/cytokinin response/wooden leg) (Kakimoto 2003).
The Arabidopsis HPt proteins (AHPs) are expressed ubiquitously and their transcription in not
affected by CK treatment (Tanaka et al. 2004). They are suggested to play a role as
intermediates in multistep phosphorelay to transfer the signal from AHKs to corresponding
response regulators.
Fig. 5. Cytokinin signalling. According to Werner and Schmülling (2009).
14
The Arabidopsis response regulators are divided into three groups:
1) Type-A response regulators (type-A ARRs) are relatively small and contain a receiver
domain and a short carboxyl terminus and their transcription is rapidly elevated in
response to exogenous cytokinin. They are considered to be primary response genes
(D´Agostino et al. 2000, Kiba et al. 2002, Rashotte et al. 2003). They are negative
regulators of CK signalling.
2) Type-B response regulators (type-B ARRs) are comprised of a receiver domain,
carboxy-terminal output domain that has a DNA-binding GARP domain and a
transcriptional activation domain. This type of ARRs transfers the CK signal to CK
early response genes (Imamura et al. 1999, Kiba et al. 1999).
3) Pseudo response regulators (APRRs) contain a receiver-like domain, but the invariant
amino acids including the phosphorylated Asp are substituted by other amino acids
(Hwang et al. 2002). Some observations suggest that APRRs may function as nuclear
transcription factors (Strayer et al. 2000). Other evidences suggested the crucial role
of APRRs as intrinsic elements of Arabidopsis circadian clock (Hayama and Coupland
2003, Eriksson et al. 2003).
1. 7. Polyamines – plant growth regulators
Polyamines (PA) are small organic, ubiquitous polycations with several aminogroups in their
molecules, found in all living organisms. They play an important role in regulation of growth
and development in prokaryotes and eukaryotes (Tiburcio et al. 1997). In plants, they have
been implicated in a wide range of biological processes, including stimulation of DNA
replication, transcription and translation, response to senescence, environmental stress and
infection by viruses and fungi. These biological activities are fastened on their cationic nature
(Galston et al. 1997, Bouchereau et al. 1999, Thomas and Thomas 2001, Theiss et al. 2002,
Capell et al. 2004). The major PAs found in plants are diamine putrescine (Put), triamine
spermidine (Spd) and tetramine spermine (Spm). They occur in the free form or as conjugates
bound to phenolic acids or to macromolecules such as proteins and nucleic acids. PAs were
reported to participate in stabilization of membranes and protection against free radicals. They
influence protein and nucleic acid synthesis and enzyme activities (RNAse, protease, etc.).
They have intensive cross-talk with phytohormones such as CKs, auxin, ethylene.
15
1. 8. Polyamine metabolism
The PAs are synthetized from arginine or ornithine by arginine decarboxylase (ADC) and
ornithine decarboxylase (ODC), respectively. The intermediate agmatine is then converted to
Put, which can be further transformed to Spd and Spm by transfer of aminopropyl groups
from decarboxylated S-adenosylmethionine (dSAM) catalysed by specific Spd and Spm
synthases (Slocum 1991a, Martin-Tanguy 2001). The catabolism of PAs is performed by
oxidases. It has been found that each PA is catabolized by a specific oxidase (Smith and
Marshall 1988). In order to understand their biological roles, many studies have been done to
localize PAs and their biosynthetic enzymes in plants. They have been found in cell wall
vacuoles, mitochondria and chloroplasts (Torrigiani et al. 1986, Slocum 1991b, Tiburcio et al.
1997).
PAs are involved in many plant developmental processes such as cell division, reproductive
organ development, root growth, floral initiation and development, fruit development and
ripening, leaf senescence and abiotic stress response. Changes in free and conjugated PAs
occur during these developmental processes. In general, cells undergoing division contain
high levels of free PAs synthetized via ODC, whereas cells undergoing expansion and
elongation contain low levels of free PAs synthetized via ADC (reviewed by Galston and
Kaur-Sawhney 1995).
16
2. Aims and scope
Cytokinins (CKs) are one of the most important plant hormones, which exhibit, in cooperation
with other phytohormones, multiple physiological functions.
This work has been focused on different aspects of cytokinin research, namely on:
1) monitoring of diurnal changes in CK levels (in correlation with other phytohormones
- auxin and abscisic acid) and polyamine content in tobacco plants, in order to
contribute to better understanding of the complex network of light and hormone signal
transduction pathways.
2) elucidation of the extent of chloroplast autonomy in the maintenance of CK
homeostasis determined by comparison of the effect of modulation of CK metabolism
in cytoplasm on CK content at chloroplast and whole leaf. Manipulation of CK
metabolism was achieved by over-expression of the genes for CK biosynthesis,
degradation and CK glucosylation/deglucosylation, respectively. This study included
monitoring of CK effect on changes in chloroplast ultrastructure in tobacco plants.
3) evaluation of the role of CKs (together with other hormones - auxin and ABA) in the
drought response of tobacco plants, comparing the wild-type plants with those
exhibiting increased CK turn-over before drought stress initiation or during the stress
progression (achieved by the expression of trans-zeatin O-glucosyltransferase gene,
ZOG1, under 35S or SAG12 promoters, respectively).
17
3. Plant material and methods
Plant material
1) Plants of Nicotiana tabacum L. cv. W 38 (prof. Machteld and David Mok, Oregon
State University)
2) Tobacco in vitro lines expressing an IPT homologue Sho (Shooting) gene from P.
hybrida under the control of a constitutive cauliflower mosaic virus (CaMV) 35S
promoter or construct with four 35S enhancer elements (Prof. Břetislav Brzobohatý,
Mendel University in Brno)
3) Eleven-week-old tobacco plants (35S:AtCKX3) overexpressing a gene for CK
oxidase/dehydrogenase from A. thaliana under a constitutive CaMV 35S promoter
(Prof. Thomas Schmülling, Free University, Berlin)
4) Nine-week-old Nicotiana tabacum L. cv. Samsun NN plants (Prof. Thomas
Schmülling, Free University, Berlin).
5) Eight-to nine-week-old tobacco plants (35S:P60) overexpressing a maize βglucosidase Zm-p60.1 naturally targeted to plastids under a CaMV 35S promoter
(Prof. Břetislav Brzobohatý, Mendel University in Brno).
6) Nicotiana tabacum L. cv. Petit Havana SR1 plants (Prof. Břetislav Brzobohatý,
Mendel University in Brno).
7) Eight-to nine-week-old
tobacco plants (35S:ZOG1) harbouring a trans-zeatin O-
glucosyltransferase gene from P. lunatus under a constitutive CaMV 35S promoter
(prof. Machteld and David Mok, Oregon State University)
Methods
3. 1. Growing conditions and collecting samples
3. 1. 1. For isolation of CKs and study of diurnal rhythms
see supplement 1, p. 49
3. 1. 2. For isolation of PAs and study of diurnal rhythms
see suplement 2, p. 56
3. 1. 3. For isolation of CKs and study of altered CK metabolism
see supplement 3, p. 65
18
3. 1. 4. For isolation of CKs and study of water deficit
see supplement 4, p. 78
3. 1. 5. Incubation of samples with tritiated trans-zeatin
see supplement 1, p. 49
3. 1. 6. Chloroplast isolation
see supplement 3, p. 66
3. 2. Plant hormone extraction and purification
3. 2. 1. Extraction and purification of CKs
see supplement 1, p. 49 ; supplement 3, p. 66 and supplement 4, p. 79
3. 2. 2. Extraction and purification of PAs
see supplement 2, p. 56
3. 3. Chromatographic methods
3. 3. 1. HPLC of radiolabelled trans-zeatin and its metabolic products
see supplement 1, p. 49
3. 3. 2. HPLC of auxin and abscisic acid
see supplement 1, p. 49 ; supplement 3, p. 66 and supplement 4, p. 80
3. 3. 3. HPLC/MS
see supplement 1, p. 49 ; supplement 3, p. 66 and supplement 4, p. 80
3. 3. 4. PA analysis
see supplement 2, p. 56
3. 3. 5. Chlorophyl determination
see supplement 3, p. 66
3. 3. 6. Chloroplast integrity
see supplement 3, p. 66
3. 4. Determination of enzymes activities
3. 4. 1. Determination of CKX activity
see supplement 1, p. 49 and supplement 4, p. 80
3. 4. 2. Determination of β−glucosidase activity
see supplement 1, p. 50
3. 4. 3. Determination of ODC, ADC activity
see supplement 2, p. 57
19
3. 4. 4. Determination of DAO activity
see supplement 2, p. 57
3. 5. Microscopic method
3. 5. 1. Transmission electron microscopy
see supplement 3, p. 66
3. 6. Molecular methods
3. 6. 1. RNA extraction and reverse transcription
see supplement 4, p. 79
3. 6. 2. qRT-PCR
see supplement 4, p. 79
3. 7. Determination of relative water content
see supplement 4, p. 79
20
4. Results and Discussion
4. 1. Diurnal rhythms of cytokinins, auxin and abscisic acid
Plants growth and development, as well as their interactions with the environment are
mediated by plant hormones. One of the most important environmental factors is light. In
order to cope with regular rhythm of light/dark periods, plants have evolved circadian
regulatory system, endogenous mechanism that allows organisms to correlate their
physiological activities with predictable day/night cycles. Circadian rhythms are the subset of
biological rhythms with period defined as the time to complete one cycle of ~ 24 h (Dunlap et
al. 2004). These rhythms have been found in a wide range of organisms from plants to
mammals, indicating its importance in life processes, including germination, developmental
processes as hypocotyl elongation and flowering, stomatal opening and gas exchange,
photosynthesis and control of the contents of various metabolites. Also many enzymes like
glutamate dehydrogenase, nitrate reductase, sucrose-phosphate synthase are liable to circadian
and/or diurnal rhythms.
Several studies on circadian rhythms were done using the model plant Arabidopsis thaliana
(Alabadi et al. 2001, Mizoguchi et al. 2005, Nakamichi et al. 2009), followed by soybean
plants (Rufty et al. 1983), tomato (Tucker et al. 2004, Mizoguchi et al. 2007), maize
(Bowsher et al. 1991), tobacco (Masclaux-Daubresse et al. 2002) and barley (Kurapov et al.
2000).
In presented work, we focused on elucidation of diurnal rhythms of regulations of plant
hormone levels, including mechanisms governing their metabolism. Analysis of CKs, auxin
(IAA), ABA (publication 1) and polyamines (publication 2) was performed using leaves of
tobacco plants (Nicotiana tabacum L. cv. Wisconsin 38) grown under 16/8h photoperiod.
The main peak of physiologically active CKs, including trans-zeatin, isopentenyladenine,
dihydrozeatin and their ribosides,was detected 1 h after the middle of the light period (Fig. 1,
p. 50). This finding is in agreement with data of Kurapov et al. (2000), who measured diurnal
variation of phytohormones in barley leaves. The midday peak of CKs was accompanied by
the maximum level of IAA (Fig. 6, p. 51). Since IAA and CKs are necessary for the
progression of cell division, these results indicate that cell division may be initiated at this
time, as demonstrated Kurapov et al. (2000), where the maximal plant growth coincided with
time right after the midday peak of CKs and IAA, whereas the minimal growth was measured
right before dawn. The behaviour of cis-zeatin and its riboside was similar to the time course
21
of active CKs, the minor peak coincided with the maximum of active CKs, whereas the major
peak detected 3 h after light/dark transition had 2 h delay in relation to the bioactive CKs.
We also measured concentration of deactivation metabolites of CKs (N-glucosides, Fig. 2, p.
50) with two peaks coinciding with active CKs. This result seems to indicate precise control
of the content of bioactive CKs.
Another way of regulation of CK levels is their conversion to storage forms (O-glucosides,
Fig. 3, p. 51). They reached the maximum 1 h after the dark/light transition, then the
concentration started declining to exhibit another minor peak 1 h before the middle of the
light period. These data correspond well with time course of bioactive CKs - while levels of
O-glucosides were decreasing probably due to the activity of β-glucosidase, concentration of
bioactive CKs started rising to reach the maximum after the vehement decline of Oglucosides, shortly after the middle of the light period. To evaluate the role of reversible
conjugation in regulation of CK levels, β-glucosidase activity was measured (Fig. 3, p. 51).
Right after the light was switched on, enzyme activity sharply increased, which might
coincide enhanced transport of CKs (especially of CK O-glucosides) from the roots by
transpiration stream. The occurrence of β-glucosidase peak right before and after the
maximum of CKs might indicate that hydrolysis of CK O-glucosides could be involved in the
fine-tuning of bioactive CK concentration at midday, even if this deglucosylation could not be
the only source of the midday peak of bioactive CKs. Elevation of CK phosphates, the
immediate CK biosynthesis precursors indicates that CK biosynthesis is likely to play the
crucial role in sharp midday increase of bioactive CKs.
As determination of total β-glucosidase activity was done, hydrolysis of other
glucoconjugates (e.g., ABA-glucosylester) should be also taken into consideration.
Measuring ABA levels (Fig. 6, p. 51), we found one minor peak following the middle day
maximum of IAA and bioactive CKs, where ABA probably serves to counterbalance the
levels of bioactive CKs to regulate the stomatal aperture by CK/ABA ratio (Fusseder et al.
1992). The sharp maximum detected at the beginning of the dark phase was consistent with
the data of Tallman (2004), who reported a stimulation of ABA biosynthesis at dark, to keep
the stomata closed during the night to ensure maximal tissue rehydration. The decrease in
ABA concentration found after the beginning of the light period is probably associated with
the exhaustion of endogenous ABA and the activation of its catabolism (Tallman 2004,
Kraepiel et al. 1994). The decline in ABA concentration in the guard cells and surrounding
apoplast in the morning allows stomatal opening with possitive effects on photosynthesis.
22
As the roots are the main site of CK biosynthesis, it was suggested that high concentration of
xylem CKs in the morning originated from the steady rate of CK biosynthesis in the roots and
of their exudation into the xylem sap during the night, when the transpiration rate is low
(Beck and Wagner, 1994). In the morning, the accumulated CKs are delivered throughout the
plant as the flow of the xylem sap increases. Due to the enhanced volume of the transpiration
stream during the day and the steady rate of CK biosynthesis in roots, CK concentration in the
xylem sap gradually decreases. The importance of this process in translocation of rootsynthetised CKs was recently shown by Aloni et al. (2005). When the activity of the main CK
degrading enzyme was followed, the main peak of CKX activity (Fig. 4, p. 51) occurred early
in the light period. The timing of the peak might coincide with enhanced CK delivery in the
morning by increased xylem flow. Other two minor CKX peaks (detected after midday and
after light/dark transition) coincided well with the decrease of endogenous CKs.
Another approach in the study of CK diurnal variations was the use of radiolabelled transzeatin to follow its metabolic conversion (Fig. 5, p. 51). We found a sharp maximum of
adenine and adenosine, which are products of CK breakdown, right after the dark/light
transition and one minor peak around 9 h of light. Both of them preceded the maxima of CKX
activity. Also the peak of non-metabolized trans-zeatin around 6 h of light corresponds well
with the period od the diminished trans-zeatin breakdown as well as with the minimum of the
ribosylated and phosphoribosylated forms.
4. 2. Diurnal rhythms of polyamines
Our next paper was focused on diurnal patterns of polyamines and their metabolic enzymes
(publication 2). Polyamines play an imporatnt role in the regulation of different processes in
plants due to their cationic nature, which enables interaction with anionic biomolecules, such
as nucleic acids, proteins or phospholipids.
They are also known to affect many processes regulated by CKs and auxins, which is the
reason for the study of diurnal rhythms of CKs, auxins and PAs simultaneously.
The results presented in the publication 2 show detailed changes in the contents both free and
conjugated (soluble and insoluble) forms of Put, Spd, Spm and Cad in tobacco leaves grown
under the 16/8 h light/dark period, as well as in the activities of main enzymes participating in
PA metabolism as ADC, ODC and diamine oxidase (DAO).
The ratio among PA forms was changing during the 24 h cycle (Fig. 2, p. 58). Whereas the
most abundant PA in free and PCA-soluble form was Put and the least abundant Cad, in PCAinsoluble conjugates predominated Cad before Put, Spm, Spd (Fig. 3, p. 58). The endogenous
23
levels of free PAs can be regulated also by conjugation with hydroxycinnamic acid (MartinTanguy 1985, Creus et al. 1991, Bagni and Tassoni 2001, Biondi et al. 2001), which seems to
be the common phenomena in Nicotiana (Martin-Tanguy 1985, Pfosser et al. 1990,
Gemperlová et al. 2005), because about 75 % of the total Put and Spd pool detected in the
midday peak was presented in PCA-soluble conjugates. Nevertheless, the biological function
of conjugated PAs remains unclear, speculating the reversible conjugates may supply the cells
with an additional amine reserve (Bonneau et al. 1994) and/or protect cells from harmful
effects of intracellular accumulation of free PAs on the maintenance of cellular pH, ion
homeostasis and on many other processes influenced by PAs.
The time-course of the level of PCA-soluble conjugates of Put and Spd was quite similar (Fig.
2A, B, p. 58) showing the first maximum 1 h after the middle of the light period. Further
transient peaks were detected 2 h before the light was switched off and at the beginning of the
dark period. A minor peak was observed at the end of the dark phase. It is very interesting but
not surprising that the observed midday maximum of PAs coincided well with the main peak
of CKs and IAA, as was shown in our previous work (Fig. 6, p. 51). These results only
confirm our suggestion that cell division occurs at this time period. In tobacco leaves grown at
the 16/8 h light/dark period, the level of free and PCA-soluble PAs seems to be regulated
especially by the activities of biosynthetic enzyme ODC and catabolic enzyme DAO. Light
dramatically stimulated activity of ODC (Fig. 5, p. 59 ), whereas no increase in activity of the
second biosynthetic enzyme ADC was observed at the beginning of the light phase. On the
contrary, the enzyme activity declined during the first hours of light and started increasing at
midday, with main peak at the light/dark transition. This peak was accompanied by the
accumulation of free PAs and PCA-soluble conjugates and also by the increase in ABA levels
(Nováková et al. 2005). Those results seems to be in accordance with findings of Koetje et al.
(1993) and Galston et al. (1997) who suggested that the pathway via ADC is generally linked
to stress responses and morphogenetic processes, whereas ODC is involved in the regulation
of cell division in actively growing plant tissue.
Activity of DAO, the catabolic enzyme, was induced by light in the similar manner as ODC
activity. We measured a sharp maximum 1 h after the dark/light transition, which may be the
reason why only a slight increase in levels of free Put was observed after the photoinduction
of ODC activity at the beginning of light phase. The second peak measured shortly after the
middle of the light period is in agreement with results of Andreadakis and Kotzabasis (1996)
who observed maximum of enzyme activity in isolated etioplasts from Hordeum vulgare after
light exposure for 9 h.
24
We must also remark that according to information from literature, catabolic degradation of
Put by DAO is not only a way of regulation of cellular content of this diamine (Kumar et al.
1997, Bagni and Tassoni 2001), as the other reaction products, like pyrroline and hydrogen
peroxide, play important roles in subsequent processes. Pyrroline can be further catabolised to
γ-aminobutyric acid (GABA) and finally incorporated into the Krebs cycle (Tiburcio et al.
1997) and/or function in signal transduction pathways during stress response (Shelp et al.
1999). Hydrogen peroxide might trigger a programmed cell death (Walters 2003) or might be
utilised in lignification both during the normal growth and stress response (Angelini et al.
1993).
4. 3. The study of chloroplast autonomy in regulation of CK levels
To study the impact of manipulation of CK metabolism on the endogenous CK pool in leaves
and isolated intact chloroplast, and on chloroplast ultrastructure in tobacco (Nicotiana
tabacum L.) we used plants overproducing CKs (plants were expressing the Sho gene coding
for a Petunia hybrida IPT), plants with decreased CK levels (overexpressing cytokinin
oxidase/dehydrogenase AtCKX3 from Arabidopsis thaliana), and plants with altered CK
glucoconjugation (overexpressing maize β-glucosidase Zm-p60.1 or trans-zeatin Oglucosyltransferase ZOG1 from Phaseolus lunatus).
The total CK content in leaves after Sho induction was eight times higher than in the
corresponding WT (SR1) (Fig. 2, p. 67) and the most elevated CK metabolites were
phosphates and N-glucosides. The differences between Sho transformants and the WT was
much lower in isolated chloroplasts than in leaves. Following Sho induction in leaves, major
increase was observed in iP7G and iPMP, which is in line with previous results reported for
petunia and tobacco constitutively overexpressing Sho (Zubko et al. 2002). The CK content
was also increased in chloroplasts but to a relatively lower extent. Recent analysis of Pssu-ipt
tobacco (Synková et al. 2006) showed that, similarly to Sho plants, CK levels increased in
chloroplasts to much lower levels compared with leaf samples, indicating that chloroplasts are
relatively autonomous and keep their CK homeostasis quite well in both Z- and iP-type CKoverproducing plants. The content of IAA in leaves of Sho-expressing plants did not differ
significantly from that in WT, whereas chloroplasts contained a lower level of IAA than WT
chloroplasts (Table 1, p. 68). A higher ABA level was found in chloroplasts and leaves (when
expressed per FW) of Sho plants compared with WT (Table 1, p. 68).
It was shown that constitutive overexpression of AtCKX3 significantly delayed development
of transgenic plants (Werner et al. 2001, 2003). Retarded ontogenesis was the reason why the
25
hormone content and chloroplast structure of transgenic plants were analysed in older plants
than corresponding WT (SNN) plants. The total CK content was markedly reduced in leaves
(Fig. 3, p. 68), particularly levels of N-glucosides and to a lesser extent bases and ribosides. It
is obvious that after a tremendous increase of the activity of CKX, the activity of the other CK
deactivation pathway (N-glucosylation) was suppressed. No significant differences were
found between the IAA levels (expressed on FW basis), in leaves of AtCKX3 and WT plants
(Table 2, p. 69). A lower ABA content was observed in AtCKX3 leaves compared with WT
leaves, which might be associated with prolonged life span and retarded senescence of
transgenic plants (Table 2, p. 69).
Maize β-glucosidase Zm-p60.1 was the first plant enzyme shown to release active CKs from
CK metabolites O- and N3-glucosides in vivo (Brzobohatý et al. 1993). In tobacco plants
overexpressing maize β-glucosidase Zm-p60.1 targeted to plastids, we observed an increase in
CK bases and ribosides and CK N-glucosides in leaves (Fig. 4A,B, p. 69 ). These results are
in accordance with recent data by Kiran et al. (2006). Accumulation of CK bases and
ribosides was found also in chloroplasts (Fig. 4C, p. 69). The very dramatic decrease in CK
O-glucosides levels shown here for chloroplasts isolated from Zm-p60.1 plants corresponded
well with plastid/chloroplast location of Zm-p60.1 (Kristoffersen et al. 2000). Measurements
of IAA levels revealed a significantly lower concentration of IAA both in leaves and
chloroplasts of transgenic tobacco plants compared with the WT IAA level (Table 3, p. 70),
which is in agreement with Kiran et al. (2006), who measured a fall in IAA gradient from the
youngest leaves downward and a decreased IAA content in the apex and first internodes in
Zm-p60.1 tobacco plants. In ABA levels, we observed lower levels in leaves and chloroplasts
of transgenic plants than in WT, which is in contrast with previous work of Kiran et al.
(2006), who found higher ABA accumulation especially in leaves of Zm-p60.1 tobacco plants.
Tobacco transformed with ZOG1 gene from Phaseolus lunatus under the control of the CaMV
35S promoter was constructed by Martin et al. (2001a). When a ZOG1 gene was
constitutively expressed in tobacco, CK O-glucosides accumulated in leaves (Fig. 5A, p. 70).
No significant difference in other CK metabolites was observed. By contrast, in chloroplasts a
very low levels of CK O-glucosides were found (Fig. 5C, p. 70). This indicates that CK Oglucosides are accumulated mainly out of the plastids, which is in agreement with the
previous data that transgenic plants overexpressing Zm-p60.1 can accumulate high levels of tZOG when grown on a medium supplemented with t-Z despite high Zm-p60.1 β-glucosidase
activity in chloroplasts (Kiran et al. 2006). As the main storage compartment for CK O-
26
glucosides was proposed vacuole (Fusseder and Ziegler 1988). In leaves of transgenic plants,
a 2-fold higher IAA level was measured compared with WT (Table 4, p. 71), whereas the
ABA content was not significantly affected. The IAA and ABA levels in chloroplasts were
higher in ZOG1 plants than in WT.
A significant changes were observed in the chloroplast ultrastructure of tobacco plants
constitutively expressing the Sho gene, including an increased number of plastoglobuli and
the appearance of crystalloids in plants grown in vitro (Fig. 6, p. 71). Except for Sho tobacco
transformants, marked changes in the ultrastructure of chloroplasts were not found in the
other CK transformants analysed. The most obvious feature observed was the different starch
accumulation associated with a change in chloroplast shape (Fig. 7, p. 72) which is in
accordance with Wilhelmová and Kutík (1995) and Stoynova et al. (1996).
All these results indicate that chloroplasts are relatively independent organelles with respect
to the regulation of CK metabolism and that different compartmentation of individual CK
metabolites should be taken into account.
4. 4. The role of CK in responses to water deficit
Water stress represents one of the main limiting factors of plant growth and development.
Water availability has become one of the most discussed problems. Detailed understanding of
mechanisms of plant response to water deficit is a necessary prerequisite for the elucidation of
an optimal strategy for enhancement of plant drought stress tolerance.
We studied the impact of water deficit progression on CKs, auxin and ABA, using tobacco
plants over-expressing trans-zeatin O-glucosyltransferase gene under 35S or SAG12
promoters and the corresponding wild-type. The SAG12 promoter has been, since the
pioneering work of Gan and Amasino (1995), successfully used for targeted manipulation of
CK levels.
The transformants were characterized by ZOG1 expression profile. The amount of ZOG1
mRNA was quantified in the individual leaves and roots of control (well-watered) and
drought-treated plants by real-time RT-PCR. ZOG1 expression in constitutive transformants
was relatively high and even within the plant. The SAG12 promoter was significantly less
active. As a promoter of senescence associated gene, it exhibited strict dependence on the leaf
position. The highest expression levels of ZOG1 were found in SAG::ZOG1 transformants in
older (lower) leaves (Table 1, p. 81).
For better orientation in results of endogenous CK analysis, we divided individual CKs into
seven groups according to their structure and function. Namely bioactive CKs, CK
27
deactivation forms (N-glucosides), CK storage forms (O-glucosides), CK phosphates (CK
biosynthesis intermediates), cis-zeatin derivatives, cis-N-glucosides and cis-O-glucosides.
In plants not suffering from water deficiency, the concentration of bioactive CKs is usually
quite high in the young leaves and declines as the leaves start senescing (e.g. Singh et al.
1992). In accordance with these results, we observed in young plants at the beginning of the
experiment a similar content of bioactive CKs in all leaves. With stress induced senescence
progression, a gradient of bioactive CKs in favour of upper leaves was observed (Fig. 5, p.
82).
In senescing well-watered SAG12::ZOG1 plants, we found an elevation of CK content,
especially in lower leaves, which is in accordance with Cowan et al. (2005), who reported an
increase in trans-zeatin-type CKs in senescing well-watered SAG12::ipt plants. After
stimulation of SAG12::ZOG1 expression, we observed, apart of the expected increase of the
level of CK O-glucosides, also an elevation of bioactive CKs. Both control and stressed
35S::ZOG1 transformants had considerably higher total CK levels than WT (Fig. 4, p. 82),
predominantly of O-glucosides, both of the trans- and cis-zeatin type. As the ZOG1
expressing plants have elevated pool of CK O-glucosides, we compared the β-glucosidase
activity in WT and ZOG1 transformants to check the ability of these plants to utilise the
elevated levels of CK O-glucosides. Mild elevation of β-glucosidase activity in ZOG1 plants
(results not shown) seemed to prove their capacity to hydrolase the CK storage pool.
In order to elucidate CK function during the drought stress, we compared the water deficit
response of WT plants, those with a uniform elevation of CK content before stress initiation
and those exhibiting CK elevation targeted to senescing tissues.
The mild water stress caused in WT plants formation of a gradient of bioactive CKs in favour
of the upper leaves (Fig. 5, p. 82). This observation reflects necessity for the preferential
protection of upper leaves under adverse conditions. The importance of CKs in sink-source
polarisation during mild stress was recently discussed by Cowan et al. (2005). SAG::ZOG1
exhibited similar CK response as WT, however, the content of bioactive CKs was in lower
leaves less suppressed by the drought. During the prolonged drought, decrease in bioactive
CK levels was in 35S::ZOG1 plants more profound than in WT. The elevated CK content
before stress initiation might be the reason of the delay in the response of 35S::ZOG1 plants
to unfavourable environmental conditions, which might have negative consequence in case of
severe, prolonged drought.
The ABA content of plants correlated well with the strength of applied water stress (Fig. 8, p.
85). When mild stress was applied, only a slight increase in ABA concentration occurred in
28
the upper and middle leaves of WT plants and in the middle leaves of SAG::ZOG1 plants. In
constitutive ZOG1 transformants we observed no increase of ABA levels. Under mild stress,
an elevated level of free IAA was found in WT lower leaves (Fig. 9, p. 85), whereas in roots
of both transgenics we observed slight increase of IAA levels. In all genotypes, the CKX
activity was much higher in roots than in leaves (Fig. 7, p. 84).
Moderate drought stress had two main effects on the concentrations of bioactive CKs. Firstly,
the progressive drought led to a decrease of the bioactive CK levels of (especially of transzeatin/riboside) in all leaves of all genotypes tested, which is in agreement with Goicoechea
et al. (1995). Secondly, the steepness of the apex-to-base gradient in bioactive CKs increased
substantially, especially in 35S::ZOG1 plants, in which only minor response to mild stress
was observed. The relatively small difference in bioactive CKs levels between the middle and
lower leaves of SAG::ZOG1 plants might be the consequence of enhanced SAG promoter
activity in stressed lower leaves (Fig. 3, p. 81). Under moderate drought stress conditions,
ABA levels were increasing substantially in all stressed plants (Fig. 8, p. 85), which is in
agreement with previous reports (e.g. Mansfield and McAinsh 1995, Riccardi et al. 2004). As
ABA is an important component of the plant stress defence system, its higher levels, observed
in the upper leaves, might reflect their preferential protection. As far as IAA is considered,
moderate drought stress resulted in all genotypes in a substantial increase in IAA levels, both
in the lower leaves and roots (Fig. 9, p. 85).
Severely stressed plants had all the leaves completely wilty. The bioactive CK content
decreased in all genotypes, compared with that in moderate stress and CK gradient was
suppressed in the WT and SAG::ZOG1 plants, while it was still visible in 35S::ZOG1 plants.
The flattening of the CK gradient might indicate that the strength of the stress nearly reached
the limits of the plant's ability to cope with the drought. Nevertheless, the decrease of CKX
activity in the upper leaves and, especially, the large increase in CKX activity in the lower
leaves seems to indicate a tendency for the maintenance of the gradient of bioactive CKs (Fig.
7, p. 84). In all genotypes tested, severe stress led to the further accumulation of total CKs in
the roots.
Severe stress also led to a further increase in ABA content (by more than one order magnitude
in comparison with control conditions). The highest concentration occurred in the upper
leaves (Fig. 8, p. 85).
Large elevation of IAA content, observed in the stressed lower leaves of all genotypes and
also in the severely stressed middle leaves of 35S::ZOG1 plants (Fig. 9, p. 85), is in the
agreement with the results of Quirino et al. (1999), who found an increase in the free IAA
29
levels in senescing Arabidopsis leaves. The function of the increased IAA levels in senescing
leaves is not clear. As senescence is a strongly regulated process, high levels of IAA might be
necessary for its control. A possible explanation could be that the increased protein
breakdown in senescing leaves leads to an increase in the content of tryptophan, which may
be converted to IAA.
After re-watering, a decrease in the total CK content was observed in the roots in all tested
genotypes. Simultaneously, the level of bioactive CKs in the leaves increased significantly in
comparison with severely stressed plants (Fig. 5, p. 82 ), which might be, at least partially,
result of the re-establishment of xylem sap flow from the roots. The importance of the
transpiration stream in CK transport from the roots was recently demonstrated by Aloni et al.
(2006). After rehydration, the stress indicators (SAG12 promoter activity and ABA levels)
sharply decreased (Table 1, p. 81 and Fig. 8, p. 85).
The slightly faster growth initiation in SAG::ZOG1 plants in comparison with WT is in
agreement with the reported positive effect of CKs on plant recovery (e.g. Itai et al. 1978).
The slower recovery in the case of 35S::ZOG1 plants might indicates that elevated CK levels
need not be advantageous in case of prolonged, severe drought stress, which is in accordance
with Synková et al. (1999).
To conclude, data on SAG12 promoter activity demonstrated that drought strongly accelerated
senescence of tobacco plants, which coincided with the gradient of bioactive CKs in favour of
the upper leaves. The gradient was maintained by the stimulation of CKX activity in the lower
leaves and by its suppression in the upper leaves. Targeted, senescence-induced elevation of
ZOG1 expression exhibited a positive effect on both the delay of senescence of the stressed
lower leaves and the speeding up of the recovery.
30
5. Summary
➔ Cytokinins, auxin, abscisic acid and polyamines levels undergo significant diurnal
variations in leaves. These rhythms have physiological relevance and correlate with
circadian changes of plant growth and development.
➔ The activity of cytokinin oxidase/dehydrogenase and cytokinin N- and O-glucosylation
are also undergoing diurnal changes.
➔ Arginine decarboxylase and ornithine decarboxylase activities, as well as putrescine
oxidizing activity, correlates well with the alternations in polyamine content during the
day/night period.
➔ Highly non-uniform distribution of cytokinins found between chloroplasts and bulk
leaf tissue in different transgenic tobacco lines with modulated cytokinin content indicates
that chloroplasts are relatively independent organelles with respect to the regulation of
cytokinin metabolism.
➔ Data on SAG12 promoter activity demonstrated that drought strongly accelerated the
senescence of tobacco plants, which coincided with the establishment of the gradient of
bioactive cytokinins in favour of the upper leaves.
➔ Targeted, senescence-induced elevation of expression of gene for trans-zeatin Oglucosyltransferase exhibited a positive effect both on the delay of senescence of the
stressed lower leaves and during the recovery.
➔ Uniform elevation of cytokinin content in constitutive 35S::ZOG1 transformants
before stress initiation might have a negative effect during the prolonged, severe drought.
31
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Journal of Experimental Botany, Vol. 56, No. 421, pp. 2877–2883, November 2005
doi:10.1093/jxb/eri282 Advance Access publication 12 September, 2005
This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details)
RESEARCH PAPER
Diurnal variation of cytokinin, auxin and abscisic acid
levels in tobacco leaves
Marie Nováková1,2, Václav Motyka1, Petre I. Dobrev1, Jiřı́ Malbeck1, Alena Gaudinová1
and Radomı́ra Vanková1,*
1
Institute of Experimental Botany AS CR, Rozvojova 135, 165 02 Prague 6, Czech Republic
2
Department of Biochemistry, Faculty of Science, Charles University, Hlavova 2030/8, 128 43 Prague 2,
Czech Republic
Received 15 April 2005; Accepted 5 August 2005
Abstract
As many processes are regulated by both light and
plant hormones, evaluation of diurnal variations of
their levels may contribute to the elucidation of the
complex network of light and hormone signal transduction pathways. Diurnal variation of cytokinin, auxin,
and abscisic acid levels was tested in tobacco leaves
(Nicotiana tabacum L. cv. Wisconsin 38) grown under
a 16/8 h photoperiod. The main peak of physiologically
active cytokinins (cytokinin bases and ribosides) was
found after 9 h of light, i.e. 1 h after the middle of the
light period. This peak coincided with the major auxin
peak and was closely followed by a minor peak of
abscisic acid. Free abscisic acid started to increase at
the light/dark transition and reached its maximum 3 h
after dark initiation. The content of total cytokinins
(mainly N-glucosides, followed by cis-zeatin derivatives and nucleotides) exhibited the main peak after
9 h of light and the minor peak after the transition to
darkness. The main, midday peak of active cytokinins
was preceded by a period of minimal metabolic conversion of tritiated trans-zeatin (less than 30%). The
major cytokinin-degrading enzyme, cytokinin oxidase/
dehydrogenase (EC 1.5.99.12), exhibited maximal activity after the dark/light transition and during the
diminishing of the midday cytokinin peak. The former
peak might be connected with the elimination of the
long-distance cytokinin signal. These cytokinin oxidase/dehydrogenase peaks were accompanied by increased activity of b-glucosidase (EC 3.2.1.21), which
might be involved in the hydrolysis of cytokinin
O-glucosides and/or in fine-tuning of active cytokinin levels at their midday peak. The achieved data indicate that cytokinin metabolism is tightly regulated
by the circadian clock.
Key words: Abscisic acid, auxin, cytokinin, cytokinin oxidase/
dehydrogenase, diurnal rhythm, b-glucosidase, tobacco.
Introduction
It is assumed that plant growth and development are
regulated by the interactions between the environment
and endogenous factors, especially hormones. Light is an
important environmental signal that activates and modifies
various growth and developmental processes in plants, for
example, seed germination, flower induction, leaf movements, stem elongation, and de-etiolation. All of these
processes are regulated by plant hormones as well. Effects
of light and phytohormones can be synergistic (e.g.
chloroplast development and cytokinins [CKs]), additive
(e.g. inhibition of hypocotyls elongation and CKs), or
antagonistic (e.g. germination and abscisic acid [ABA]).
There are numerous reports that light itself may influence
the levels of different hormones. Light was found to
regulate directly the biosynthesis of active gibberellins
(Foster and Morgan, 1995) and ethylene (Jasoni et al.,
2002) as well as to promote the degradation of ABA
(Kraepiel et al., 1994). The effect of light on auxin levels is
less transparent and may be realized by influencing auxin
* To whom correspondence should be addressed. Fax: +420 220 390 446. E-mail: [email protected]
Abbreviations: ABA, abscisic acid; CK, cytokinin; CKX, cytokinin oxidase/dehydrogenase; DHZ, dihydrozeatin; DHZR, dihydrozeatin riboside; IAA, b-indolylacetic acid; iP, N 6-(D2-isopentenyl)adenine; iPR, N 6-(D2-isopentenyl)adenosine; iP7G, N6-(D2-isopentenyl)adenine 7-glucoside; iP9G, N6-(D2-isopentenyl)
adenine 9-glucoside; Z, trans-zeatin; ZR, trans-zeatin riboside; ZOG, trans-zeatin O-glucoside; ZROG, trans-zeatin riboside O-glucoside; Z7G, trans-zeatin
7-glucoside; Z9G, trans-zeatin 9-glucoside.
ª The Author [2005]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
The online version of this article has been published under an Open Access model. Users are entitled to use, reproduce, disseminate, or display the Open Access version of this article
for non-commercial purposes provided that: the original authorship is properly and fully attributed; the Journal and the Society for Experimental Biology are attributed as the original
place of publication with the correct citation details given; if an article is subsequently reproduced or disseminated not in its entirety but only in part or as a derivative work this must be
clearly indicated. For commercial re-use, please contact: [email protected]
48
2878 Nováková et al.
polar transport (Tian and Reed, 2001) or by the modulation
of cell sensitivity towards auxin (Jones et al., 1991).
The data on the effect of light on the metabolism of
cytokinins are rather scarce. The assumption of a positive
effect of light on CK accumulation has been based on the
decreased level of trans-zeatin in phytochrome mutants
(Kraepiel et al., 1995). Diurnal variation of CK levels was
followed in barley (Kurapov et al., 2000) and carrot
(Paasch et al., 1997), but these studies were limited only
to the determination of CK bases and ribosides. The recent
development of analytical methods (especially of mass
spectrometry) has enabled the study of the diurnal variation
of a broad array of CK metabolites.
Diurnal changes in the levels of 27 CK metabolites were
studied in tobacco leaves. The potential contribution of the
cleavage of CK O-glucoconjugates to the fast changes of
the level of physiologically active CKs (CK bases and
ribosides) was estimated via the activity of b-glucosidase
(EC 3.2.1.21). In order to study CK catabolism, the activity
of the main CK-degrading enzyme, cytokinin oxidase/
dehydrogenase (CKX, EC 1.5.99.12) was determined. As
CK levels are also affected by other hormones, the diurnal
variation of auxin and ABA levels was followed.
USA) and Oasis MCX mixed mode (cation exchange and reversephase) columns (150 mg, Waters, USA) and evaporated.
HPLC of radiolabelled trans-zeatin and its metabolic products
Evaporated samples were resuspended in 200 mm3 20% (v/v)
acetonitrile. Radiolabelled metabolites of [3H]Z were analysed
according to Blagoeva et al. (2004) using a Series 200 Quaternary
HPLC Pump (Perkin Elmer, Wellesley, MA, USA) coupled to a
235C Diode Array Detector (Perkin Elmer, USA) and a RAMONA
2000 flow-through radioactivity detector (Raytest, Straubenhardt,
Germany), column: Luna C18(2), 150 mm/4.6 mm/3 lm (Phenomenex, Torrance, CA, USA). The radioactive metabolites were identified on the basis of coincidence of retention times with authentic
standards.
HPLC of auxin and abscisic acid
Indolyl acetic acid and ABA were determined using two-dimensional
HPLC according to Dobrev et al. (2005). The instrumental set-up
consisted of a series 200 autosampler (Perkin Elmer, Norwalk, CT,
USA), two HPLC gradient pump systems (first pump: ConstaMetric
3500 and 3200 with 500 ll mixer, TSP, Riviera Beach, FL, USA;
second pump: Series 200 Quaternary Pump, Perkin Elmer), two
columns (first column: ACE-3CN, 15034.6 mm, 3 mm, ACT,
Aberdeen, Scotland, UK; second column: Luna C18(2), 15034.6
mm, 3 mm, Phenomenex, Torrance, CA, USA), one 2-position, fluid
processor SelectPRO with 1 ml loop (Alltech, Deerfield, IL, USA).
The segment containing IAA and ABA obtained in the first dimension was collected in the loop of the fluid processor and
redirected to the second HPLC dimension. IAA was quantified
using fluorescence detector LC 240 (Perkin Elmer). Quantification
of ABA was performed on the basis of UV detection using diode
array-detector 235C (Perkin Elmer).
Materials and methods
Plant material
Tobacco plants (Nicotiana tabacum L. cv. Wisconsin 38) were grown
in soil in a growth chamber for 6 weeks at a 16 h photoperiod and
130 lmol mÿ2 sÿ1, a day/night temperature of 25/23 8C, and a relative
humidity of c. 80%. Mixed samples of leaves (3rd–5th of 7, i.e. with
the exception of the youngest and oldest ones) from three plants were
collected every hour during a 24 h period. After removal of the main
vein the samples were immediately frozen in liquid nitrogen.
HPLC/mass spectrometry
LC-MS analysis was performed as described by Lexa et al. (2003)
using a Rheos 2000 HPLC gradient pump (Flux Instruments,
Switzerland) and HTS PAL autosampler (CTC Analytics, Switzerland) coupled to an Ion Trap Mass Spectrometer Finnigan MAT
LCQ-MSn equipped with an electrospray interface. Detection and
quantification were carried out using a Finnigan LCQ operated in the
positive ion, full-scan MS/MS mode using a multilevel calibration
graph with deuterated cytokinins as internal standards. The levels of
27 different cytokinin derivatives were measured. The detection limit
was calculated for each compound as 3.3 r/S, where r is the standard
deviation of the response and S the slope of the calibration curve.
Three independent experiments were done. Each sample was injected
at least twice.
Incubation with tritiated trans-zeatin
Tip parts of the 3rd or 4th leaf (c. 1 g FW) were cut at 3 h intervals and
their cut surface was submerged into aqueous tritiated trans-zeatin
([3H]Z, 33104 Bq, specific activity 0.6 TBq mmolÿ1, prepared by
Dr Jan Hanuš, Isotope Laboratory, Institute of Experimental Botany
AS CR, Prague, Czech Republic). After 1 h incubation (under the
same light regime as the intact plants) leaf tips were frozen in liquid
nitrogen. The uptake of [3H]Z by the vascular tissue was efficient
(35–50% of the applied radioactivity).
Determination of CKX activity
The enzyme preparations were extracted and partially purified using
the method of Chatfield and Armstrong (1986) as modified by
Motyka et al. (2003). The CKX activity was determined by in vitro
assays based on the conversion of [2-3H]isopentenyladenine (prepared by Dr Jan Hanuš) to [3H]adenine. The assay mixture (50 mm3
final volume) included 100 mM TAPS-NaOH buffer containing
75 lM 2,6-dichloroindophenol (pH 8.5), 2 lM substrate ([2-3H]
isopentenyladenine, 7.4 Bq molÿ1) and the enzyme preparation
equivalent to 250 mg tissue FW (corresponding to 0.02–0.26 mg
protein gÿ1 FW). After incubation at 37 8C the reaction was
terminated by the addition of 10 mm3 Na4EDTA (200 mM) and
120 mm3 95% (v/v) ethanol. Separation of the substrate from the
product of the enzyme reaction was achieved by the same HPLC
system as described for tritiated trans-zeatin metabolites according
to Blagoeva et al. (2004). The CKX activity was determined in two
independent experiments. As the data from both experiments showed
Plant hormone extraction and purification
Phytohormones were extracted and purified according to Dobrev and
Kamı́nek (2002). Frozen leaf samples (c. 1 g FW) were ground in
liquid nitrogen and extracted overnight with 10 cm3 methanol/water/
formic acid (15/4/1, by vol., pH;2.5, ÿ20 8C). For analyses of
endogenous CKs, 50 pmol of each of the following 12 deuteriumlabelled standards were added: [2H5]Z, [2H5]ZR, [2H5]Z7G,
[2H5]Z9G, [2H5]ZOG, [2H5]ZROG, [2H6]iP, [2H6]iPR, [2H6]iP7G,
[2H6]iP9G, [2H5]DHZ, [2H5]DHZR (Apex Organics, Honington,
UK). Tritiated internal standards were used for the determination of
auxin (3[5(n)-3H] indolyl acetic acid, Amersham, UK, specific
activity 74 GBq mmolÿ1, 53103 Bq) and ABA (Amersham, UK,
specific activity 1.74 TBq mmolÿ1, 53103 Bq). The extracts were
purified using Si-C18 columns (SepPak Plus, Waters, Milford, MC,
49
Cytokinin, auxin, and ABA diurnal rhythms
2879
the same tendencies (but different absolute values), the results of
one representative experiment (mean value of three parallel assays for
each of three replicate protein preparations) are presented.
Determination of b-glucosidase activity
Tobacco leaves (0.2 g FW) were ground in liquid nitrogen and
transferred into 0.4 cm3 ice-cold extraction buffer [25 mM TRIS-Cl,
pH 7.5, 5 mM MgCl2, 1 mM EDTA, and protease Complete EDTAfree cocktail tablet (Roche)]. Samples were centrifuged at 13 000 g
for 15 min (4 8C) and 0.02 cm3 supernatant were added into 0.28 cm3
citrate-phosphate buffer (0.02 M citric acid; 0.06 M NaH2PO4; pH
5.5). The reaction was initiated by 0.1 cm3 20 mM p-nitrophenyl
a-D-glucopyranoside. After 120 min at 30 8C the reaction was
stopped by adding 0.6 cm3 2 M Na2CO3. The released p-nitrophenol
was measured at 405 nm (UNICAM 5625 UV/VIS spectrometer).
Results
Cytokinins
Physiologically active cytokinins, i.e. trans-zeatin, isopentenyladenine, dihydrozeatin, and their corresponding ribosides, exhibited in tobacco leaves a sharp maximum after
9 h of light, i.e. 1 h after the middle of the light period
(Fig. 1). Another, considerably lower, peak was detected
after the transition to darkness and the lowest one was
found at the beginning of the light phase. The content of
cis-zeatin and its riboside exhibited a minor peak coinciding with the maximum of active CKs. Their maximum
followed the second peak of active CKs after the light/
dark transition (with 2 h delay). The content of total CKs
(the sum of physiologically active CKs together with CK
phosphates, cis-zeatin derivatives, N- and O-glucosides)
have an increasing trend during the light period, up to
a maximum 9 h after the light was switched on (i.e. 1 h after
midday). The second half of the light period was characterized by a sharp decrease in CKs. After the light/dark
transition another transient CK peak occurred. During the
course of the dark period, the total CK level gradually
declined. (The levels of individual CKs are given in a table
as supplementary material that can be found at JXB online.)
The concentration of the CK deactivation metabolites
(N-glucosides) increased at the end of the dark period and
remained at an elevated level up to 6 h of light (Fig. 2). The
main maximum of CK N-glucosides coincided with (or
closely followed) the peak of active CKs after the middle
of the light period. Then, a sharp decrease took place.
The second main peak, nearly as large as the first one, was
found after the light was switched off, coinciding with
a minor peak of physiologically active cytokinins. The
level of cis-zeatin derivatives followed a trend similar to
the N-glucosides, with the exception of the second maximum after the light/dark transition, which was, in the case
of cis-zeatin derivatives, delayed and much less profound.
The levels of CK phosphates exhibited a transient peak
around 4 h of light. Their maximum was found after the
middle of the light period.
Fig. 1. Endogenous levels of total cytokinins, physiologically active
cytokinins (trans-zeatin, isopentenyladenine, dihydrozeatin, and the corresponding ribosides) and cis-zeatin (riboside) in tobacco leaves during
a 24 h period. The photoperiod is represented by the black and white box on
the abscissa. Means 6SE of three independent experiments are shown.
Fig. 2. Endogenous levels of CK N-glucosides, CK phosphates, and ciszeatin derivates in tobacco leaves during a 24 h period. Other details as
for Fig. 1.
Cytokinin O-glucosides and activity of b-glucosidase
The level of CK storage forms (O-glucosides) was found
to increase around the dark/light transition. Another peak
of O-glucosides was detected after 7 h of light, preceding
the peak of active CKs as well as that of CK deactivation metabolites. However, this CK O-glucoside peak was
significantly lower than those of the deactivation products.
Minor peaks of CK O-glucosides were detected after 10 h
of light, when the level of active CKs was decreasing, and
2 h after the light/dark transition.
The activity of b-glucosidase, the enzyme that catalyses
the cleavage of glucosyl moieties from O-glucosides releasing, in the case of CK O-glucosides, active CKs,
exhibited the first maximum after 1 h of light, which correlated with the decrease of the peak of CK O-glucosides
found around the dark/light transition (Fig. 3). The major
peak of b-glucosidase activity was found 7 h after the light
was switched on, preceding the midday peak of active CKs.
50
2880 Nováková et al.
Cytokinin oxidase/dehydrogenase
adenosine, products of CK breakdown, was found close to
the dark/light transition and a minor one around 9 h of light,
i.e. at the periods preceding the detected maxima of CKX
activity. The peak of non-metabolized trans-zeatin, corresponding to the period of the diminished trans-zeatin
breakdown, preceded the midday peak of active CKs. The
maximum of the non-metabolized labelled trans-zeatin
correlated with the minimum level of the ribosylated and
phosphoribosylated forms.
A comparison of the diurnal variations in CKX activity and
in the levels of CKs, which are the substrates of this enzyme,
i.e. isopentenyladenine, trans-zeatin, and their corresponding ribosides (but not dihydrozeatin or its derivatives), is
shown in Fig. 4. The main CKX maximum was found 3 h
after the transition from dark to light. The second maximum followed the midday peak of active CKs. The third,
least profound peak of CKX activity accompanied the
minor peak of CKs after the light/dark transition (with
a 1 h delay).
Indolyl acetic acid and abscisic acid
The midday CK peak coincided with the maximum level
of indolyl acetic acid and was accompanied (or closely
followed) by a minor peak of ABA (Fig. 6). No other
profound peak of indolyl acetic acid was detected during
the diurnal cycle. At the end of the light period, a gradual
increase in ABA level was found. This enhancement was
accelerated after the light was switched off, the maximum
Metabolic conversion of radiolabelled trans-zeatin
Diurnal changes in the levels of endogenous CKs and the
activity of CK metabolic enzymes were also evaluated
following the metabolic conversion of radiolabelled
trans-zeatin (Fig. 5). A high peak of labelled adenine and
Fig. 3. Endogenous levels of CK O-glucosides and activity of
b-glucosidase in tobacco leaves during a 24 h period. Other details as
for Fig. 1.
Fig. 5. Metabolism of radiolabelled trans-zeatin in tobacco leaves
during a 24 h period (incubation time 1 h). Other details as for Fig. 1.
Fig. 4. The activity of cytokinin oxidase/dehydrogenase (CKX) and
endogenous levels of cytokinins, which are the substrates of this enzyme
(trans-zeatin, isopentenyladenine, and the corresponding ribosides), in
tobacco leaves during a 24 h period. Other details as for Fig. 1.
Fig. 6. Endogenous levels of indole-3-acetic acid (IAA), abscisic acid
(ABA), and active cytokinins in tobacco leaves during a 24 h period.
Other details as for Fig. 1.
51
Cytokinin, auxin, and ABA diurnal rhythms
being reached after 3 h of dark. The high level of ABA was
only transient. A low, sharp peak of ABA was also detected
immediately after the light was switched on.
Discussion
Timing of the main CK peak in tobacco leaves shortly (1
h) after the middle of the light period is in accordance
with the diurnal profile in barley leaves (main peaks of
CK bases and ribosides around 14.00 h; Kurapov et al.,
2000) and slightly differs from that in carrot plants kept
under continuous light (maximum of CK bases and ribosides around 16.00 h; Paasch et al., 1997). As both CKs and
auxin are necessary for the progression of cell division,
this finding of the coincidence of the main CK peak with
the maximum of IAA indicates that cell division may be
initiated at this time. The close connection between CK
increase and growth stimulation was reported in barley
(at the stage of the third leaf), when the CK peak at 14.00 h
was shown to precede the period of maximal growth from
15.00 h to 18.00 h (Kurapov et al., 2000). The minimal
growth was determined before dawn (at 04.00–05.00 h). A
correlation between growth and the hormone level was
also found in the case of auxin, the link between the oscillation in the level of IAA in the rosette leaves and the stem
growth rate in Arabidopsis was reported by Jouve et al.
(1998, 1999).
The results on the midday peak of IAA are in accordance
with the data of Pavlova and Krekule (1984), who reported
a peak of IAA content after the middle of the light phase in
Chenopodium rubrum plants grown under very long day
(20 h). In shorter periods (16 h, 12 h, 8 h), connected with
flower induction, this peak was shifted towards the dark
phase.
It should be kept in mind that these results reflect the
situation in still developing but not the youngest tobacco
leaves, in other tissues and/or species hormones may
exhibit a different diurnal pattern. The effect of developmental stage was observed in barley. Leaves at the stage of
the third leaf had the main CK peak at 14.00 h, while at the
stage of tillering the peak was shifted to 17.00 h (Kurapov
et al., 2000). Another point is that the experiments were
aimed at the determination of the diurnal profile of isoprenoid CKs. Due to the differences in physiological function, diurnal rhythms of isoprenoid CKs are likely to differ
from those of their aromatic counterparts. The peaks of
aromatic CKs m-methoxytopolin and its riboside were
detected at 09.30 h in poplar leaves (Tarkowska et al.,
2003), which corresponded approximately to 3 h after the
light period initiation, i.e. 6 h earlier than for isoprenoid
CKs in tobacco.
The finding of the ABA maximum at the beginning of
the dark phase is in accordance with the survey of diurnal
changes in ABA metabolism summarized by Tallman (2004),
2881
who reported a stimulation of ABA biosynthesis at dark,
in order to maintain the stomata in the closed position
during the night to ensure maximal tissue rehydration.
The sharp decrease in ABA concentration, which was
found shortly after the beginning of the light period, seems
to be connected with the depletion of endogenous
ABA and the activation of its catabolism. According
to Tallman (2004), after irradiation ABA levels drop
due to the activation of the xanthophyll cycle (conversion
of ABA precursors to zeaxanthin), isomerization into
physiologically inactive trans-forms, as well as to ABA
degradation by cytochrome P450 mono-oxygenase.
Phytochrome-mediated activation of ABA degradation
was also reported by Kraepiel et al. (1994). The decrease
in ABA concentration in the guard cells and surrounding
apoplast in the morning enables stomatal opening, which
positively affects plant photosynthesis.
Later, at the midday phase, elevated apoplastic ABA
around guard cells, delivered by transpiration, is necessary
for the regulation of the stomata aperture in order to balance
the rate of photosynthesis and plant turgor maintenance
(Tallman, 2004). The minor peak of ABA, which was found
1 h after the middle of the light phase, might be related
to this ‘after-midday’ regulation or, more probably, may
serve to counterbalance the midday peak of CKs, as under
temperate conditions stomatal aperture is regulated by the
CK/ABA ratio (Fusseder et al., 1992).
Fusseder et al. (1992) reported that the increase in
xylem-delivered CKs preceded the increase in leaf conductance. They found the peak of xylem CKs in the morning
and their rapid decrease in the afternoon in desert-grown
almond trees. Beck and Wagner (1994) suggested that the
high concentration of xylem CKs in the morning originated
from the steady rate of CK biosynthesis in the roots and of
their exudation into the xylem sap during the night when
the transpiration rate is low. In the morning, the flow of
xylem sap sharply increases, delivering accumulated CKs
throughout the plant. Due to the increased volume of the
transpiration stream during the day and the steady rate of
CK biosynthesis in roots, CK concentration in the xylem
sap gradually decreases. The importance of the velocity
of the transpiration stream on the translocation of CKs
produced in roots to other parts of plant was recently
demonstrated by Aloni et al. (2005). In our experiments
xylem transport of CKs was not measured. Indirect evidence of the existence of such translocation may be given
by the position of the main maximum of CKX activity that
was detected early in the light period. The other peaks of
CKX activity (after midday and following the light/dark
transition) correlated well with the decrease of endogenous
CKs in leaf blades, indicating that CK breakdown by CKX
is involved in the down-regulation of these endogenous CK
peaks. However, the main maximum of CKX activity,
which appeared early in the light phase, did not coincide
with substantial changes of endogenous CK levels in leaf
52
2882 Nováková et al.
Supplementary data
blades, suggesting that this CKX peak may serve in the
regulation of xylem CKs.
To evaluate the role of CK reversible conjugation in
the regulation of their levels, the diurnal rhythm of bglucosidase activity was followed. In the first hours of
the light period, the increase in b-glucosidase activity
created conditions for the cleavage of CK O-glucosides
(one of the CK transport forms) necessary for their further
degradation with CKX. The b-glucosidase increase could,
of course, also result in hydrolysis of other unidentified
compounds, for example, ABA-glucosylester. However, at
this time-point, when the level of free ABA was decreasing
(Tallman, 2004), the function of b-glucosidase activity
stimulation did not seem to be ABA-glucoside deconjugation. The occurrence of b-glucosidase peak in the period
before and after the main peak of CKs indicates that
hydrolysis of CK O-glucosides may be involved in the finetuning of active CK levels at midday, even if this deglucosylation cannot be the only source of the midday peak
of physiologically active CKs. As CK translocation from
the roots, which is the primary site of CK biosynthesis
(Letham, 1963; Aloni et al., 2005), was reported to be
high in the morning, declining rapidly during the day (Beck
and Wagner 1994), CK biosynthesis seems to be a more
probable source of the sharp midday increase of CKs. As
only free ABA was determined, hardly any conclusion may
be drawn about the role of b-glucosidase in the regulation
of free ABA levels at the ‘after-midday’ peak. This will
be the subject of further research. Determination of the
bulk leaf ABA content represents another methodological limitation, as for regulation of stomata aperture ABA
concentration in the guard cells and the surrounding apoplast is crucial, and the bulk ABA concentration includes
the ABA stored in mesophyll chloroplasts as well.
The physiological significance of CK peaks may be
related, apart from the stimulation of cell division in leaves
or the regulation of stomatal aperture, to the generation
of a potential CK signal to other plant parts (e.g. flower
induction, Macháčková et al., 1996) or the induction/
repression of CK responsive genes (Hare and van Staden,
1997). These genes usually display circadian oscillations
in their levels of abundance with maximal expression
during the light period (Thomas et al., 1997). The circadian
clock represents an important adaptation of life to the
conditions on the earth and provides the necessary external
and internal co-ordination of the metabolic processes.
The results here have shown that CKs, auxin, and ABA
levels undergo significant variations in leaves during the
day and night. These data provide, to our knowledge, the
first report of a diurnal rhythm of CK metabolic enzymes
and CK metabolites other than CK bases and ribosides. The
coincidence of the peak of CKs and auxin indicates that
the changes in plant hormone levels have physiological
relevance and correlate with the diurnal changes of plant
growth and development.
Supplementary data for this paper can be found at JXB
online.
Acknowledgements
The authors wish to acknowledge Eva Kobzová and Marie Korecká
for skilful technical assistance. This work was supported by grants
from the Grant Agency of the Czech Republic, No. 206/03/0369,
and from the Ministry of Education, Youth and Sports, LN
LN00A081.
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54
Journal of Experimental Botany, Vol. 57, No. 6, pp. 1413–1421, 2006
doi:10.1093/jxb/erj121 Advance Access publication 23 March, 2006
This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details)
RESEARCH PAPER
Diurnal changes in polyamine content, arginine and
ornithine decarboxylase, and diamine oxidase in
tobacco leaves
Lenka Gemperlová1,2, Marie Nováková1,3, Radomı́ra Vaňková1, Josef Eder1 and Milena Cvikrová1,*
1
Institute of Experimental Botany, Academy of Sciences of Czech Republic, Rozvojová 135, 165 02 Prague 6,
Czech Republic
2
Department of Plant Physiology, Faculty of Sciences, Charles University, Viničná 5, 128 44 Prague 2,
Czech Republic
3
Department of Biochemistry, Faculty of Sciences, Charles University, Hlavova 2030/8, 128 44 Prague 2,
Czech Republic
Received 29 August 2005; Accepted 13 January 2006
Abstract
Changes in the contents of polyamines (PAs) in
tobacco leaves (Nicotiana tabacum L. cv. Wisconsin
38) grown under 16 h photoperiod were correlated with
arginine and ornithine decarboxylase (EC 4.1.1.19 and
EC 4.1.1.17) and diamine oxidase (EC 1.4.3.6) activities.
The maximum of free and soluble conjugated forms of
PAs occurred 1–2 h after the middle of the light period
and was followed by two distinct peaks at the end of
the light and at the beginning of the dark phase.
Putrescine was the most abundant and cadaverine
the least abundant PA in both free and PCA-soluble
forms. However, cadaverine was predominant in PCAinsoluble conjugates, followed by putrescine, spermidine, and spermine. Both arginine and ornithine
decarboxylases are involved in putrescine biosynthesis in tobacco leaves. Light dramatically stimulated the
activity of ornithine decarboxylase, while no photoinduction of arginine decarboxylase activity was observed. Ornithine decarboxylase was found mainly in
the particulate fraction. Only one peak, just after light
induction, occurred in the cytosolic fraction, with 35%
of the total ornithine decarboxylase activity. By contrast, the total arginine decarboxylase activity was
equally divided between the soluble and pellet fractions. A sharp increase in diamine oxidase activity
occurred 1 h after exposure to light, concomitant with
the light-induced increase in ornithine decarboxylase
activity. After a decline, diamine oxidase activity increased again, together with the rise in the amount of
free Put. The roles of both conjugation of PAs with
hydroxycinnamic acids and oxidative degradation of
putrescine in maintaining free PA levels during the
24 h light/dark cycle are discussed. The presented
results have shown that the parameters studied here
followed rhythmical changes and were not only affected by light.
Key words: Arginine decarboxylase, diamine oxidase, ornithine
decarboxylase, polyamines, tobacco.
Introduction
Polyamines (PAs) are ubiquitous organic polycations that
are involved in the regulation of different cellular processes
in animals and plants. The biological activity of PAs is
attributed to the cationic nature of these molecules, which
enables their interactions with anionic biomolecules (e.g.
DNA, RNA, proteins, and phospholipids). In plant cells,
PAs are usually present in amounts varying from micromolar to more than millimolar, i.e. levels significantly
higher than those of plant hormones. Nevertheless, PAs
affect many processes regulated by cytokinins and auxins
* To whom correspondence should be addressed. E-mail: [email protected]
Abbreviations: ADC, arginine decarboxylase; Cad, cadaverine; DAO, diamine oxidase; EDTA, ethylenediaminetetraacetic acid; GABA, c-aminobutyric acid;
ODC, ornithine decarboxylase; PA, polyamine; PAL, phenylalanine ammonia-lyase; PCA, perchloric acid; Put, putrescine; SAMDC, S-adenosylmethionine
decarboxylase; Spd, spermidine; Spm, spermine.
ª The Author [2006]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
The online version of this article has been published under an Open Access model. Users are entitled to use, reproduce, disseminate, or display the Open Access version of this article
for non-commercial purposes provided that: the original authorship is properly and fully attributed; the Journal and the Society for Experimental Biology are attributed as the original
place of publication with the correct citation details given; if an article is subsequently reproduced or disseminated not in its entirety but only in part or as a derivative work this must be
clearly indicated. For commercial re-use, please contact: [email protected]
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55
1414 Gemperlová et al.
described in leaves of Pharbitis nil (Yoshida and Hirasawa,
1998). Transcription of the SAMDC gene was shown to
be under circadian control (Yoshida et al., 1999). Changes
in PA content, ADC and ODC activities during light/
dark phases in maize calluses did not only respond to light,
but they were also found to be regulated by a daily
rhythm (Bernet et al., 1999). Photoperiodic control of
PA accumulation was observed in petal explants of
Araujia sericifera (Moysset et al., 2002).
The purpose of the current work was to identify the daily
variation of PA levels in tobacco plants in order to find
the optimal time for sample collection. These data will be
used in further studies of the role of PAs in abiotic stress
response and PA cross-talk with cytokinins using transgenic tobacco plants over-expressing cytokinin metabolic
enzymes under different (constitutive as well as inducible)
promoters. Moreover, determination of diurnal variation
of the levels of individual PAs might contribute to the
understanding of mechanisms involved in the regulation
of physiologically active PAs.
Data presented here show detailed changes in the contents of free Put, Spd, Spm, and Cad, their soluble and insoluble conjugates in tobacco leaves grown under a long
day. The focus was on the evaluation of the effect of light
on ADC and ODC activities, as well as on the elucidation
of the potential correlation of ADC and/or ODC activities
and Put and, subsequently, Spd and Spm accumulation.
The role of PA conjugation and oxidative deamination in
PA homeostasis during the 24 h cycle was followed, too.
and, in co-operation with these plant hormones, modulate
various morphogenic processes (Altamura et al., 1993). In
plants, PAs participate in the control of such important
processes as cell division and growth (Theiss et al., 2002)
morphogenesis and differentiation (Paschalidis et al.,
2001), programmed cell death (Serafini-Fracassini et al.,
2002; Pignatti et al., 2004), stabilization of nucleic acids
and membranes (Thomas and Thomas, 2001), and stress
responses (Bouchereau et al., 1999).
The polyamine content in cells can be regulated by both
metabolic and transport processes. In higher plants PAs
are biosynthesized by two main pathways. The diamine
putrescine (Put) may be formed directly from ornithine by
ornithine decarboxylase (ODC; EC 4.1.1.17) or indirectly
from arginine by arginine decarboxylase (ADC; EC
4.1.1.19). S-adenosylmethionine decarboxylase (SAMDC;
EC 4.1.1.50) is essential for spermidine (Spd) and spermine
(Spm) biosynthesis. Spd is formed from Put as well as Spm
from Spd by successive addition of aminopropyl groups
derived from decarboxylated S-adenosylmethionine generated by the activity of SAMDC (Wallace et al., 2003). A
clear relationship between the diamine cadaverine (Cad)
and other PAs has not yet been found (Bagni and Tassoni,
2001). The intracellular concentrations of free PAs may be
regulated by the rate of biosynthesis, which is controlled
at the transcriptional, post-transcriptional, translational, and
post-translational levels (Kumar et al., 1997). Apart from
biosynthesis, PA levels may be modulated by conjugation
either with small molecules, especially hydroxycinnamic
acids (soluble conjugated PAs; Martin-Tanguy, 1985;
Bagni and Tassoni, 2001), or with high molecular-mass
substances like hemicelluloses, lignin, and, to a lesser
extent, proteins as well (insoluble conjugated PAs; Creus
et al., 1991). Cytoplasmic levels of PAs are also affected
via storage in the vacuoles, mitochondria, and chloroplasts and by extracellular transport (Flores, 1991;
Beranger-Novat et al., 1997; Bagni and Tassoni, 2001).
The mechanisms which regulate polyamine transport in
eukaryotes remain largely unknown. Only translocation of
free polyamines has been reported (Antognoni et al., 1998).
Another important process of PA level regulation is
degradation through oxidative deamination. Two catabolic
enzymes were found in plants. Diamine oxidase (DAO;
EC 1.4.3.6) which primarily catalyses the conversion of Put
to ammonia, H2O2 and pyrroline, and polyamine oxidase
(PAO; EC 1.5.3.) with high affinity towards Spd and Spm.
Data on the relationship between the physiological
effects of PAs and light are not very extensive. Alterations
in the contents and forms of PAs during photoperiodic
flower induction were observed in soybean (Caffaro and
Vicente, 1994). Photosynthetically active radiation caused
Put and Spd accumulation in soybean (Kramer et al.,
1992), which is consistent with the hypothesis that phytochrome can regulate the activity of ADC in chloroplasts
(Besford et al., 1993). Photo-induction of ADC was also
Materials and methods
Plant material
Tobacco (Nicotiana tabacum L. cv. Wisconsin 38) plants were grown
in soil in a growth chamber for 6 weeks at a 16/8 h photoperiod
(130 lmol mÿ2 sÿ1) with a day/night temperature of 25/23 8C and
relative humidity c. 80%. Mixed samples of leaves (3rd–5th of 7,
i.e. with the exception of the youngest and the oldest ones) from
three plants were pooled and collected every h during a 24 h
period. The sampling during the dark period was realized in dim
light and did not exceed 1 min. The samples were immediately frozen
in liquid nitrogen.
Polyamine analysis
The leaves were ground in liquid nitrogen and extracted overnight
at 4 8C with 5% (v/v) perchloric acid (PCA) (100 mg fresh weight
tissue cmÿ3 5% PCA). 1,7-Diaminoheptane was added as an internal standard. The extracts were centrifuged at 21 000 g for 15 min,
and then PCA-soluble free PAs were analysed in one-half of the
supernatant. The remaining supernatant and pellet were acid hydrolysed in 6 M HCl for 18 h at 110 8C to obtain PCA-soluble and
PCA-insoluble conjugates of PAs as described by Slocum et al.
(1989). Standards (Sigma-Aldrich, St Louis, MO, USA), PCAsoluble free PAs, and acid hydrolysed PA conjugates were benzoylated. HPLC analysis of benzoyl-amines was performed on a
Beckman-Video Liquid Chromatograph equipped with a UV detector
(detection at 254 nm) and C18 Spherisorb 5 ODS2 column (particle
size 5 lm, column length 25034.6 mm) according to the method
of Slocum et al. (1989).
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56
Diurnal changes in polyamine content 1415
Statistical analyses
Means 6SE of two independent experiments with two replicates
are shown in the figures. Statistical tests were analysed using the
Student’s t distribution criteria.
Results
Polyamine contents
Free and bound PAs (PCA-soluble and PCA-insoluble
conjugates) were analysed in tobacco leaves each h during
a 24 h light/dark cycle. All four PAs (Put, Spd, Spm, and
Cad) were present in both free and conjugated forms.
Although the content of PAs is usually expressed on a fresh
weight basis, it was decided to express the PA contents in
nmol gÿ1 protein as well (Fig. 1) because the focus of this
work was on the evaluation of the potential correlation
between biosynthetic enzyme activities and the contents of
PAs. Differences between the graphs are related to changes
in protein in the course of the light/dark cycle, which were
higher during the light, especially in the first half of the
light period. As a consequence, the peaks of PAs per
fresh weight at the end of the light and at the beginning of
the dark phase (Fig. 1A) became more pronounced when
expressed per protein (Fig. 1B).
Total contents of PAs (nmol mg-1 protein)
Diamine oxidase assay
Diamine oxidase (DAO, EC 1.4.3.6) activity was assayed by
a spectrophotometric method based on detection of the aldehyde
with cis-1,4-diamino-2-butene as the substrate (Peč et al., 1991).
Samples were homogenized in 0.1 M TRIS–HCl buffer, pH 8.5,
containing 2 mM mercaptoethanol and 1 mM EDTA, and centrifuged at 20 000 g for 15 min at 4 8C. The reaction mixture contained
0.1 M TRIS–HCl buffer, pH 8.5, catalase (25 lg) and 0.01 M
cis-1,4-diamino-2-butene. The reaction was started by the addition
of 0.2 cm3 supernatant, incubated for 1 h at 37 8C and stopped by
adding 1 cm3 of Ehrlich’s reagent. The reaction mixture was incubated at 50 8C for 5 min, and then chilled in an ice bath before
reading the absorbance of produced pyrrol at 563 nm. Enzymatic
activity is expressed in pkat mgÿ1 protein.
Protein content was measured according to Bradford’s method
using bovine serum albumin as a standard (Bradford, 1976).
Total contents of PAs (nmol g-1 FW)
Ornithine decarboxylase and arginine decarboxylase assays
Ornithine decarboxylase (ODC; EC 4.1.1.17) and arginine decarboxylase (ADC; EC 4.1.1.19) were determined by a radiochemical
method described by Tassoni et al. (2000). Samples were extracted in
3 vols of ice-cold 0.1 M TRIS–HCl buffer, pH 8.5, containing 2 mM
b-mercaptoethanol, 1 mM EDTA and 0.1 mM pyridoxal phosphate,
and centrifuged at 20 000 g for 30 min at 4 8C. Aliquots (0.1 cm3)
of both supernatant (soluble fraction) and resuspended pellet (particulate fraction) were used to determine ODC and ADC activity.
Enzyme activity assays were performed by measuring the 14CO2
evolution from 7.4 kBq L-[1-14C]ornithine (1.92 GBq mmolÿ1,
Amersham Pharmacia Biotech UK) or 7.4 kBq L-[U-14C]arginine
(11.5 GBq mmolÿ1 Amersham Pharmacia Biotech UK), for ODC
and ADC, respectively, in the presence of 2 mM unlabelled
substrate during a 1.5 h incubation at 37 8C. CO2 was trapped in
hyamine hydroxide and the radioactivity was counted by a liquid
scintillation analyser (Tri-Carb 2900TR; Packard).
2500
2000
Put
Cad
Spd
Spm
A
Put
Cad
Spd
Spm
B
1500
1000
500
0
500
400
300
200
100
0
0
4
8
12
16
20
Time (h)
Fig. 1. Changes in total contents of Put, Spd, Spm, and Cad (represented
by the sum of free, PCA-soluble and insoluble conjugates) in tobacco
leaves grown under a 16/8 h photoperiod. (A) PA contents expressed
as nmol gÿ1 FW; (B) PA contents expressed as nmol mgÿ1 protein. The
photoperiod is represented by the black and white boxes on the abscissa.
Means 6SE of two independent experiments with two replicates are
shown.
The ratio among PA forms changed during the 24 h cycle
(Fig. 2). The most abundant PA in free and PCA-soluble
form was Put and the least abundant was Cad. However,
Cad prevailed in PCA-insoluble conjugates, followed by
Put, Spm, and Spd (Fig. 3). The endogenous level of free
Put showed a transient increase at the second and the third
hours of the light period (P <0.05 compared with the value
at the end of the dark period) and exhibited a maximum
after 9 h of light (P <0.005 compared with the value at 7 h).
The largest peak occurred at the light/dark transition
(P <0.005 compared with the value at 11 h; Fig. 2A).
The contents of PCA-soluble Put conjugates showed a first
peak 1 h after the middle of the light period (P <0.005 compared to the value at 7 h), and further transient peaks were
detected 2 h before the light was switched off and at the
beginning of the dark period (statistical significance of both
peaks P <0.005 compared with the value at 13 h). A slight
rise was observed at the end of the dark phase (Fig. 2A).
The peaks in the free Spd content were less marked than
those of Put (statistical significance of the increase at 11 h
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57
1416 Gemperlová et al.
F1
F2
200
A
Cad (nmol mg-1 protein)
Put (nmol mg-1 protein)
250
200
150
100
50
0
120
F1
F2
B
100
80
60
40
20
Cad (nmol mg-1 protein)
Spm (nmol mg-1 protein)
0
50
40
150
100
50
0
Put, Spd, Spm (nmol mg-1 protein)
Spd (nmol mg-1 protein)
140
A
C
F1
F2
30
B
Put
Spd
Spm
20
10
0
0
4
8
12
16
20
Time (h)
30
Fig. 3. Changes in PCA-insoluble conjugates of Cad (A) and Put, Spd,
Spm (B) in tobacco leaves grown under a 16/8 h photoperiod. Other
details as for Fig. 2.
20
10
peaks, the contents of soluble conjugates of Put and Spd
were significantly higher than those of their free forms
(PCA-soluble forms represented about 75% of the total
content at the midday maximum). The contents of free
Spm and Cad and of their soluble conjugates did not
change significantly during the diurnal cycle, except that
of Spm conjugates, which increased 5 h before the end
of the light (P <0.05 compared with the value at 10 h
of light) and at the beginning of the dark period (P <
0.005 compared to the value at 10 h of light; Fig. 2C, D).
The time-course of the levels of PCA-insoluble conjugates showed different trends from those observed in
free and soluble forms (Fig. 3). Three dominant peaks of
insoluble Cad conjugates were detected in tobacco leaves
grown under the 16/8 h photoperiod. After the decline at the
dark/light transition the first maximum occurred at the
beginning (at the second and third hours) of the light period
(P <0.005 compared with the value at 1 h of light), the
second one 4 h after midday (P <0.005 compared with the
value at 10 h of light), and the third one at the beginning
of the dark period (2 h after the light was switched
off, P <0.005 compared with the value at the light/dark
transition). Insoluble conjugates of Put, after the transient
0
15
D
F1
F2
10
5
0
0
4
8
12
16
20
Time (h)
Fig. 2. Changes in free and PCA-soluble conjugates of PAs in tobacco
leaves grown under 16/8 h photoperiod. (A) Put contents; (B) Spd
contents; (C) Spm contents; (D) Cad contents. F1, free PAs; F2, PCAsoluble conjugates. The photoperiod is represented by the black and
white boxes on the abscissa. Means 6SE of two independent experiments with two replicates are shown. For further details, see text.
of light was P <0.05 compared with the value at 7 h, which
was similar to the peaks at the light/dark transition
compared with 13 h). PCA-soluble conjugates of Spd
showed a similar trend to those of Put (Fig. 2B). In the
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Diurnal changes in polyamine content 1417
decline at the dark/light transition, reached their high value
in the middle of the light period (P <0.005 compared with
the value at 4 h of light), and then decreased. Their second
peak coincided with that of Cad (2 h after the light was
switched off, P <0.005 compared with the value at 14 h
of light). Relatively high level of insoluble conjugates of
Cad and Put were observed during the dark period. However, the amounts of insoluble Put conjugates were much
lower than those of Cad. The content of insoluble Spd
conjugates was the highest at the first third of the light
period. The contents of Spm conjugates followed a trend
similar to Cad but the peaks were much less pronounced.
ADC activity (pkat mg-1 protein)
10
ADC and ODC activities
DAO activity
DAO activity was induced by light in a similar way as the
ODC activity. A sharp peak of activity was observed 1 h
after the start of light (P <0.005 compared with the value at
the dark/light transition), after which the activity declined.
The main DAO maximum was found 2 h after midday
(P <0.05 compared with the value at 6 h of light) and then
the enzyme activity steadily declined. The peak of activity
at the light/dark transition (P <0.05 compared with the
value at 14 h of light) coincided with the increase in the
activities of both biosynthetic enzymes (Fig. 6).
8
6
4
2
0
0
4
8
12
16
20
Time (h)
Fig. 4. Time-course of ADC activity found in soluble and particulate
fractions in tobacco leaves grown under a 16/8 h photoperiod. Other
details as for Fig. 2.
ODC activity (pkat mg-1 protein)
Biosynthetic enzyme activities were measured both in the
soluble and pellet fractions. No increase in ADC activity
was observed in light, by contrast, the enzyme activity in
the first hour of light decreased. At midday, ADC activity
started increasing, reaching a high 1 h later (P <0.005
compared with the value at 7 h of light). After a decline,
a second main peak of activity occurred at the light/dark
transition (P <0.005 compared with the value at 12 h of
light). The enzyme activity in the pellet fraction represented
about 45% and 48% of the entire biosynthetic activity in
the first and second enzyme peak, respectively (Fig. 4).
Light dramatically stimulated the activity of the second
PA biosynthetic enzyme, ODC (Fig. 5). The sharp increase
at 1 h of light (P <0.005 compared with the value at the
dark/light transition) was followed by a transient decrease and then further peaks were observed prior to
and after the middle of the light period (statistical significance of both peaks P <0.005 compared with the
value at 3 h) and at the beginning (P <0.005 compared
with the value at the light/dark transition) and the end of
the dark period (P <0.05 compared with the value at 4 h
of dark). The ODC enzyme activity was present predominantly in the pellet fraction and was rather low in
the soluble fraction except for a peak 1 h after light induction (P <0.005 compared with the value at the dark/light
transition), at which time it represented about 35% of
total ODC activity. From the third hour of light until the
end of the dark period the soluble ODC activity was very
low again.
soluble
particulate
10
soluble
particulate
8
6
4
2
0
0
4
8
12
16
20
Time (h)
Fig. 5. Time-course of ODC activity found in soluble and particulate
fractions in tobacco leaves grown under a 16/8 h photoperiod. Other
details as for Fig. 2.
Discussion
Although the number of studies on PAs has been growing
constantly, it is still unclear whether ODC or ADC plays
a decisive role in Put biosynthesis in plants. The existence
of two alternative routes for Put synthesis could be explained by the necessity of specific regulation of different
processes affected by Put during growth and development
(Koetje et al., 1993). It has been suggested that the pathway via ADC is generally linked to stress responses and
morphogenetic processes, whereas ODC is involved in the
regulation of cell division in actively growing plant cell
tissue (Koetje et al., 1993; Galston et al., 1997). However,
it seems that species and/or tissue specificity should be
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59
1418 Gemperlová et al.
non-photosynthetic tissues of tobacco, the particulate ADC
activity appeared to be located mainly in nuclei (Bortolotti
et al., 2004). The different compartmentation of biosynthetic enzymes may be related to distinct functions in
different cell types (Bortolotti et al., 2004).
The results presented here show that, in tobacco leaves,
the total ADC activity was equally divided between the
soluble and pellet fractions while the ODC activity was
mainly localized in the pellet fraction and only very little
activity was present in the cytosolic fraction (there was
one peak of activity after light induction in which 35%
of the total ODC activity was found).
A significant peak of free and soluble conjugated PA
forms observed in tobacco leaves 1–2 h after the middle of
the light period coincided with the high activity of ODC
pellet fraction. It is well documented that in many plant
systems dividing tissues have high PA levels and activities
of their biosynthetic enzymes (Pfosser et al., 1990; Bezold
et al., 2003). In vitro experiments have shown that free
PAs affect many functions of nucleic acids, including transcription and translation (for a review see Kumar et al.,
1997). The observed midday peak in the contents of free
Put and Spd in tobacco leaves may be related to high
physiological activity of the leaf tissue at this time
period. This hypothesis might be supported by the abovementioned fact that ODC expression coincided with early
cell divisions observed in hypogenous tobacco tissues
(Paschalidis and Roubelkis-Angelakis, 2005). Furthermore, this peak coincided with the main peak of cytokinins and with the maximum of indole-3-acetic acid
(IAA) in tobacco leaves cultivated under identical light/
dark conditions (Nováková et al., 2005). Since cytokinins,
auxin, and PAs are necessary for the progression through
the cell cycle, this result leads to the assumption that cell
division occurs at this period.
At midday, growth in ADC activity was observed and
the main peak of the activity was found at the light/dark
transition. This distinct peak of enzymatic activity was
accompanied by the accumulation of free PAs and PCAsoluble conjugates. The PA maximum occurring at the
light/dark transition coincided with the increase in abscisic acid content determined in tobacco leaves cultivated
under the identical light/dark conditions (Nováková et al.,
2005). As ABA is a key component of plant defence, the
coincidence of its maximum and ADC peak is in a good
accordance with the presumption of a potential link of
ADC to stress response.
The observed midday and dark phase peaks in Put and
Spd contents are in agreement with the accumulation
pattern of PAs in maize calli during the light/dark phases
(Bernet et al., 1999). Put was the most abundant PA in
maize callus, as in tobacco leaves, reaching peaks at 8 h and
12 h in the light and then in the middle of the dark period.
Large intracellular accumulation of free PAs would have
harmful effects on the maintenance of cellular pH, ion
DAO activity (pkat mg-1 protein)
5
4
3
2
1
0
0
4
8
12
16
20
Time (h)
Fig. 6. Time-course of DAO activity in tobacco leaves grown under
a 16/8 h photoperiod. Other details as for Fig. 2.
taken into consideration in plants. In apple, the primary
biosynthetic pathway for Put is biosynthesis through ADC
(Hao et al., 2005), while in the green alga Chlamydomonas
reinhardtii the activity of ODC exceeded that of ADC by
orders of magnitude, being highly induced by light (Voigt
et al., 2000). On the contrary, in leaves of Pharbitis nil
a very rapid photoresponse of ADC with the major activity
peak 1 h after illumination was observed, while the activity of ODC did not respond to illumination (Yoshida and
Hirasawa, 1998). In tobacco plants, ADC seemed to be responsible for Put synthesis in old hypergenous vascular
tissues, whereas ODC expression coincided with early cell
divisions and catalysed Put synthesis in hypogenous tissues
(Paschalidis and Roubelkis-Angelakis, 2005).
In tobacco leaves a different effect of light was observed
on the induction of PA biosynthetic enzymes. Marked
photoinduction of ODC activity 1 h after the exposure to
light was found. Further peaks of ODC were observed prior
to and after the middle of the light period with a slight
increase at the beginning and the end of the dark period.
By contrast, ADC activity decreased at the first hour of
the light period and started increasing at midday with the
main peak at the light/dark transition.
There is still a lack of knowledge about the tissue and
subcellular localization of the enzymes involved in PA
metabolism. In animal tissues, despite extensive research,
the exact intracellular localization of ODC remains to be
clarified. It varies from an exclusively cytoplasmic to both
a cytoplasmic and a nuclear localization (Schipper et al.,
2004). The assays performed on leaves of Arabidopsis
thaliana showed that high particulate ODC activity was
associated with the nuclei and chloroplast-enriched fractions (Tassoni et al., 2003). In photosynthetic tissues of
tobacco plants ADC was found mainly in chloroplasts,
mitochondria, and cytoplasm (Slocum, 1991), whereas in
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60
Diurnal changes in polyamine content 1419
homeostasis, and on a number of physiological functions
where PAs are implied. The endogenous free PA levels
could be regulated, apart from the control of PA biosynthesis, also by conjugation with hydroxycinnamic acids
in some plant species (Martin-Tanguy, 1985; Creus et al.,
1991; Bagni and Tassoni, 2001; Biondi et al., 2001). The
detected high levels of PCA-soluble conjugates of Put and
Spd (about 75% of the entire Put and Spd pool in the peak
1–2 h after the midday) seem to be the common phenomena
in Nicotiana and have been reported by several authors
(Martin-Tanguy, 1985; Pfosser et al., 1990; Gemperlová
et al., 2005). It has been shown in alfalfa cell suspension
cultures and in oak embryogenic cultures that the rate of
PA conjugation with hydroxycinnamic acids influenced
the endogenous free PA level and, indirectly, cell division (Cvikrová et al., 1999, 2003). The biological function
of conjugated PAs remains unclear. It has been speculated
that enzymatic hydrolysis of reversible conjugates could
supply the cells with an additional amine reserve (Bonneau
et al., 1994).
The origin and function of bound PAs are still an open
issue. Some reports demonstrated high levels of Put
and Spd binding to proteins in actively dividing young
tissues. The incorporation of Put, Spd, and Spm into
thylakoid and stromal proteins was stimulated by light
(Aribaud et al., 1995). Spm was the most efficiently conjugated PA (Della Mea et al., 2004). In tobacco leaves
grown under a 16/8 h photoperiod the most abundant PA
in PCA-insoluble conjugates was Cad, followed by Put,
Spm, and Spd. Cad in animal tissues does not normally
participate in any particular metabolic process, but it is
completely excreted from cells into the culture medium
(Hawel et al., 1994; Hawel and Byus, 2002). Synthesized
Cad is, in tobacco leaves, most probably immediately incorporated into the PA-insoluble fraction, as the levels of
both free and soluble conjugated forms are very low during
the light/dark cycle. In this experiment, the peaks of
insoluble Spm conjugates coincided with the peaks of
Cad. It may be hypothesized that Cad participates in the
protection of chloroplasts from photo damage or stress.
Significant light-affected conjugation of PAs to endogenous proteins was observed in isolated chloroplasts from
Helianthus tuberosus (Dondini et al., 2003). It should be
noted that light quality is also important for PA metabolism. Lettuce explants cultured under blue light contained
a higher proportion of PCA-insoluble conjugated PAs
compared with explants grown under white or red light,
which contained more of free PA and/or soluble conjugates
(Hunter and Burritt, 2005).
The level of free and PCA-soluble PAs seems to be
during the light/dark cycle regulated by the activities of
biosynthetic enzymes (in tobacco especially ODC) and
the catabolic one, DAO, activity of which was found to
be substrate inducible (Srivastava et al., 1977). The sharp
increase in DAO activity in tobacco leaves 1 h after the
exposure to light closely followed or was even concomitant
with the light-induced increase in the activity of ODC. This
may be the reason why only a slight increase in free Put
content was observed after the photoinduction of ODC
activity at the beginning of the light phase. The role of
both PA conjugation with hydroxycinnamic acids and oxidation in maintaining free PA levels was documented with
leaf explants of Chrysanthemum after inhibition of DAO
activity. High levels of PAs, especially soluble PA conjugates accumulated in these explants (Aribaud et al., 1994).
Timing of the main peak of DAO activity shortly (1 h)
after the middle of the light phase is in agreement with the
enzyme maximum observed in isolated etioplasts from
Hordeum vulgare after light exposure for 9 h (Andreadakis
and Kotzabasis, 1996).
The DAO was found to be localized in cell walls in pea
and maize tissues (Slocum and Furey, 1991) and imunohistochemical analyses revealed that DAO expression
occurs in cells destined to undergo lignification in tobacco
tissues (Paschalidis and Roubelakis-Angelakis, 2005). However, the comparable soluble and cell debris-associated
DAO activity was measured in tobacco thin layers (Biondi
et al., 2001) and the decline in Put content in the early Sphase of the cell cycle in Helianthus tuberosus was
probably due to the high DAO activity occurring in the
early S-phase (Serafini-Fracassini, 1991). From the published literature it is clear that the catabolic degradation of
Put by DAO is not only a means to regulate the cellular
content of this diamine (Kumar et al., 1997; Bagni and
Tassoni, 2001). The reaction product of DAO, pyrroline,
can be further catabolized to c-aminobutyric acid (GABA)
and subsequently trans-aminated and finally incorporated
into the Krebs cycle (Tiburcio et al., 1997) and/or GABA
has been shown to play a key role in signal transduction
pathways during stress response of many plants (Shelp
et al., 1999). The other reaction product of DAO, hydrogen peroxide, might trigger the hypersensitive response,
which is thought to be a form of programmed cell death
(Walters, 2003) or might be utilized in lignification both
during the normal growth and stress response (Angelini
et al., 1993).
The results presented here have shown that ADC and
ODC activities, as well as putrescine oxidizing activity,
correlated well with the alterations in PA content during
the day/night period. The data indicate that the parameters studied here were not only light-affected but also
appear to be regulated by an internal rhythm.
Acknowledgements
We are grateful to Dr I Macháčková and Professor M Mok for
critical reading of the manuscript. We thank N Hatašová for skilful
technical assistance. This work was supported by grants from the
Grant Agency of the Czech Republic No. 206/03/0369.
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61
1420 Gemperlová et al.
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Journal of Experimental Botany Advance Access published December 14, 2006
Journal of Experimental Botany, Page 1 of 13
doi:10.1093/jxb/erl235
RESEARCH PAPER
Altered cytokinin metabolism affects cytokinin, auxin, and
abscisic acid contents in leaves and chloroplasts, and
chloroplast ultrastructure in transgenic tobacco
Lenka Polanská1,2,*, Anna Vičánková1,2, Marie Nováková1,3, Jiřı́ Malbeck1, Petre I. Dobrev1,
Břetislav Brzobohatý4, Radomı́ra Vaňková1 and Ivana Macháčková1
1
Institute of Experimental Botany AS CR, Rozvojová 135, 165 02 Prague 6-Lysolaje, Czech Republic
Charles University in Prague, Faculty of Science, Department of Plant Physiology, Viničná 5, 128 44 Prague 2,
Czech Republic
2
3
Charles University in Prague, Faculty of Science, Department of Biochemistry, Hlavova 2030/8, 128 43 Prague 2,
Czech Republic
4
Institute of Biophysics AS CR, Královopolská 135, 612 65 Brno, Czech Republic
Received 2 August 2006; Revised 12 October 2006; Accepted 16 October 2006
Abstract
Cytokinins (CKs) are involved in the regulation of plant
development including plastid differentiation and function. Partial location of CK biosynthetic pathways in
plastids suggests the importance of CKs for chloroplast
development. The impact of genetically modified CK
metabolism on endogenous CK, indole-3-acetic acid,
and abscisic acid contents in leaves and isolated
intact chloroplasts of Nicotiana tabacum was determined by liquid chromatography/mass spectrometry
and two-dimensional high-performance liquid chromatography, and alterations in chloroplast ultrastructure
by electron microscopy. Ectopic expression of Sho,
a gene encoding a Petunia hybrida isopentenyltransferase, was employed to raise CK levels. The increase
in CK levels was lower in chloroplasts than in leaves.
CK levels were reduced in leaves of tobacco harbouring a CK oxidase/dehydrogenase gene, AtCKX3. The
total CK content also decreased in chloroplasts, but
CK phosphate levels were higher than in the wild type.
In a transformant overexpressing a maize b-glucosidase
gene, Zm-p60.1, naturally targeted to plastids, a decrease of CK-O-glucosides in chloroplasts was found.
In leaves, the changes were not significant. CK-Oglucosides accumulated to very high levels in leaves,
but not in chloroplasts, of plants overexpressing
a ZOG1 gene, encoding trans-zeatin-O-glucosyltransferase from Phaseolus lunatus. Manipulation of the CK
content affected levels of indole-3-acetic and abscisic
acid. Chloroplasts of plants constitutively overexpressing Sho displayed ultrastructural alterations including the occasional occurrence of crystalloids and
an increased number of plastoglobuli. The other transformants did not exhibit any major differences in
chloroplast ultrastructure. The results suggest that
plant hormone compartmentation plays an important
role in hormone homeostasis and that chloroplasts are
rather independent organelles with respect to regulation of CK metabolism.
Key words: Abscisic acid, auxin, chloroplast ultrastructure,
cytokinin metabolism, cytokinin oxidase/dehydrogenase, bglucosidase, isopentenyltransferase, Nicotiana tabacum, Sho,
zeatin-O-glucosyltransferase.
Introduction
The plant hormones cytokinins (CKs) are known to be
involved in many processes related to plastid development
and functions. Within 10 years of the discovery of kinetin
(6-furfurylaminopurine) as a plant growth regulator CK
(Miller et al., 1956) in cultured tobacco tissue, it was
* To whom correspondence should be addressed. E-mail: [email protected]
Abbreviations: ABA, abscisic acid; BA, benzyladenine; CaMV, cauliflower mosaic virus; CK, cytokinin; DMSO, dimethylsulphoxide; FW, fresh weight; IAA,
indole-3-acetic acid; IPT, isopentenyltransferase; LCMS, liquid chromatography/mass spectrometry; WT, wild type.
ª The Author [2006]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
64
2 of 13 Polanská et al.
CKs may be synthesized in plastids by the methylerythritol phosphate pathway and that four isopentenyltransferases (IPTs) are localized to plastids in Arabidopsis
cells. Unexpectedly, Sakakibara et al. (2005) have found
that Agrobacterium IPT functions in plastids, even though
it lacks a typical plastid-targeting sequence, and it prefers
a substrate different from that of genuine plant IPTs. It
was previously shown that the enhanced content of CKs
in chloroplasts of tobacco plants overexpressing the A.
tumefaciens ipt gene driven by a light-inducible promoter
of a Pisum sativum small subunit of Rubisco (Pssu-ipt)
correlated with highly anomalous chloroplast ultrastructure (Synková et al., 2006). Interestingly, although a clear
increase in CKs was observed in chloroplasts, it remained
about an order of magnitude lower than the levels found
in whole leaf extracts.
In the present study, the impact of manipulation of CK
metabolism on the endogenous CK pool in leaves and
isolated intact chloroplasts, and on chloroplast ultrastructure in tobacco (Nicotiana tabacum L.) was evaluated.
CK-overproducing plants (expressing the Sho gene coding
for a Petunia hybrida IPT), plants with decreased CK
levels (overexpressing CK oxidase/dehydrogenase
AtCKX3 from Arabidopsis thaliana), and plants with
altered CK glucoconjugation (overexpressing maize bglucosidase Zm-p60.1 or zeatin-O-glucosyltransferase
ZOG1 from Phaseolus lunatus) were compared. The simplified model of CK metabolism with highlighted enzymes
involved in this study is presented in Fig. 1. Taking into
account the immense cross-talk among plant hormones,
the contents of auxin (IAA) and abscisic acid (ABA) were
also determined.
shown that kinetin could stimulate chloroplast differentiation from proplastids, including grana formation (Stetler
and Laetsch, 1965). Kinetin was also found to almost
double chloroplast number per cell in etiolated tobacco
leaf discs incubated in the light (Boasson and Laetsch,
1969).
It has been demonstrated that exogenously applied CKs
induce partial development of chloroplasts from proplastids, amyloplasts, and etioplasts, especially in darkness
(Chory et al., 1994; Kusnetsov et al., 1994). For example,
benzyladenine (BA) treatment of excised watermelon
cotyledons accelerated degradation of reserve material
and differentiation of plastids with a more developed inner
membrane system, as well as increased levels of plastid
pigments and enzymes (Longo et al., 1979). Moreover,
applied BA accelerated the redevelopment of grana and
stroma in regreening plastids, the increase of reappeared
NADPH-protochlorophyllide oxidoreductase, and the contents of chlorophyll and protein in regreening tobacco
leaves (Zavaleta-Mancera et al., 1999a, b).
The action of CKs in plastids seems to be mediated by
the stimulation of the expression of several plastid-related
genes, of both nuclear and plastid origin, in particular that
of the nuclear gene for chlorophyll a/b-binding polypeptide of the light-harvesting complex and the gene
coding for the small subunit of Rubisco in Nicotiana
tabacum (Abdelghani et al., 1991), Arabidopsis (Chory
et al., 1994), or Dianthus caryophyllus (Winiarska et al.,
1994). For extensive reviews, see Schmülling et al. (1997)
or Parthier (1989).
Recently, Brenner et al. (2005) identified by genomewide expression profiling five rapidly CK-induced plastid
transcripts in Arabidopsis seedlings, indicating a fast
transfer of the CK signal to plastids or its direct perception
there. The CK effect on gene expression may be mediated
via the hormone interaction with specific proteins. In the
presence of trans-zeatin, a 64 kDa chloroplast zeatinbinding protein was found to activate the chloroplast but
not the nuclear genome (Lyukevich et al., 2002).
The occurrence of endogenous CKs in plastids has been
proven. CKs are present in chloroplast tRNAs (Vreman
et al., 1978). Zeatin-type CKs were detected in isolated
spinach chloroplasts by paper chromatography and soybean callus bioassay (Davey and Van Staden, 1981).
Immunolocalization of CKs revealed low labelling in the
stroma of plastids of Tilia cordata embryo cotyledons
and roots (Kärkönen and Simola, 1999). Eventually, CK
occurrence in chloroplasts was rigorously confirmed by
LCMS (liquid chromatography/mass spectrometry) which
allows identification of a whole range of CK metabolites
with high accuracy (Benková et al., 1999).
The importance of CKs for plastid development and
function may be deduced from the partial localization of
the CK biosynthetic pathway to this compartment.
Kasahara et al. (2004) showed that the prenyl group of
Materials and methods
Plant materials and growth conditions
Samples of mature leaves from four different types of transgenic
tobacco plants with altered CK metabolism and the corresponding
wild types (WTs), all in the vegetative stage, collected imediately
after the dark period, were used.
1. Tobacco in vitro lines expressing an IPT homologue Sho
(Shooting) gene from P. hybrida under the control of a constitutive
cauliflower mosaic virus (CaMV) 35S promoter or construct with
four 35S enhancer elements (Zubko et al., 2002) and WT N.
tabacum L. cv. Petit Havana SR1 cultivated on solid hormone-free
MS medium (Murashige and Skoog, 1962) supplemented with
sucrose (30 g l1). To achieve inducible Sho expression, a dexamethasone-inducible gene expression system (pOp6/LhGR;
Šámalová et al., 2005) was employed. Sho expression was induced
in 8–9-week-old transgenic plants with 20 lM dexamethasone [in
0.05% dimethylsulphoxide (DMSO), 5350 ml per plant watered
for 13 d]. As controls, transgenic plants treated with water or
0.05% DMSO and WT SR1 plants treated with water, 0.05%
DMSO, or 20 lM dexamethasone in 0.05% DMSO were used. The
plants were cultivated in a growth chamber (16/8 h photoperiod at
130 lE m2 s1, day/night temperature of 25/23 C and relative
humidity ~80%) for 5 weeks and then transferred, before Sho
induction, to a greenhouse (in summer, without additional light
supplementation).
65
Cytokinins in chloroplasts of transgenic tobacco 3 of 13
(1000 g; 2 min; 4 C). The pellet was resuspended in medium. All
the procedures were done at 4 C.
Chlorophyll determination
Chlorophyll was extracted into 80% (v/v) acetone. The total
chlorophyll content (a+b) was calculated from the absorbance at
652 nm of the clear extract after centrifugation (500 g, 5 min)
according to Arnon (1949).
Chloroplast integrity
The integrity of chloroplasts was determined by the latency of
glyceraldehyde-3-phosphate dehydrogenase according to Latzko
and Gibbs (1968). Chloroplasts were incubated in a reaction
mixture (0.33 M TRIS–HCl, pH 8.5; 17 mM Na2HAsO47H2O;
4 mM cysteine; 20 mM NaF; 40 lM NADP+). The reaction was
initiated with 0.02 M glyceraldehyde-3-phosphate. Reduction of
NADP+ was followed at 340 nm for 5 min. The same assay was run
with chloroplasts disrupted with 0.01 M MgCl2. The percentage of
intact chloroplasts was calculated from the difference between the
original and disrupted sample.
Fig. 1. Simplified model of CK metabolism including known localization of examples of involved enzymes proved by in planta experiments.
The enzymes employed in this study are highlighted. AtCKX,
Arabidopsis CK oxidase/dehydrogenase (Werner et al., 2003); AtIPT,
Arabidopsis isopentenyltransferase (Kasahara et al., 2004); Sho, petunia
isopentenyltransferase (Zubko et al., 2002); UGT, Arabidopsis Nglucosyltransferase (Hou et al., 2004); Zm-p60.1, maize b-glucosidase
(Kristoffersen et al., 2000); ZOG1, bean trans-zeatin-O-glucosyltransferase (Martin et al., 2001a); cisZOG, maize cis-zeatin-O-glucosyltransferase (Martin et al., 2001b); ZOX1, bean zeatin-O-xylosyltransferase
(Martin et al., 1993); ?, unknown or uncertain localization. The
interconversions among iP-, transZ-, cisZ-, and DHZ-type CKs and
their bases, ribosides, and phosphates, CK transport and tissue localization are not depicted. For more details see Sakakibara (2006).
All the other plants were cultivated in a soil substrate in a growth
chamber (16/8 h photoperiod at 150 lE m2 s1, 26/20 C and
relative humidity ~80%).:
2. Eleven-week-old tobacco plants (35S:AtCKX3) overexpressing
a gene for CK oxidase/dehydrogenase from A. thaliana under
a constitutive CaMV 35S promoter (Werner et al., 2001) and 9week-old WT N. tabacum L. cv. Samsun NN.
3. Eight- to 9-week-old tobacco plants (35S:P60) overexpressing
a maize b-glucosidase Zm-p60.1 naturally targeted to plastids
under a CaMV 35S promoter (Kiran et al., 2006) and WT Nicotiana
tabacum L. cv. Petit Havana SR1.
4. Eight- to 9-week-old tobacco plants (35S:ZOG1) harbouring
a trans-zeatin-O-glucosyltransferase gene from P. lunatus under a
constitutive CaMV 35S promoter (Martin et al., 2001a) and WT N.
tabacum L. cv. Wisconsin 38.
Chloroplast isolation
Intact chloroplasts were isolated and purified as described by Kiran
et al. (2006). The chloroplast fraction was recovered from the
homogenate of deribbed leaves in homogenization medium [0.33 M
sorbitol; 50 mM TRIS–HCl, pH 7.8; 0.4 mM KCl; 0.04 mM
Na2EDTA; 0.1% (w/v) bovine serum albumin; 1% (w/v) polyvinylpyrrolidone; 5 mM isoascorbic acid] by centrifugation, layered
on a Percoll density gradient [40% and 80% (v/v) Percoll solution
in resuspension medium (0.33 M sorbitol; 2 mM Na2EDTA; 1 mM
MgCl2; 1 mM MnCl2; 50 mM HEPES, pH 7.6)] and centrifuged
(1000 g; 15 min; 4 C). Intact chloroplasts were collected at the
interface of the gradient, diluted with medium, and centrifuged
Extraction and purification of IAA, ABA, and CK
The detailed procedure for hormone extraction, purification, and
quantification has been described in Kiran et al. (2006). IAA, ABA,
and CKs were extracted overnight at –20 C with Bieleski solvent
(Bieleski, 1964). [3H]IAA and [3H]ABA (Sigma, USA) and 12
deuterium-labelled CKs ([2H5]t-Z, [2H5]t-ZR, [2H5]t-Z7G, [2H5]tZ9G, [2H5]t-ZOG, [2H5]t-ZROG, [2H3]DHZ, [2H3]DHZR, [2H6]iP,
[2H6]iPR, [2H6]iP7G, and [2H6]iP9G; Apex Organics, UK) were
added as internal standards. The extracts were purified using Sep-Pak
C18 cartridges (Waters Corporation, Milford, MA, USA) and an
Oasis MCX mixed mode, cation exchange, reverse-phase column
(150 mg, Waters) (Dobrev and Kamı́nek, 2002). After a wash with
1 M HCOOH, IAA and ABA were eluted with 100% MeOH and
evaporated to dryness. Further, CK phosphates were eluted with 0.34
M NH4OH in water, and CK bases, ribosides, and glucosides were
eluted with 0.34 M NH4OH in 60% (v/v) MeOH. Phosphates were
converted to ribosides with alkaline phosphatase. IAA and ABA were
separated and quantified by two-dimensional high-performance liquid
chromatography (2D-HPLC) according to Dobrev et al. (2005).
Purified CK samples were analysed by an LCMS system consisting
of an HTS PAL autosampler (CTC Analytics, Switzerland), Rheos
2000 quaternary pump (FLUX, Switzerland) with Csi 6200 Series
HPLC oven (Cambridge Scientific Instruments, UK), and an LCQ
Ion Trap mass spectrometer (Finnigan, USA) equipped with an
electrospray. A 10 ll aliquot of sample was injected onto a C18
column (AQUA, 2 mm3250 mm35 lm, Phenomenex, USA) and
eluted with 0.0005% acetic acid (A) and acetonitrile (B). The HPLC
gradient profile was as follows: 5 min 10% B, then increasing to 17%
for 10 min, and to 46% for a further 10 min at a flow rate of 0.2 ml
min1. The column temperature was kept at 30 C. The effluent was
introduced into a mass spectrometer being operated in the positive
ion, full-scan MS/MS mode. Quantification was performed using a
multilevel calibration graph with deuterated CKs as internal standards. Plant hormone content is usually presented in picomoles per
gram fresh weight (FW) of plant tissue. Here, hormone concentration
is also expressed per unit of chlorophyll, allowing the comparison
with hormone levels in isolated chloroplasts. The differences were
estimated according to the value of the standard error.
Transmission electron microscopy
Leaf blade segments (2 mm32 mm3leaf thickness) were fixed for
2 h with 2.5% (v/v) glutaraldehyde in 0.1 M phosphate buffer,
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4 of 13 Polanská et al.
pH 7.2, followed by 2 h in 2% (w/v) osmic acid. After dehydration in
a graded ethanol series up to 100% ethanol, samples were embedded
via propylene oxide in low viscosity Spurr’s resin. The ultrastructure
was evaluated on transverse ultrathin sections of embedded objects
contrasted with a saturated solution of uranyl acetate in 70% (v/v)
aqueous ethanol, followed by a lead citrate solution treatment
according to Reynolds (1963) using a transmission electron
microscope (Philips Morgani) at an operating voltage of 70 kV.
Results
Hormone analyses
In order to investigate the role of hormone compartmentation and to study the interactions among plant hormones,
the contents of CKs, free IAA, and ABA were determined
in mature leaves and in isolated intact chloroplasts of four
different types of transgenic tobacco with altered CK
metabolism and in the corresponding WTs during the
vegetative stage of plant development. The integrity of the
analysed chloroplasts immediately after their isolation was
>90%, as checked by the latency of the activity of the
stromal enzyme glyceraldehyde-3-phosphate dehydrogenase.
Plants expressing isopentenyltransferase Sho
The total CK content in leaves after Sho induction by
dexamethasone was eight times that in the corresponding
WT (SR1) (Fig. 2A). The most elevated CK metabolites
were phosphates (predominantly iPMP) and N-glucosides
(especially iP7G). The level of physiologically active CK
bases and ribosides (predominantly of iP and iPR) increased to a lesser extent. No significant difference in the
content of CK-O-glucosides was observed. As CK levels
did not differ significantly among WT plants treated with
water, with dexamethasone in DMSO, or with DMSO
solution or non-induced Sho gene-carrying plants, only
the CK content of WT plants treated with water is shown.
The CK elevation in induced Sho leaves was lower when the
CK content was expressed per mg of chlorophyll, as the
transgenics had a higher chlorophyll content than the WT
(Fig. 2B). The difference between Sho transformants and the
WT was much lower in isolated chloroplasts (Fig. 2C) than
in leaves. Indeed, except for CK-O-glucosides, the percentage of leaf CKs sequestered in chloroplasts was lower in Sho
compared with WT (see supplementary Table 1 available at
JXB online). Nevertheless, the CK content of chloroplasts
from plants expressing Sho was more than twice as high as
that of the WT. The contents of individual CKs in leaves and
isolated chloroplasts are given in supplementary Table 2
available at JXB online.
On a FW basis, the higher CK content in Sho leaves
compared with the WT was accompanied by a slightly
higher content of IAA and ABA (Table 1). These differences were not significant when expressed per unit of
chlorophyll. Chloroplasts of Sho plants had lower IAA,
but higher ABA content than chloroplasts of WT plants.
Fig. 2. Cytokinin content (B+R, free bases and ribosides; NG,
N-glucosides; OG, O-glucosides; P, phosphates) in leaves (per fresh
weight A and per unit of chlorophyll B) and in isolated intact
chloroplasts (C) of transgenic dexamethasone-induced Sho (SHO,
black) and control SR1 (WT, striped) tobacco. Error bars display
the SE.
Plants expressing CK oxidase/dehydrogenase AtCKX3
The constitutive overexpression of AtCKX3 significantly
delayed development of transgenic plants; consequently,
hormone contents and chloroplast ultrastructure were
67
Cytokinins in chloroplasts of transgenic tobacco 5 of 13
Table 1. Endogenous contents of free IAA and ABA in leaves
and in isolated intact chloroplasts of transgenic dexamethasoneinduced Sho and WT (SR1) tobacco
Values represent the mean of two replicates (individual measurements
are shown in parentheses).
Sho
IAA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol mg1
chlorophyll)
ABA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol mg1
chlorophyll)
90.7 (117.9; 63.5)
73.7 (95.8; 51.6)
2.5 (2.8; 2.2)
WT (SR1)
71.2 (81.1; 61.3)
74.2 (84.4; 63.9)
3.4 (2.8; 4.0)
453.5 (749.1; 157.8)
368.7 (609.0; 128.3)
360.0 (628.1; 91.9)
375.0 (654.3; 95.7)
16.8 (18.7; 14.9)
12.8 (13.3; 12.2)
analysed in transgenic plants that were older than the
corresponding WT (SNN) plants. The total CK content
was reduced in leaves (Fig. 3A) to about one-half of that
in the WT. Particularly altered were the levels of CK-Nglucosides and, to a lesser extent, bases and ribosides. The
chlorophyll content was reduced in transgenic plants, so
that the differences between transgenic and WT tobacco
are reduced when expressed per unit of chlorophyll (Fig.
3B). The CK content was changed to a lesser extent in
isolated intact chloroplasts (Fig. 3C). Further, when
AtCKX3 and WT were compared, the percentage of leaf
CK bases and ribosides remained almost unchanged,
whilst those of phosphates sequestered in chloroplasts
increased; the values for CK-glucosides dropped significantly (see supplementary Table 1 available at JXB online).
The content of IAA was not significantly changed in
AtCKX3 leaves (Table 2) compared with WT when
expressed per FW, but it was higher when expressed per
unit of chlorophyll. The ABA level was lower in leaves of
transgenic tobacco. No significant changes of IAA or
ABA levels were observed in chloroplasts (Table 2).
Plants expressing b-glucosidase Zm-p60.1
Compared with WT (SR1) leaves, a trend towards an
increase in CK bases and ribosides and CK-N-glucosides
was observed in leaves (Fig. 4A, B) of tobacco overexpressing maize b-glucosidase Zm-p60.1 targeted to
plastids. A dramatic decrease in the levels of CK-Oglucosides remained restricted to the CK pool found in
chloroplasts (Fig. 4C). Interestingly, a moderate decrease
was also observed for CK-N-glucosides in the chloroplast
fraction. The trend towards an increase in CK bases and
ribosides observed in whole leaf extracts was also seen in
the chloroplast fraction. Accordingly, the percentage of
leaf CK-O- and N-glucosides sequestered in chloroplasts
was significantly lower in the transgenic plants than in
WT plants, while the values changed only marginally for
Fig. 3. Cytokinin content (B+R, free bases and ribosides; NG, Nglucosides; OG, O-glucosides; P, phosphates) in leaves (per fresh
weight A and per unit of chlorophyll B) and in isolated intact
chloroplasts (C) of transgenic 35S:AtCKX3 (CKX, black) and control
SNN (WT, striped) plants. Error bars display the SE.
CK bases and ribosides, and for phosphates (see supplemetary Table 1 available at JXB online).
The IAA measurements revealed a significantly lower
level of IAA in leaves and in chloroplasts of transgenic
plants compared with the WT IAA level (Table 3). The
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6 of 13 Polanská et al.
Table 2. Endogenous contents of free IAA and ABA in leaves
and in isolated intact chloroplasts of transgenic 35S:AtCKX3
and WT (SNN) tobacco
Values represent the mean of two replicates (individual measurements
are shown in parentheses).
35S:AtCKX3
IAA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol
mg1 chlorophyll)
ABA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol
mg1 chlorophyll)
125.0 (177.4; 72.6)
200.5 (279.1; 121.8)
5.2 (4.3; 6.1)
WT (SNN)
122.1 (147.3; 96.9)
114.4 (151.6; 77.2)
5.2 (9.9; 0.4)
359.5 (447.7; 271.3) 1102.3 (822.9; 1381.6)
579.9 (704.4; 455.4) 974.1 (847.2; 1100.9)
68.0 (44.0; 92.0)
58.8 (106.6; 10.9)
ABA content was reduced in Zm-p60.1 leaves compared
with WT (Table 3). In chloroplasts, the difference in ABA
level was less pronounced.
Plants expressing zeatin-O-glucosyltransferase ZOG1
When a ZOG1 gene was constitutively expressed in
tobacco, CK-O-glucosides accumulated in leaves (Fig.
5A). The level of t-ZOG was two orders of magnitude
higher in leaves of transformants than in those of WT
(W38). No significant difference in the content of CK-Nglucosides, and only marginal reduction of the remaining
CK forms was observed. These differences reduced
further when the CK content was expressed per unit of
chlorophyll (Fig. 5B), while that of CK-O-glucosides was
more profound, as the chlorophyll content was lower in
ZOG1 than in WT. By contrast, in chloroplasts, very low
levels of CK-O-glucosides were found (Fig. 5C). The
predominant CK forms were bases and ribosides, the
contents of which, as well as that of CK phosphates, were
higher in ZOG1 chloroplasts than in WT chloroplasts.
Indeed, out of the transgenic plants analysed, only in the
plants overexpressing ZOG1 was the percentage of leaf
CK bases and ribosides sequestered in chloroplasts
significantly higher in the transgenic plants than in WT
plants (see supplementary Table 1 available at JXB online).
In leaves of transgenic plants, a 2-fold higher IAA
content was detected (Table 4). The ABA level was not
significantly affected in ZOG1 leaves on a FW basis, but
was higher when expressed per unit of chlorophyll. IAA
and ABA contents were higher in ZOG1 chloroplasts than
in WT chloroplasts.
Fig. 4. Cytokinin content (B+R, free bases and ribosides; NG, Nglucosides; OG, O-glucosides; P, phosphates) in leaves (per fresh
weight A and per unit of chlorophyll B) and in isolated intact
chloroplasts (C) of transgenic 35S:Zm-p60.1 (P60, black) and control
SR1 (WT, striped) tobacco. Error bars display the SE.
enhancers (Fig. 6A, detailed view in Fig. 6B) as well as
the 35S promoter. However, some chloroplasts from these
plants did not show apparent changes in the structure
compared with the WT (Fig. 6C). Chloroplasts from Shooverexpressing plants contained more plastoglobuli than
WT chloroplasts (Fig. 6D). The crystalloids were never
present in chloroplasts of WT plants cultivated in vitro.
Ultrastructure observations
The most striking anomaly in chloroplast ultrastructure
was the occasional occurrence of crystalloid structures in
chloroplasts of in vitro-cultivated plants harbouring the
Sho gene under transcriptional control of four 35S
69
Cytokinins in chloroplasts of transgenic tobacco 7 of 13
Table 3. Endogenous contents of free IAA and ABA in leaves
and in isolated intact chloroplasts of transgenic 35S:Zm-p60.1
and WT (SR1) tobacco
Values represent the mean of two replicates (individual measurements
are shown in parentheses).
35S:Zm-p60.1
IAA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol
mg1 chlorophyll)
ABA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol
mg1 chlorophyll)
85.7 (95.1; 76.2)
60.3 (66.9; 53.6)
0.8 (1.4; 0.2)
WT (SR1)
144.0 (146.5; 141.5)
135.2 (137.5; 132.8)
1.2 (2.1; 0.3)
481.6 (486.9; 476.3)
338.7 (342.4; 334.9)
839.0 (785.9; 892.1)
787.5 (737.6; 837.4)
23.7 (21.5; 25.8)
26.3 (40.4; 12.1)
After induction of the Sho gene expression, no crystalloids
were found, only an increased grana stacking was
observed (Fig. 6E). The crystalloids did not appear in
chloroplasts of WT plants treated either with water (Fig.
6F), with dexamethasone in DMSO, or with DMSO
solution, or in chloroplasts of non-induced plants carrying
the Sho gene.
Representative chloroplasts from the other types of
transgenic tobacco and corresponding WTs are shown in
Fig. 7. The most obvious feature observed was the different starch accumulation. The chloroplasts of tobacco
overexpressing AtCKX3 (Fig. 7A) and Zm-p60.1 (Fig. 7C)
have less starch inclusions than chloroplasts of WT SNN
(Fig. 7B) and SR1 (Fig. 7D), respectively. In contrast, the
chloroplasts of ZOG1 transformants (Fig. 7E) contained
more starch inclusions than WT W38 chloroplasts (Fig.
7F). With increasing starch content, the shape of chloroplasts altered from lens-shaped to more loaf-like.
Discussion
Hormone analyses
In order to elucidate the degree of chloroplast CK
autonomy, the impact of altered CK metabolism on the
CK pool in isolated intact chloroplasts and in bulk leaf
tissue was compared. Bearing in mind hormone cross-talk,
the effect of altered CK metabolism on IAA and ABA
levels was also followed. The presence of these hormones
in chloroplasts was demonstrated in earlier studies through
the use of immunocytochemical (Sossountzov et al., 1986;
Ohmiya and Hayashi, 1992; Kärkönen and Simola, 1999)
or fractionation techniques, followed by HPLC or LCMS
analysis (Sandberg et al., 1990; Benková et al., 1999).
Chloroplasts were found to be the compartment where
most of the ABA in leaf tissue is formed by the methylerythritol phosphate pathway (Milborrow and Lee, 1998).
Fig. 5. Cytokinin content (B+R, free bases and ribosides; NG, Nglucosides; OG, O-glucosides; P, phosphates) in leaves (per fresh
weight A and per unit of chlorophyll B) and in isolated intact
chloroplasts (C) of transgenic 35S:ZOG1 (ZOG, black) and control
W38 (WT, striped) tobacco. Error bars display the SE.
This pathway also provides most of the prenyl group of
CKs (Kasahara et al., 2004). Nordström et al. (2004)
demonstrated de novo CK synthesis in Arabidopsis shoots
and tobacco leaves, and suggested that the presence of
chloroplasts might be a prerequisite for the iPMPindependent pathway of CK biosynthesis.
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Plants expressing isopentenyltransferase Sho
Numerous studies of plants transformed with the ipt gene,
encoding an IPT from Agrobacterium tumefaciens, have
demonstrated enhanced biosynthesis of endogenous CKs,
especially of the Z-type (Redig et al., 1996; Motyka
et al., 2003). Recently, genuine plant IPT genes (AtIPT1–
9) were identified in the A. thaliana genome (Kakimoto,
2001; Takei et al., 2001) and their homologue Sho in P.
hybrida (Zubko et al., 2002). The overexpression of the
plant homologue AtIPT8 (Sun et al., 2003), as well as Sho
(Zubko et al., 2002), in contrast to the bacterial ipt, leads
to profound accumulation of the iP-type of CKs. A
dexamethasone-inducible gene expression system (pOp6/
LhGR; Šámalová et al., 2005) was used to drive Sho
expression in transgenic tobacco. Following Sho induction
in leaf extracts, major increases were observed in iP7G
and iPMP, which is in line with previous results reported
for petunia and tobacco overexpressing Sho constitutively
(Zubko et al., 2002). The CK content was also increased
in chloroplasts, although to a relatively lower extent. This
is in accordance with preliminary results of feeding experiments (data not shown) showing that this IPT preferentially utilizes mevalonate as a substrate, indicating its
cytosolic localization. In a recent analysis of Pssu-ipt
tobacco (Synková et al., 2006), similarly to Sho plants,
Table 4. Endogenous contents of free IAA and ABA in leaves
and in isolated intact chloroplasts of transgenic 35S:ZOG1 and
WT (W38) tobacco
Values represent the mean of two replicates (individual measurements
are shown in parentheses).
35S:ZOG1
IAA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol
mg1 chlorophyll)
ABA
Leaves (pmol g1 FW)
Leaves (pmol mg1
chlorophyll)
Chloroplasts (pmol
mg1 chlorophyll)
88.8 (102.1; 75.5)
77.2 (88.8; 65.6)
1.5 (1.8; 1.1)
528.1 (662.4; 393.8)
459.2 (575.9; 342.4)
13.1 (20.9; 5.2)
WT (W38)
44.2 (38.4; 50.0)
24.0 (20.8; 27.1)
1.0 (0.5; 1.4)
523.3 (566.9; 479.6)
284.0 (307.7; 260.3)
10.1 (15.0; 5.2)
Fig. 6. Transmission electron micrographs of representative chloroplast cross-sections taken from the intact leaves of tobacco expressing the Sho
gene and four 35S enhancer elements (A, C) with detail of crystalloid (B) and control SR1 (D) tobacco growing under sterile conditions; and
chloroplast cross-section detail taken from the leaf of transgenic dexamethasone-induced Sho (E) and control SR1 (F) tobacco cultivated in
a greenhouse. C, crystalloid; CW, cell wall; GT, granal thylakoids; IT, intergranal thylakoids; M, mitochondrion; P, plastoglobuli; S, stroma; SI,
starch inclusion. Bar¼2 lm in A, C and D; and 500 nm in B, E and F.
71
Cytokinins in chloroplasts of transgenic tobacco 9 of 13
Fig. 7. Transmission electron micrographs of representative chloroplast cross-sections taken from the intact leaves of transgenic 35S:AtCKX3 (A) and
control SNN (B), transgenic 35S:Zm-p60.1 (C) and control SR1 (D), transgenic 35S:ZOG1 (E) and control W38 (F) tobacco. CW, cell wall; GT,
granal thylakoids; IT, intergranal thylakoids; M, mitochondrion; P, plastoglobuli; S, stroma; SI, starch inclusion. Bar¼2 lm.
CK levels increased in chloroplasts to much lower levels
compared with leaf extracts, indicating that the main CK
pool accumulates outside chloroplasts in both Z- and iPtype CK-overproducing plants.
The content of IAA in leaves of Sho-expressing plants
did not differ significantly from that in WT, whereas the
chloroplasts contained a lower level of IAA than WT
chloroplasts. In ipt tobacco, Eklöf et al. (1997, 2000)
found lower levels of IAA and reduced rates of IAA
synthesis and turnover. This has led to the suggestion that
CKs might down-regulate IAA levels. Recently, a decrease
in auxin biosynthesis following an increase in CK levels
achieved by induction of ipt was observed in Arabidopsis,
however, with a significant delay. Thus, the decrease was
hypothesized to be mediated through an altered development rather than through a direct CK–auxin cross-talk
(Nordström et al., 2004).
A higher ABA content was found in chloroplasts and in
leaves (when expressed per FW) of Sho plants compared
with WT. After overexpression of bacterial ipt, both
a decrease (Synková et al., 1999; Chang et al., 2003) and
an increase (Macháčková et al., 1997) in ABA content
was reported.
Plants expressing CK oxidase/dehydrogenase AtCKX3
The CK oxidase/dehydrogenase selectively cleaves unsaturated N6 side chains of Z, iP, and their corresponding
ribosides, while CK phosphates, O-glucosides, and CKs
with saturated side chains are not substrates for CK
oxidase/dehydrogenase (Armstrong, 1994). It was shown
that overexpression of AtCKX genes in tobacco and
Arabidopsis plants resulted in a reduced content of
endogenous CKs and a strongly altered phenotype, with
dwarfed shoot habit, enhanced root growth, and delayed
flowering and senescence (Werner et al., 2001, 2003).
More abundant Z-derived CKs were more strongly reduced than were iP derivatives in 35S:AtCKX1 and
35S:AtCKX2 Arabidopsis, in contrast to 35S:AtCKX1 and
35S:AtCKX2 tobacco where the more profound changes
were in iP-type CKs. To our knowledge, the CK content
has never been measured in any 35S:AtCKX3 transgenic
plant. These plants are phenotypically identical to
35S:AtCKX1 and the enzyme is ultimately targeted to the
same compartment, the vacuole, as proved by in planta
experiments exploring an AtCKX–green fluorescent protein (GFP) fusion. Nevertheless, the apparent Km value for
72
10 of 13 Polanská et al.
Lower IAA levels were determined in leaves as well as
isolated chloroplasts of Zm-p60.1 plants compared with
WT. This agrees with previous results, where a steeper fall
in IAA gradient from high to low from youngest leaves
downward, and a decreased IAA content in the apex and
first internodes in Zm-p60.1 tobacco plants was found
(Kiran et al., 2006). CK effects might be partly mediated
by elevation of ethylene biosynthesis (Genkov et al.,
2003). Ethylene treatment was shown to stimulate the
oxidative decarboxylation of IAA (Winer et al., 2000).
Thus, the elevated contents of active CKs in Zm-p60.1
plants could lead to lower levels of free IAA.
ABA measurements revealed a lower ABA content in
Zm-p60.1 leaves than in WT. This finding is in contrast to
those of a previous study (Kiran et al., 2006), when the
tendency of increased ABA accumulation especially in
older leaves was observed.
AtCKX3 against iP was 14 times higher than that for
AtCKX1 (Werner et al., 2003). It has been shown that the
ectopic expression of AtCKX3 reduced the CK levels in
leaves of adult tobacco plants and also influenced the CK
levels in chloroplasts. In leaves, the most strongly reduced
CK metabolites were N-glucosides. It is obvious that after
a tremendous increase of the activity of an important CK
catabolic pathway (side chain cleavage by oxidase/dehydrogenase), the activity of the other CK deactivation
pathway (N-glucosylation) was suppressed. In young
tobacco seedlings with very low N-glucosylation activity,
oxidase/dehydrogenase overexpression had only a marginal effect on the level of N-glucosides (Werner et al.,
2001). This discrepancy could be explained by the different developmental stages of the analysed tobacco plants.
In 35S:AtCKX1 and 35S:AtCKX2 Arabidopsis seedlings,
Z9G and iPG were reduced compared with WT.
No significant differences were observed between the
IAA levels, expressed on a FW basis, in leaves of adult
AtCKX3 and WT tobacco plants. This finding contrasts
with the results of Werner et al. (2003), who found reduced
levels of IAA in 35S:AtCKX1 and 35S:AtCKX2 Arabidopsis seedlings. However, the authors speculate that CKs need
not directly regulate IAA metabolism, but that the different
tissue composition, i.e. reduced size of the shoot apical
meristem with fewer meristematic cells and reduced cell
production in leaves, might lead to a lowering of IAAproducing shoot tissue in transgenic plants.
A lower ABA level was found in AtCKX3 leaves
compared with WT leaves, which might be linked to
prolonged life span and retarded senescence of transgenic
plants. Brugière et al. (2003) demonstrated induction of
maize CKX1 gene expression in leaf discs by ABA,
suggesting a role for this hormone in lowering CK
concentrations under different abiotic stresses.
Plants expressing trans-zeatin-O-glucosyltransferase
ZOG1
Tobacco transformed with the ZOG1 gene from P. lunatus
under the control of the CaMV 35S promoter was
constructed by Martin et al. (2001a). They found
a strongly elevated t-ZOG content with no substantial
changes in the level of other CK metabolites. In the
present study, an order of magnitude higher accumulation
of ZOG in ZOG1 leaves was detected, which may have
been caused by the analysis of older material in the
present case and different cultivation conditions. The
elevated CK-O-glucoside concentrations in bulk leaf
tissue did not significantly influence the content of these
metabolites in intact chloroplasts isolated from the transgenic plants. This indicates that CK-O-glucosides are
accumulated mainly outside chloroplasts, which agrees
with the previous observation that transgenic plants overexpressing Zm-p60.1 can accumulate high levels of t-ZOG
when grown on a medium supplemented with t-Z despite
high Zm-p60.1 b-glucosidase activity in chloroplasts
(Kiran et al., 2006). The vacuole has been proposed as
the main storage compartment for CK-O-glucosides,
based on feeding experiments in Chenopodium rubrum
(Fußeder and Ziegler, 1988).
A 2-fold higher IAA content was observed in ZOG1
leaves compared with WT. Martin et al. (2001a) reported
that IAA does not serve as a substrate for zeatin-Oglucosyltransferase and therefore excluded changes in
IAA glucosylation as triggering the observed variations in
morphology of transformants (e.g. aerial roots, multiple
shoots, more compact stature than WT).
Plants expressing b-glucosidase Zm-p60.1
Maize b-glucosidase Zm-p60.1 was the first plant enzyme
shown to release active CKs from CK-O- and N3-glucosides in vivo (Brzobohatý et al., 1993). Recently, it was
shown that ectopic overexpression of Zm-p60.1 can
perturb CK homeostasis in transgenic tobacco (Kiran
et al., 2006). Higher levels of CK bases and ribosides in
upper leaves and internodes of transgenic plants were
found. The present results are consistent with those of the
previous report. Moreover, it was found that a tendency to
higher accumulation of CK bases and ribosides is also
apparent in chloroplasts. The dramatic decrease in CK-Oglucoside levels reported here for chloroplasts isolated
from Zm-p60.1 plants corresponded well with the plastid/
chloroplast location of Zm-p60.1 (Kristoffersen et al.,
2000) and a strong decrease in apparent Km for t-ZOG
when the values obtained for the enzyme purified in vitro
and present in isolated chloroplasts were compared (Kiran
et al., 2006).
Ultrastructure observations
Considerable changes were observed in the chloroplast
ultrastructure of tobacco constitutively expressing the Sho
gene, including an increased number of plastoglobuli and
the appearance of crystalloids in plants grown in vitro.
73
Cytokinins in chloroplasts of transgenic tobacco 11 of 13
These crystalloids were not found in other CK metabolism
transformants analysed in this work and their corresponding WTs. In another type of transgenic tobacco, with
strongly elevated endogenous CKs, Pssu-ipt, anomalies in
chloroplast ultrastructure together with crystalloid occurrence were also found, and the distinct crystalline
structures were hypothesized to be formed by 2D crystals
of light-harvesting complex proteins (Synková et al.,
2006).
When dexamethasone-induced Sho plants were analysed, increased grana stacking was noticed. Likewise, an
increase in grana stacking was observed in chloroplasts of
transgenic ipt tobacco (Čatský et al., 1993) and after CK
treatment (Wilhelmová and Kutı́k, 1995; Salopek-Sondi
et al., 2002).
Except for Sho tobacco, marked changes in the
ultrastructure of chloroplasts were not found in the other
CK transformants analysed. Only a difference in starch
accumulation, associated with a change in chloroplast
shape, was perceived. This finding is in accordance with
data showing that exogenous CK application may increase
starch accumulation in chloroplasts (Wilhelmová and
Kutı́k, 1995; Stoynova et al., 1996). The involvement of
endogenous CKs in starch formation and amyloplast
development was proved by Miyazawa et al. (2002) in
tobacco BY-2 cells.
In conclusion, a highly non-uniform distribution of
CKs was found between chloroplasts and bulk leaf tissue
in transgenic tobacco lines with several distinct alterations of CK metabolism, indicating that chloroplasts are
relatively independent organelles with respect to the
regulation of CK metabolism. Thus, it is evident that the
estimation of overall CK content in plant tissue is not
sufficient to assess its level at a particular site of action
and that different compartmentation of individual CK
metabolites should be taken into account. CK glucosides
do not usually accumulate in chloroplasts, while active
forms of CKs, bases, and ribosides are often found in
them. In line with previous reports, the present data
suggest a complex nature of mutual interactions between
CKs, IAA, and ABA that apparently cannot be described
by a simple model based on the current state of the art
in this field. Thus, Werner et al. (2003) and Nordström
et al. (2004) suggested that the effect of CKs on IAA
levels might be mediated through altered development.
Very variable data can be found in the literature on the
interaction of CKs and ABA. CKs are known to
counteract many processes induced by ABA (e.g.
senescence or stomata closure). Elevated CK content
thus might be balanced by higher ABA levels, which is
also the case for Sho plants. However, the CK–ABA
cross-talk does not seem to be straightforward. It
probably strongly depends on the actual physiological
conditions of the studied material including environmental factors and/or tissue specificity.
Supplementary data
Supplementary data available at JXB online are Table 1
showing the percentage of leaf CKs sequestered to
chloroplasts and Table 2 listing the CK contents in leaves
and isolated chloroplasts.
Acknowledgements
We are very grateful to Professor Peter Meyer (Sho), Professor
Thomas Schmülling (AtCKX3), and Professors Machteld and David
Mok (ZOG1) for kind donation of transgenic material, and to
Professor David Morris for critical reading of the manuscript. This
work was supported by grant nos 206/03/0369 and 206/06/1306
(Grant Agency of the Czech Republic) and IAA600040612 (Grant
Agency of the Academy of Sciences of the Czech Republic).
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Plant, Cell and Environment (2008) 31, 341–353
doi: 10.1111/j.1365-3040.2007.01766.x
The role of cytokinins in responses to water deficit
in tobacco plants over-expressing trans-zeatin
O-glucosyltransferase gene under 35S or
SAG12 promoters
MARIE HAVLOVÁ1,2, PETRE I. DOBREV1, VÁCLAV MOTYKA1, HELENA ŠTORCHOVÁ1, JIŘÍ LIBUS1,
JANA DOBRÁ1,2, JIŘÍ MALBECK1, ALENA GAUDINOVÁ1 & RADOMIRA VANKOVÁ1
1
Institute of Experimental Botany AS CR, Rozvojova 263, 165 02 Prague 6, Czech Republic and 2Department of
Biochemistry, Faculty of Natural Sciences, Charles University, Hlavova 2030/8, 128 43 Prague 2, Czech Republic
enable plants to cope with water deficit include fast
responses, such as stomatal closure, and longer-term
changes in metabolism, growth and development. The latter
include the accelerated senescence and abscission of lower
leaves, as well as changes in source–sink relationships and
the partitioning of dry matter between root and shoot
systems (Rodrigues, Pacheco & Chaves 1995). Both fast
stress responses and transduction of stress-related signals
into changes in specific gene expression involve the mediation by phytohormones.
Most studies of the part played by plant hormones in
stress physiology have focused on the role of abscisic acid
(ABA). An elevation in ABA concentration in plants
exposed to various stresses has frequently been reported
(see Mansfield & McAinsh 1995). In response to drought,
ABA reduces the transpiration rate by inducing stomatal
closure. In addition, at least two of five signal transduction
pathways known to regulate drought- and cold-inducible
genes involve ABA action (Shinozaki & YamaguchiShinozaki 1996; Davies, Kudoyarova & Hartung 2005).
Precise control of individual processes involved in
responses to stress requires both positive and negative
regulation. For example, many ABA-mediated physiological processes induced by water deficit, including closure of
the stomata and acceleration of leaf senescence, are counteracted by cytokinins (CKs; for a review, see Pospíšilová &
Dodd 2005). CKs increase stomatal aperture and/or delay
ABA-induced stomatal closure (Stoll, Loveys & Dry 2000).
It has been suggested (Pospíšilová & Dodd 2005) that in
longer-term responses to stress, hormones such as ABA and
CK may function to regulate the production, metabolism
and distribution of metabolites essential for stress survival
and recovery.
A role for phytohormones in plant stress responses can
be inferred from plant reactions to exogenously applied
natural hormones or their synthetic analogues. Exogenous
CKs have been reported to increase tolerance to mild stress
and to speed up recovery (e.g. Itai, Benzioni & Munz 1978).
Applied CKs reduce the negative effects of water deficit on
chlorophyll and carotenoid content and on photosynthetic
ABSTRACT
The impact of water deficit progression on cytokinin (CK),
auxin and abscisic acid (ABA) levels was followed in upper,
middle and lower leaves and roots of Nicotiana tabacum
L. cv. Wisconsin 38 plants [wild type (WT)]. ABA content
was strongly increased during drought stress, especially
in upper leaves. In plants with a uniformly elevated total
CK content, expressing constitutively the trans-zeatin
O-glucosyltransferase gene (35S::ZOG1), a delay in the
increase of ABA was observed; later on, ABA levels were
comparable with those of WT.
As drought progressed, the bioactive CK content in
leaves gradually decreased, being maintained longer in the
upper leaves of all tested genotypes. Under severe stress
(11 d dehydration), a large stimulation of cytokinin oxidase/
dehydrogenase (CKX) activity was monitored in lower
leaves, which correlated well with the decrease in bioactive
CK levels. This suggests that a gradient of bioactive CKs in
favour of upper leaves is established during drought stress,
which might be beneficial for the preferential protection of
these leaves.
During drought, significant accumulation of CKs
occurred in roots, partially because of decreased CKX
activity. Simultaneously, auxin increased in roots and lower
leaves. This indicates that both CKs and auxin play a role in
root response to severe drought, which involves the stimulation of primary root growth and branching inhibition.
Key-words: abscisic acid; auxin; cytokinin; drought; tobacco.
INTRODUCTION
Under the selection pressures resulting from their sessile
growth habit in a variable environment, plants have evolved
a considerable capacity to cope with adverse environmental
conditions. Water availability represents a major limiting
factor for plant growth and development. Strategies that
Correspondence: R. Vanková. Fax: +420 225 106 446; e-mail:
[email protected]
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd
341
77
342 M. Havlová et al.
activity (e.g. Metwally, Tsonev & Zeinalov 1997) and
improve the recovery of stomatal conductance and net photosynthesis after rehydration (Rulcová & Pospíšilová 2001).
Moreover, the transcription of many stress-induced genes is
stimulated by CKs (Hare & Van Staden 1997).
Modulation of hormone levels may also be achieved by
the over-expression of genes coding for their synthesis or
metabolism. In the case of CKs, a biosynthetic gene, ipt
(coding for the Agrobacterium tumefaciens isopentenyltransferase), has been frequently used. In constitutive transformants, the highly elevated bioactive CK content may
itself, unfortunately, lead to substantial changes in plant
morphology, which in turn may complicate the interpretation of the potential role of CKs in the response of plants to
stress conditions. To minimize this limitation, the ipt gene
expression was given under the control of the highly
senescence-specific SAG12 promoter (Gan & Amasino
1995). In these transformed plants, the activation of ipt transcription is triggered mainly in senescing leaves, and plants
develop normally. SAG12::ipt plants have been used to
evaluate the effect of CKs on leaf senescence (Gan &
Amasino 1995; Jordi et al. 2000), photosynthesis and nitrogen partitioning (Jordi et al. 2000), flooding tolerance
(Zhang et al. 2000; Huynh et al. 2005), seed germination,
and response to mild water stress (Cowan et al. 2005; but
see further).
An alternative approach has been to over-express the
gene coding for trans-zeatin O-glucosyltransferase
(ZOG1), which results in the elevation of CK O-glucoside
content. ZOG1 gene was isolated from Phaseolus lunatus
(Martin, Mok & Mok 1999). Tobacco plants transformed
with ZOG1 under the control of the constitutive (35S) promoter were prepared by Martin et al. (2001). In 35S::ZOG1
plants, the total CK content (especially of CK O-glucosides)
was greatly increased, while the level of bioactive CKs
(bases, ribosides) was not changed significantly (Martin
et al. 2001). CK O-glucosides represent CK storage forms,
as they can be converted to bioactive CKs by the action of
b-glucosidases (e.g. Campos et al. 1992).
A study of the role of CKs in ontogenetic and stressinduced transitions in the tobacco plants was recently
published by Cowan et al. (2005). They proposed that
senescence-dependent ontogenetic and water stressinduced transitions in plants may be influenced by CK regulation of sink–source relations. This proposal was based,
inter alia, on a comparison of the CK and soluble sugar
content and distribution, pigment (chlorophylls and carotenoids) distribution, ABA content, and photosynthetic
activity in leaves of different ages in wild-type (WT) plants
and plants transformed with the SAG12::ipt gene. However,
these authors did not provide data on the CK and soluble
sugar composition of water-stressed plants of the two genotypes. Thus, a direct evidence for their hypothesis that
CK-mediated, stress-induced changes in sink–source relationships play a role in the plant response to water deficit
has been still missing.
In the work described here, we sought to clarify the role of
CKs in the responses of plants to water deficits.We compared
the impact of constitutive and senescence-induced ZOG1
over-expression (using the 35S and SAG12 promoters,
respectively) on CK levels, distribution and metabolism, on
leaf production and senescence, and on indolyl-3-acetic acid
(IAA) and ABA content during the onset and progression of
water deficit in tobacco plants and during the early stages of
their recovery following rehydration. Measurement of the
changes in ABA levels was undertaken because the ABA
content is known to correlate well with the degree of stress to
which a plant is exposed (Mansfield & McAinsh 1995).
Similarly, as auxins and CKs interact in the regulation of
many physiological processes, including those likely to be
involved in the recovery from stress, the effects of water
deficit on IAA levels were also determined. Furthermore,
as the transgenic plants accumulate high levels of CK
O-glucosides, we also investigated the potential contribution
of this readily mobilizable CK reserve to the pool of bioactive CKs following the onset of stress.
MATERIALS AND METHODS
Plant material and stress application
WT tobacco (Nicotiana tabacum L. cv. Wisconsin 38) and
transgenic SAG12::ZOG1 and 35S::ZOG1 tobacco plants
(for detailed information, see Martin et al. 2001) were sown
in sterilized soil (Garden Substrate B; Rašelina, Soběslav,
Czech Republic; pot volume 350 mL) without added fertilizer. Plants were raised in a growth chamber (Sanyo MLR
350H; Sanyo, Osaka, Japan) for 6 weeks on a 16 h photoperiod at 1000 mmol m-2 s-1, day/night temperature of 25/23 °C
and relative humidity ca. 80%. Water was permanently
available to plants. After 6 weeks, half of the plants of each
genotype were transferred into another growth chamber
under the same light and temperature regimes, but with the
relative humidity decreased to 35%. Plants of the same leaf
number were selected for the stress experiments in order
to exclude the variability within the population (highest in
the case of 35S::ZOG1 plants). The surface of the soil was
covered with a film impervious to water vapour (Parafilm;
Pechiney, Menasha, WI, USA). No further water was
applied, and the impact of water deficit on various plant
parameters was determined 1 d (‘mild stress’), 6 d (‘moderate stress’) and 11 d (‘severe stress’) after the onset of dehydration. The stressed plants were then watered, and the
same parameters were measured 1 d later (‘recovery’).
Samples of upper leaves (the two youngest unfolded),
lower leaves (the two lowest, still viable) and the middle
ones (two in between the upper and lower ones) were collected, cut into pieces and immediately frozen in liquid
nitrogen (Linde Gas, Pardubice, Czech Republic) (after
removal of the main vein). The roots were shaken to
remove the soil, washed briefly with cold tap water (ca.
1 min), dried with filter paper and frozen in liquid nitrogen.
For each treatment, a total of three (control) or six (stress)
plants were harvested (collected in three independent
experiments); the presented results are the means of the
three samples.
© 2008 The Authors
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78
Role of cytokinins in responses of tobacco to water deficit 343
(a)
RWC was measured using individual cut leaves (upper and
middle ones), which were weighed [fresh mass (FM)], saturated with water in the beakers [water saturated mass
(SM)] and dried at 88 °C [dry mass (DM)]. RWC was
counted as: RWC (%) = 100 - [(SM - FM)/(SM - DM)] ¥
100 (Salisbury & Ross 1992).
12
10
150
8
6
100
4
50
2
Leaf number
200
Determination of relative water content (RWC)
14
WT
RNA extraction and reverse transcription
0
200
Total RNA from the leaves or roots was extracted by means
of the RNeasy Plant Kit (Qiagen, Hilden, Germany) and
treated with DNAse I on column. One microgram of RNA
and random hexamer oligonucleotides (4 mg) or oligo dT
primers (500 ng) were heated 5 min at 65 °C, chilled on ice
and mixed with buffer, 0.5 mL of Protector RNase Inhibitor
(Roche, Mannheim, Germany), 2 mL of 10 mM dNTPs and
10 units of Transcriptor Reverse Transcriptase (Roche). The
first strand cDNA was synthesized at 55 °C for 30 min,
which was preceded by 10 min at 25 °C if random hexamers
were used.
14
SAG:ZOG
12
10
150
8
6
100
4
50
Leaf number
(b)
2
0
0
200
(c)
35S:ZOG
14
qRT-PCR
12
First strand cDNA was 10¥ or 20¥ diluted and qRT-PCR
was performed using the FastStart DNA MasterPLUS SYBR
Green I Kit (Roche) in a final volume of 10 mL with 300 nM
of each of the high-performance liquid chromatography
(HPLC)-purified primers (Metabion, Plantegg, Martinsried,
Germany).The LightCycler 1.2 (Roche) was programmed as
follows: 10 min of initial denaturation at 95 °C, then 45 cycles
for 10 s at 95 °C, 4 s at the appropriate annealing temperature (Supplementary Table S1; annealing temperature determined by experiment), and 7 s at 72 °C. cDNA derived from
calibrator RNA was included into each LighCycler run to
correct for run-to-run differences. PCR efficiencies (Supplementary Table S1) were estimated from the calibration
curves generated from the serial dilution of cDNAs. The
calibrator normalized relative ratio of the relative amount of
the target and reference gene was calculated as follows:
10
150
8
6
100
4
50
Leaf number
% of plant initial weight
0
2
0
0
1 3 5 7 9 11
1 3 5 7 9 11
Control
Drought
Time (days)
Figure 1. Time course of the water loss and leaf number
(•) in control and drought-stressed (a) wild-type (WT),
(b) SAG12::ZOG1 and (c) 35S::ZOG1 plants. The plants were
cultivated in soil (pot size 350 mL). The age of the plants was
6 weeks at the beginning of the experiment. Results from three
independent experiments are presented (mean values ⫾ SE are
shown).
ET C pT (C )− C pT (S) × ER C pR (S)− C pR (C )
where ET/ER is the efficiency of target/reference amplification, CpT/CpR is the cycle number at the target/reference
detection threshold (crossing point), S is the sample and C
is the calibrator.
ZOG1 transcript levels were normalized against Act9
(Volkov, Panchuk & Schoffl 2003) and 18S rRNA (in case of
random primed cDNA).
The degree of stress was followed via the decrease in soil
moisture (estimated as a loss of pot weight). In the stressed
plants, the greatest loss of water occurred during the first
24 h (Fig. 1). The weight of the control pots did not change
significantly during the experiments. The time course of
fresh weight/dry weight (FW/DW) ratio (mean values) was:
control plants – all leaves ca. 10.6, roots 5.6; mild stress – all
leaves 9.2, roots 5.3; moderate stress – leaves (upper,
middle, lower) 6.8, 6.8, 5.6, roots 4.9; severe stress – leaves
5.3, 5.0, 4.0, roots 4.2; rehydration – leaves 7.9, 8.5, 8.0,
roots 4.6.
Plant hormone extraction and purification
Phytohormones were extracted and purified according to
Dobrev & Kaminek (2002). For analyses of endogenous
CKs, 50 pmol of each of the following 17 deuteriumlabelled standards were added - [2H5]Z, [2H5]ZR, [2H5]Z7G,
[2H5]Z9G, [2H5]ZOG, [2H5]ZROG, [2H6]iP, [2H6]iPR,
© 2008 The Authors
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79
344 M. Havlová et al.
[2H6]iP7G,
[2H6]iP9G,
[2H3]DHZ,
[2H3]DHZR,
[2H3]DHZ9G, [2H7]DHZOG, [2H5]ZMP, [2H3]DHZMP and
[2H6]iPMP (Apex Organics, Honiton, UK). cis-Zeatin
derivatives were determined using the retention times and
mass spectroscopy (MS) spectra of unlabelled standards
and the response ratio of their trans-zeatin counterparts.
The term ‘(total) CK pool’ is used for the sum of all isoprenoid CK metabolites. Tritiated internal standards were
used for the determination of auxin (3[5(n)-3H] IAA;Amersham Pharmacia Biotech, Little Chalfont, Bucks, UK;
specific activity 74 GBq/mmol, 5 ¥ 103 Bq) and ABA
(Amersham; specific activity 1.74 TBq/mmol, 5 ¥ 103 Bq).
(anova) and Tukey’s post hoc test implemented in XLSTAT
(Addinsoft, Deutschland, Andernach, Germany) or t-test as
implemented in Microsoft Excel.
RESULTS
Effects of water deficit on leaf formation and
leaf loss
In well-watered plants, leaf number increased throughout
the experiment, from ca. 8 to 12–13 leaves per plant by
day 12. The rate of leaf formation did not differ significantly
among the genotypes tested (Fig. 1). Accelerated senescence of the lowest leaves induced by the water stress treatment resulted in a decrease in viable leaf number in all
genotypes. This started around day 7, and by the end of the
stress period (11 d from start) the water-stressed plants
had lost, on average, around three true leaves (Fig. 1).
In transformed plants constitutively expressing ZOG1
(35S::ZOG1), leaf loss started about 1 d earlier than in WT
and SAG::ZOG1 transformants (Fig. 1). Within 24 h of
rehydration, leaf formation resumed and the number of
leaves per plant in WT and SAG::ZOG1-transformed plants
began to increase (Fig. 1); the most rapid recovery occurring in the SAG::ZOG1 plants. Recovery of leaf formation
in water-stressed 35S::ZOG1 plants was barely detectable
after 24 h of rehydration.
In the stress treatments, the RWC of leaves decreased
with the duration of the stress conditions (Fig. 2), with
RWC falling by ca. 20% at day 6 (moderate stress) and by
ca. 40% at day 11 (severe stress). One day after rehydration, RWC increased in all water-stressed plants, almost to
the level observed in the control plants (Fig. 2).
HPLC of auxin and ABA
IAA and ABA were determined using two-dimensional
HPLC according to Dobrev et al. (2005). IAA was quantified using a fluorescence detector LC 240 (Perkin Elmer,
Wellesley, MA, USA). Quantification of ABA was performed on the basis of ultraviolet (UV) detection using a
diode array detector 235C (Perkin Elmer). The method was
verified by comparison with GC-MS.
HPLC/MS
LC-MS analysis was performed as described by Dobrev
et al. (2002) using an Ion Trap Mass Spectrometer Finnigan
MAT LCQ-MSn (Thermo Fisher, Waltham, MA, USA)
equipped with an electrospray interface. Detection and
quantification were carried out using a Finnigan LCQ operated in the positive ion, full-scan MS/MS mode using a
multi-level calibration graph with deuterated CKs as internal standards.The levels of 27 different CK derivatives were
measured. The detection limit was calculated for each compound as 3.3 s/S, where s is the standard deviation of the
response and S is the slope of the calibration curve. Three
independent experiments were performed. Each sample
was injected at least twice.
Quantification of ZOG1 expression in
transgenic plants
In order to determine the extent of ZOG1 expression in
transgenic plants, the amount of ZOG1 mRNA was quantified by real-time RT-PCR in the individual leaves and
roots of the control (well watered) and drought-treated
plants. Normalization of ZOG1 expression to actin (Act9)
and 18S rRNA led to similar results. No specific product was
amplified using ZOG1 primers from WT cDNAs.
The highest expression levels of ZOG1 were found
in SAG::ZOG1 transformants in lower (older) leaves
(Table 1). ZOG1 expression increased dramatically in
lower leaves following the onset of water deficit, and it
continued to increase throughout the stress period. SAG
promoter activity sharply decreased after rehydration
(Table 1). The impact of moderate stress (6 d) on the
pattern of SAG::ZOG1 expression is shown in more detail
in Fig. 3. Considerable stimulation of SAG promoter activity started in the fourth leaf (numbered from the base) of
the drought-stressed plants and in the fifth leaf of the
control plants. Physiologically, these two leaf positions
approximately corresponded to each other, as the stressed
Determination of CKX activity
The enzyme preparations were extracted and partially purified using the method described by Motyka et al. (2003).
The CKX activity was determined by in vitro assays based
on the conversion of [2-3H]isopentenyladenine (prepared
by Dr Jan Hanuš, Institute of Experimental Botany AS CR,
Prague, Czech Republic) to [3H]adenine. Separation of the
substrate from the product was achieved by HPLC. The
CKX activity was determined in two independent experiments. As the data from both experiments showed the same
tendencies (but different absolute values), the results of one
representative experiment (mean value of three parallel
assays for each of three replicate protein preparations) are
presented.
Statistical analysis
The significance of the differences among the measured
values was assessed by a four-factor analysis of variance
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 341–353
80
Role of cytokinins in responses of tobacco to water deficit 345
RWC (%)
Moderate
100
80
80
60
60
40
40
20
20
0
0
100
Severe
Recovery
100
80
80
60
60
40
40
20
20
RWC (%)
Middle leaves – control
Upper leaves – drought
Middle leaves – drought
Mild
100
Figure 2. Relative water content
(RWC) during the drought stress
progression in wild-type (WT),
SAG12::ZOG1 (SAG) and 35S::ZOG1
(35S) plants. Mild stress: 1 d dehydration;
moderate stress: 6 d dehydration; severe
stress: 11 d dehydration; recovery:
subsequent 1 d rehydration. C,
well-watered plants; D, drought-stressed
plants.
0
0
C D
WT
C D
SAG
C D
35S
C D
WT
C D
SAG
C D
35S
Stress
Table 1. SAG::ZOG1 expression after
mild, moderate and severe drought stress
and subsequent rehydration quantified by
real-time RT-PCR
Recovery
Mild (1 d)
Moderate (6 d)
ZOG1 expression (relative units)
Severe (11 d)
1d
Leaves
Upper
Middle
Lower
0.0006
0.0261
0.9792
0.0025
1.2087
2.3478
0.0002
0.0089
0.0041
0.0036
0.0283
1.7981
ZOG1 expression levels were normalized to actin (Act9). Results from two distinct experiments on the basis of triplicate determination were combined.
ZOG1, trans-zeatin O-glucosyltransferase gene.
ZOG1 expression (relative units)
plants had three leaves less than the control ones at the
time of measurement because of drought-induced growth
cessation.
Different levels of ZOG1 expression driven by the constitutive 35S promoter were detected under drought and
control conditions (Fig. 3). Analysis of individual leaves of
two plants grown under moderate and severe stress and two
control plants revealed that ZOG1 transcript level was significantly higher (more than double) in drought-stressed
plants than in the watered ones (t-test, P = 0.0001). As no
correlation between 35S::ZOG1 expression and leaf position was observed, the mean values ⫾ SE for control and
stressed leaves are presented.
SAG12::ZOG1 – control
SAG12::ZOG1 – drought
35S::ZOG1 – control
35S::ZOG1 – drought
12
10
8
6
4
2
0
1
1
2
2
3
4
5
6
7
8
Leaf position (drought)
3
9
4 5 6 7 8 9 10 11 12
Leaf position (control)
CK content
Figure 3. Quantitation of SAG12::ZOG1 and 35S::ZOG1
CK content has been expressed on a DW basis in order to
eliminate the difference caused by different water contents
between control and stressed plants. CK levels in the upper,
middle and lower leaves and roots of WT, SAG::ZOG1 and
35S::ZOG1 plants under control and different drought conditions are shown in Fig. 4. Individual CK metabolites were
assigned to one of seven groups according to their structure
and function. These groups included bioactive CKs, CK
expression in plants under control and moderate drought stress
conditions (6 d dehydration) by real-time RT-PCR. ZOG1
expression levels were normalized to actin (Act9). As no
correlation between 35S::ZOG1 expression and leaf position was
observed, mean values ⫾ SE for the control and stressed leaves
are presented. Results from two distinct experiments on the basis
of triplicate determination were combined.
© 2008 The Authors
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81
346 M. Havlová et al.
cis-Z(R)
Active CK
N-glc
cis-N-glc
O-glc
cis-O-glc
Phosphates
Mild
1000
800
Moderate
cis-Z(R)
Active CK
N-glc
cis-N-glc
O-glc
cis-O-glc
Phosphates
1200
1000
800
600
600
400
400
200
200
0
1200
cis-Z(R)
Active CK
N-glc
cis-N-glc
O-glc
cis-O-glc
Phosphates
Severe
1000
800
Recovery
0
1200
cis-Z(R)
Active CK
N-glc
cis-N-glc
O-glc
cis-O-glc
Phosphates
1000
800
600
600
400
400
200
200
0
UML R UML R UML R UML R UML R UML R
C
D
WT
C
D
C
SAG
UML R UML R UML R UML R UML R UML R
D
C
35S
D
C
WT
D
SAG
C
Total CK content (pmol g–1 DW)
Total CK content (pmol g–1 DW)
1200
0
D
35S
Figure 4. Endogenous cytokinin (CK) levels in tobacco leaves and roots during the drought stress progression in wild-type (WT),
SAG12::ZOG1 (SAG) and 35S::ZOG1 (35S) plants. CK metabolites were summarized into seven groups: (1) cis-zeatin (riboside)
[cis-Z(R)]; (2) bioactive CKs (trans-zeatin, isopentenyladenine, dihydrozeatin and the corresponding ribosides); (3) N-glucosides (7N- and
9N-glucosides of trans-zeatin, isopentenyladenine and dihydrozeatin) (N-glc); (4) cis-N-glucosides (cis-zeatin 7N- and 9N-glucoside)
(cis-N-glc); (5) O-glucosides [trans-zeatin (riboside) O-glucoside and dihydrozeatin (riboside) O-glucoside] (O-glc); (6) cis-O-glucosides
[cis-zeatin (riboside) O-glucoside] (cis-O-glc); and (7) CK phosphates (trans- and cis-zeatin-, isopentenyladenine- and dihydrozeatin
riboside phosphates). In the grouped bars, the first bar corresponds to the upper leaves (U), the second one to the middle leaves (M), the
third one to the lower leaves (L) and the last one to the roots (R). SE was in the range 5–18%. Other details the same as for Fig. 2. DW,
dry weight.
deactivation forms (N-glucosides), CK storage forms
(O-glucosides) and CK phosphates (CK biosynthesis intermediates). As cis-zeatin and its riboside are recognized by
some CK receptors, but lack physiological activity in most
CK bioassays, cis-zeatin derivatives have been shown as
separate groups (with the exception of cis-zeatin phosphate,
which was included with other CK phosphates). As the
bioactive CKs represent only a small portion of the total
Actie CK content (pmol g–1 DW)
iP
iPR
DHZ
DHZR
Z
ZR
Moderate
iP
iPR
DHZ
DHZR
25
20
20
15
15
10
10
5
5
0
0
25 Severe
Z
ZR
iP
iPR
DHZ
DHZR
Recovery
Z
ZR
iP
iPR
DHZ
DHZR
25
20
20
15
15
10
10
5
5
0
UML R UML R UML R
C
D
WT
UML R UML R UML R
C
D
SAG
C
D
35S
UML R UML R
C
D
WT
UML R
C
UML R UML R
D
SAG
UML R
C
D
35S
0
Actie CK content (pmol g–1 DW)
Z
ZR
25 Mild
CK pool, their contents are shown in detail in Fig. 5. Similarly, the dynamics of cis-zeatin/riboside contents is given in
Fig. 6.When expressed on a DW basis, the total CK contents
in roots are relatively lower than those in leaves, but per
gram FW, they are similar to, or higher than, those in leaves.
After mild stress, a significant elevation of bioactive CKs
was observed in WT upper leaves (t-test, P = 0.039; Fig. 5).
This increase was mainly a result of the elevation of
Figure 5. Endogenous bioactive
cytokinin (CK) levels (trans-zeatin,
isopentenyladenine, dihydrozeatin and
the corresponding ribosides) in tobacco
leaves and roots during the drought
stress progression in wild-type (WT),
SAG12::ZOG1 (SAG) and 35S::ZOG1
(35S) plants. Mean values ⫾ SE are
given. Other details the same as for
Figs 2 and 4.
© 2008 The Authors
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82
Role of cytokinins in responses of tobacco to water deficit 347
Upper leaves
Middle leaves
Lower leaves
Roots
Mild
Moderate
25
20
20
15
10
10
5
0
0
30
Severe
Recovery
30
20
20
10
10
0
C
D
WT
C
D
SAG
C
D
35S
C
D
WT
C
D
SAG
trans-zeatin/riboside and, to a low extent, also of
isopentenyladenine/riboside. The most remarkable increase
was monitored in the case of trans-zeatin. No significant
changes in bioactive CK levels in upper leaves were
observed between stressed and control SAG::ZOG1 or
35S::ZOG1 plants. The levels of cis-zeatin/riboside were not
significantly affected in either genotype. Mild stress did not
impose any remarkable effect on the size of the total CK
pool in either of the tested genotypes. No significant
difference was found in the total CKs between WT and
SAG::ZOG1 plants. Both control and stressed 35S::ZOG1
plants had considerably higher total CK levels than WT
(Tukey’s test, P < 0.0001), mainly as a result of higher levels
of O-glucosides, both of the trans-zeatin and the ciszeatin type. In roots, trans-zeatin O-glucosides strongly
predominated.
Moderate drought stress led to a decreased level of bioactive CKs in the leaves of all stressed plants, the highest
effect being imposed on trans-zeatin and its riboside. Nevertheless, a gradient in bioactive CKs in favour of the upper
leaves was observed in all tested genotypes, together with a
gradient in CK phosphates. The relatively small difference
in bioactive CK levels in the middle and lower leaves of
stressed SAG::ZOG1 plants might be the consequence of
enhanced SAG promoter activity in stressed lower leaves
(see Fig. 3). A much higher impact of stimulated SAG promoter activity was, however, observed at the distribution of
total CKs in these plants. Besides an increase in total CK
level in the upper leaves in response to water deficit, a
comparable elevation was found in lower leaves.
The increase of total CKs in leaves, caused by both water
stress and natural senescence, was found predominantly in
ZOG1 transgenics. It was mainly the result of raised levels
of the inactive CK forms – CK O- and N-glucosides, especially of the cis-zeatin type. Elevation of free cis-zeatin/
riboside was observed as well. In roots, the levels of CK
N- and O-glucosides were generally much lower. However,
in the roots of stressed plants, a mild elevation of the bioactive CK content was found.
C
D
cis-Z(R) content (pmol g–1 DW)
cis-Z(R) content (pmol g–1 DW)
30
0
35S
Figure 6. Levels of cis-zeatin and
cis-zeatin riboside [cis-Z(R)] in tobacco
leaves and roots during the drought
stress progression in wild-type (WT),
SAG12::ZOG1 (SAG) and 35S::ZOG1
(35S) plants. Mean values ⫾ SE are
given. Other details the same as for
Fig. 2.
Severely stressed plants were completely wilty. The bioactive CK content decreased in all genotypes, compared
with that in moderate stress. Differences between the
upper, middle and lower leaves were diminished. The flattening of the CK gradient might indicate that the strength
of the stress nearly reached the limits of the plant’s ability
to cope with drought. In SAG::ZOG1 plants, SAG promoter
activity increased in stressed middle leaves (in comparison
with moderate stress), leading to an increase in the total CK
content of these leaves comparable with that in the upper
ones. In lower leaves, strong SAG expression caused a
further elevation of the total CK content (Tukey’s test,
P < 0.0001). Besides elevated N-glucoside levels, large
increases in cis-zeatin O-glucosides and, to a lesser extent,
in trans-zeatin O-glucosides were also detected. In control
SAG::ZOG1 plants, the activity of the SAG promoter,
driven by natural senescence, resulted in the formation of a
gradient of total CKs in favour of the lower leaves. Because
of the SAG promoter activity, the levels of bioactive CKs
in the lower leaves of control plants were maintained approximately at the levels of the upper ones. In stressed
35S::ZOG1 plants, the levels of bioactive CKs in the upper
leaves were higher than those in the middle and lower ones.
The large increase in total CKs, especially in the lower
leaves, was caused by the increases in both O- and
N-glucosides (Tukey’s test, P < 0.0001).
In all the genotypes tested, severe stress led to the further
accumulation of total CKs in the roots. An increase in bioactive CKs was observed especially in the roots of WT and
SAG::ZOG1 plants.
Following watering, a decrease in the total CK content
was observed in the roots in all tested genotypes. Simultaneously, the level of bioactive CKs in the leaves increased
significantly in comparison with severely stressed plants
(Tukey’s test, P = 0.007). Considerable elevation was found
predominantly in the case of trans-zeatin/riboside. WT
plants had an elevated bioactive CK content in the upper
and, to a lower extent, also in the middle leaves. New
leaf formation commenced within 1 d of renewed water
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 341–353
83
348 M. Havlová et al.
CKX activity (pmol Ade·mg–1 protein·h–1)
1000
1200
Moderate
1000
800
800
600
600
400
400
200
200
0
1200 Severe
0
1200
Recovery
1000
1000
800
800
600
600
400
400
200
200
0
C
D
WT
C
D
SAG
C
D
35S
C
D
WT
C
D
SAG
application. SAG::ZOG1 plants exhibited an increase in
bioactive CK content in all leaves, and new leaf formation
commenced even faster than in WT plants. The level of
inactive CKs in SAG::ZOG1 plants fell, which correlated
well with the decrease of SAG promoter activity following
rehydration (see Table 1); nevertheless, two CK peaks (in
upper and lower leaves) were still apparent. 35S::ZOG1
transgenics had relatively high levels of bioactive CKs in all
leaves, in which CK O-glucosides had accumulated during
stress progression. Considerable decrease of free cis-zeatin/
riboside content was observed after rehydration, especially
in ZOG1 transgenics.
C
D
35S
0
CKX activity (pmol Ade·mg–1 protein·h–1)
Upper leaves
Middle leaves
Lower leaves
Roots
1200 Mild
Figure 7. The activity of cytokinin
oxidase/dehydrogenase (CKX) in
tobacco leaves and roots during the
drought stress progression in wild-type
(WT), SAG12::ZOG1 (SAG) and
35S::ZOG1 (35S) plants. Other details
the same as for Fig. 2.
control plants. After rehydration, CKX activity rose almost
to the level observed in the roots of control plants.
ABA
The ABA content of the plants was clearly correlated with
the degree of applied water stress (Fig. 8). Under mild stress
conditions, a slight increase in ABA concentration occurred
in the upper and middle leaves of WT plants and in the
middle leaves of SAG::ZOG1 plants. ABA levels did not
increase in 35S::ZOG1 plants subjected to mild water stress.
Under moderate stress, ABA content had risen substantially in all stressed plants, with SAG::ZOG1 plants exhibiting the greatest increase. Severe stress led to a further
substantial increase in ABA content (by more than one
order of magnitude in comparison with control conditions).
The highest concentration of ABA occurred in the upper
leaves. Following rehydration, ABA levels began to
decrease in all treatments, although after a 1 d recovery
period, they remained significantly higher than those
observed in control plants. No significant differences in
ABA levels between the genotypes were observed during
recovery.
CKX activity
In young, well-watered plants, constitutive ZOG1 expression led to a considerable decrease in CKX activity in leaves
in comparison with WT plants (Fig. 7). After moderate
stress, an increase in CKX activity in the middle leaves in
SAG::ZOG1 plants was observed. In constitutive transformants, a decrease in CKX activity in stressed upper leaves
was found. After severe stress, a strong enhancement of
CKX activity was observed in the lower leaves of all plants,
together with the decrease in CKX activity in the upper
leaves, which might contribute to the maintenance of a
bioactive CK gradient. After rehydration, a rise in CKX
activity in the leaves was observed, presumably to regulate
bioactive CK levels after the re-establishment of the transpiration flow.
CKX activity was much higher in the roots than in the
leaves in all genotypes (Tukey’s test, P < 0.0001). After mild
stress, CKX activity in the roots decreased in WT and
35S::ZOG1 plants. After moderate stress, CKX activity fell
in the roots of all the tested genotypes.The decreasing trend
of CKX activity in the roots was reversed after severe stress,
but still, the CKX activity was well below the level in
Auxin
After 1 d of dehydration, an increased level of free IAA was
found in WT lower leaves (Fig. 9). Mild elevation was
observed in the roots of both transgenics. In all genotypes,
moderate stress resulted in a substantial increase in IAA
concentration, both in the lower leaves and roots (Tukey’s
test, P = 0.024).After severe stress, the highest IAA concentration was found in the lower leaves of all tested plants
(highly significant increase in comparison with the control,
Tukey’s test, P < 0.0003). Differences in IAA level between
the stressed lower and upper leaves were highly significant
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 341–353
84
Role of cytokinins in responses of tobacco to water deficit 349
ABA content (pmol g–1 DW)
Moderate
60000
10000
40000
5000
0
80000
20000
Severe
0
15000
Recovery
60000
10000
ABA content (pmol g–1 DW)
Upper leaves
Middle leaves
Lower leaves
Roots
15000 Mild
Figure 8. Endogenous level of abscisic
acid (ABA) in tobacco leaves and roots
during the drought stress progression in
wild-type (WT), SAG12::ZOG1 (SAG)
and 35S::ZOG1 (35S) plants. Mean
values ⫾ SE are given. Other details the
same as for Fig. 2.
40000
5000
20000
C
D
C
WT
D
C
SAG
D
C
35S
D
WT
C
D
SAG
Upper leaves
Middle leaves
Lower leaves
Roots
IAA content (pmol g–1 DW)
4000
D
0
35S
(Tukey’s test, P < 0.0002), between the stressed lower and
middle leaves were significant (Tukey’s test, P = 0.002),
and between the lower leaves and roots were significant
(Tukey’s test, P = 0.008). In all plants, rehydration led to a
fall in IAA levels in the lower leaves and roots, and in WT,
almost to the control values. In SAG::ZOG1 transgenics, a
significantly elevated IAA content was observed in all rehydrated leaves. Natural senescence led to a mild elevation of
IAA content in the lower leaves of WT and 35S::ZOG1
plants, with some delay also in SAG::ZOG1 plants.
Fast and strong drought treatment (at 35% humidity) led,
apart from the elevation of IAA in the roots, to the slight
increase of WT root biomass, mainly caused by root elongation and thickening (not shown). Because of the low SAG
promoter activity in the roots, only a negligible negative
effect on root growth was observed in SAG::ZOG1 plants.
The 35S::ZOG1 plants, which already had a longer root
5000 Mild
C
system under control conditions, exhibited a reduced ability
to increase root biomass during the drought; thus, its rootto-shoot ratio was the lowest among the plant types tested
(results not shown).
DISCUSSION
Changes in the distribution of plant hormones
in leaves in response to water deficit
In plants adequately supplied with water, the concentration
of bioactive CKs is normally greatest in the young leaves
and declines as the leaves age and start to senesce (e.g.
Singh, Letham & Palni 1992). Accordingly, we observed at
the beginning of the experiment, when the control plants
were juvenile, with no visible signs of senescence, a similar
bioactive CK content in all leaves. During the experiment,
5000
Moderate
4000
3000
3000
2000
2000
1000
1000
0
5000 Severe
0
5000
Recovery
4000
4000
3000
3000
2000
2000
1000
1000
0
C
D
WT
C
D
SAG
C
D
35S
C
D
WT
C
D
SAG
C
D
IAA content (pmol g–1 DW)
0
0
35S
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 341–353
85
Figure 9. Endogenous level of
indolyl-3-acetic acid (IAA) in tobacco
leaves and roots during the drought
stress progression in wild-type (WT),
SAG12::ZOG1 (SAG) and 35S::ZOG1
(35S) plants. Mean values ⫾ SE are
given. Other details the same as for
Fig. 2.
350 M. Havlová et al.
simultaneously with the development of the symptoms of
senescence (gradual decrease in chlorophyll content of
lower leaves), a gradient of bioactive CKs in favour of the
upper leaves was observed (Fig. 5).
In order to elucidate, at least partially, CK function
during the drought stress, we compared the water deficit
response of plants with a uniform elevation of CK content
before stress initiation and those exhibiting CK elevation
localized to senescing tissues. As constitutive overexpression of CK biosynthetic gene results in the severe
modification of plant morphology, we decided to manipulate the CK levels in more gentle way using plants overexpressing ZOG1. Constitutive over-expression of ZOG1
had only a minor effect on plant morphology, as well as on
the levels of bioactive CKs (see also Martin et al. 2001).
However, because of the permanently enhanced CK
O-glucosylation, which stimulated the CK turnover,
35S::ZOG1 plants exhibited under control conditions
symptoms of CK elevation, i.e. higher transpiration rate and
higher net photosynthetic rate (Pospíšilová, personal
communication).
For the targeted elevation of CK levels, we used the
SAG12 promoter. Since the pioneering work of Gan &
Amasino (1995) on SAG12::ipt tobacco, this promoter has
repeatedly been used for the targeted manipulation of CK
content. In accordance with Cowan et al. (2005), who found
an increase in trans-zeatin-type CKs in senescing wellwatered SAG12::ipt plants, we found an elevation of CK
content in senescing control SAG12::ZOG1 plants. Apart
from the expected increase of the level of CK O-glucosides,
we also found an elevation of bioactive CKs when SAG
promoter activity was stimulated.
In order to check the ability of ZOG1 transformants to
utilize the elevated CK O-glucoside pool, b-glucosidase
activity was compared in WT and ZOG1 expressing plants.
Mild elevation of b-glucosidase activity in ZOG1 transformants (results not shown) seemed to prove their capacity to
hydrolyse the CK storage pool. Another question arose,
whether the stress response of ZOG1 transformants could
not be modulated by a massive hydrolysis of the CK
O-glucoside pool under the stress conditions. The dynamics
of b-glucosidase activity during the stress progression
(results not shown), as well as the CK analyses, indicated
that CK O-glucoside hydrolysis has been under strict environmental control.
In WT plants, the formation of a gradient of bioactive
CKs in favour of the upper leaves was observed when mild
stress was imposed. The occurrence of this gradient might
reflect the necessity for the preferential protection of upper
leaves under unfavourable environmental conditions. The
importance of CKs in sink–source polarization during mild
water stress was recently discussed by Cowan et al. (2005).
Constitutive ZOG1 transformants did not exhibit any significant response to mild stress. No increase in ABA content
(observed in the WT upper and middle leaves) was
detected. Neither the bioactive CK nor the IAA levels were
considerably changed. The lack of difference in CK levels
between the upper and middle leaves of 35S::ZOG1 plants
coincided with the delayed decrease of stomatal aperture
and net photosynthetic rate of the middle leaves in
response to water deficit (Pospíšilová, personal communication), which is in accordance with the positive effect of
CKs on stomatal aperture (e.g. Rulcová & Pospíšilová
2001). Elevated CK content before stress initiation, thus,
might postpone the reaction of the plants to adverse environmental conditions.
The moderate water deficit (RWC decreased by ca. 20%)
had two main effects on the concentrations of bioactive
CKs. Firstly, in agreement with Goicoechea et al. (1995), it
caused a progressive fall in the concentration of bioactive
CKs (especially of trans-zeatin/riboside) in all the leaves of
all the genotypes tested. Secondly, the steepness of the
apex-to-base gradient in bioactive CKs increased substantially, which was remarkable, especially in 35S::ZOG1
plants. In SAG::ZOG1 plants, moderate stress caused the
high stimulation of ZOG1 expression in the lower leaves
(Table 1), which resulted in an increase in the total CK
levels in these leaves. However, the bioactive CK content in
the lower leaves did not differ significantly from that in the
middle ones, which indicates that bioactive CK levels are
strictly controlled under stress conditions.
ABA levels strongly correlated with the degree of stress.
This finding agrees with previous reports (e.g. Mansfield &
McAinsh 1995; Riccardi et al. 2004). As ABA is an important component of the plant stress defence system, its
higher levels, observed in the upper leaves, might reflect
their preferential protection. Interestingly, as senescence
occurred in lower leaves, these results also indicate that
while ABA levels may increase during stress-induced leaf
senescence, ABA itself is not the cause of senescence.
Under severe stress (RWC decreased by ca. 40%), all the
leaves of the tested genotypes were wilty. The bioactive CK
gradient was suppressed in the WT and SAG::ZOG1 plants,
while it was more profound in the 35S::ZOG1 ones. Nevertheless, the decrease of CKX activity in the upper leaves
and, especially, the large increase in CKX activity in the
lower leaves seems to indicate a tendency for the gradient
of bioactive CKs to be maintained.
Large elevation of IAA content, observed in the stressed
lower leaves (at mild stress only in WT, at moderate stress
in WT and SAG::ZOG1 plants, and at severe stress in all
genotypes) and also in the severely stressed middle leaves
of 35S::ZOG1 plants, is in agreement with the results of
Quirino, Normanly & Amasino (1999), who found an
increase in the free auxin levels in senescing Arabidopsis
leaves. This increase correlated with the stimulation of the
expression of nitrilase 2, an enzyme catalysing the conversion of indole-3-acetonitrile to IAA. Both senescence and
pathogen attack have been found to induce nitrilases 2
and 4 in leaves (Bartel & Fink 1994). The function of the
increased IAA content of senescing leaves is not clear. As
senescence is a strongly regulated process, high IAA levels
might be necessary for its control. The flow of auxin to the
abscission zone is necessary to prevent leaf abscission. A
possible explanation might be that the increased protein
breakdown in senescing leaves leads to an increase in the
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 341–353
86
Role of cytokinins in responses of tobacco to water deficit 351
content of tryptophan, which may be converted to IAA, in
order to facilitate the transport from senescing leaves.
Rehydration resulted in an increase in bioactive CK
levels (especially of trans-zeatin/riboside) in the leaves of
all genotypes, which might be, at least partially, result from
the re-establishment of xylem sap flow from the roots. The
crucial impact of the transpiration stream on CK transport
from the roots was recently nicely demonstrated (Aloni
et al. 2006). Mild elevation of CKX activity in leaves might
reflect the necessity to control the sudden excess of CKs.
The stress indicators (SAG promoter activity and ABA
level) exhibited a highly significant decrease. Elevation of
IAA levels in the middle leaves (and, in SAG::ZOG1 plants,
also in the upper leaves) seems to be related to growth
re-establishment. The slightly faster growth initiation in
SAG::ZOG1 plants in comparison with WT is in accordance
with the known positive effect of CKs on plant recovery
(e.g. Itai et al. 1978). The slower recovery of 35S::ZOG1
transgenics indicates that elevated CK content need not be
advantageous in case of prolonged, severe drought stress.
This conclusion agrees with data on pssu:ipt over-expressing
tobacco plants, which had a highly elevated CK content and
which were found to be more vulnerable to water deficit
(Synková et al. 1999).
roots, in contrast to the situation in senescing leaves
(Quirino et al. 1999), auxin increase in the stressed roots
might be a consequence of auxin polar transport. Increased
IAA levels were also observed under desiccation stress in
maize roots (Ribaut & Pilet 1994; Xin, Zhou & Pilet 1997).
The function of auxin during the drought-stress response of
roots remains to be elucidated. Studies on axr1-3 mutants of
Arabidopsis revealed that functional auxin signalling has
been necessary for the induction of stress-related proteins
RD29B and glutamine synthetase in roots, as well as for
drought rhizogenesis (Bianchi, Danerval & Vartanian
2002). Increased levels of auxin might cause an additional
elevation of CK content, as auxin was found to promote
the expression of genes for CK biosynthetic enzymes in
roots (namely, AtIPT5 and AtIPT7; Miyawaki, MatsumotoKitano & Kakimoto 2004). An elevated content of both
hormones might stimulate root cell division, which resulted,
because of the inhibitory effect of CKs on lateral root
formation, in primary root elongation.
In conclusion, data on SAG12 promoter activity demonstrated that drought strongly accelerated the senescence of
tobacco plants, which coincided with the gradient of bioactive CKs in favour of the upper leaves. During the drought
stress progression, the level of bioactive CKs decreased
in growth-suppressed leaves. Nevertheless, the gradient of
bioactive CKs was maintained (even if flattened) by the
stimulation of the activity of CKX, the main CK-degrading
enzyme, in the lower leaves and by its suppression in the
upper leaves. Accumulation of CKs and free auxin in the
stressed roots suggests their importance for a droughtinduced change of root morphology (enhanced root elongation). Constitutive 35S::ZOG1 transformants exhibited a
delay in the stress-induced elevation of ABA. Thus, the
uniform elevation of CK content before stress initiation
might have a negative effect during the prolonged, severe
drought. Targeted, senescence-induced elevation of ZOG1
expression exhibited a positive effect both on the delay of
senescence of the stressed lower leaves and during the
recovery.
Root growth modification
Our observation of the effect of constitutive ZOG1 expression on the morphology of the root system (primary root
elongation and diminished branching) is in agreement with
the concept of root apical dominance described by Aloni
et al. (2006). It also agrees with data on the impact of
decreased CK content (in plants over-expressing genes for
CKX; Werner et al. 2003) or after the disturbance of the CK
signal transduction pathway via mutation of individual CK
receptors (Riefler et al. 2006), when the stimulation of the
lateral and adventitious root formation was found.
However, a functioning CK signal transduction pathway
seems to be necessary for root growth, as mutations of all
three CK receptors were reported to have strong inhibitory
effects (Nishimura et al. 2004).
During the stress progression, we observed the accumulation of CKs in the roots of all tested genotypes. As CKs
are supposed to be predominantly root-borne phytohormones, which are distributed in the shoot via the xylem
stream (Beck & Wagner 1994), CK accumulation in roots
might be caused passively by a decrease in the velocity of
the transpiration stream. However, the diminished loading
of CKs into the xylem observed during the drought (Yang
et al. 2002), as well as the large decrease of CKX activity in
the stressed roots, suggest the physiological relevance of
drought-induced CK accumulation in roots. Limited transport of CKs into shoots seemed to coincide with their
growth suppression, while CK accumulation in roots might
be of importance for the increase of the root-to-shoot ratio.
During the prolonged drought stress, the elevation of
IAA concentrations was observed in the lower leaves and
roots. As no expression of nitrilase was detected in the
ACKNOWLEDGMENTS
We are very grateful to Professor Richard Amasino (University of Wisconsin, Madison, WI, USA) for his donation
of the SAG12 promoter and Professors Machteld and
David Mok and Dr Ruth Martin (Oregon State University, Corvallis, OR, USA) for supplementation of the seeds
of the 35S::ZOG1- and SAG12::ZOG1-transformed
tobacco. We express our thanks to Professor David Morris
(Palacky University, Olomouc, Czech Republic) and Dr
Miroslav Kaminek (Institute of Experimental Botany,
Prague, Czech Republic) for a critical reading of this
manuscript. For technical assistance, we thank Eva
Kobzova and Marie Korecka. This work was supported by
the Grant Agency of the Czech Republic, Project No. 522/
04/0549, and by the Ministry of Education, Youth and
Sports CR, Project Nos. LC06034 and ME868.
© 2008 The Authors
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Received 28 September 2007; received in revised form 20 November
2007; accepted for publication 30 November 2007
SUPPLEMENTARY MATERIAL
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Table S1. Primers for Nicotiana tabacum genes used in
qRT-PCR.
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