Journal of Plant Physiology 167 (2010) 1457–1465
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Journal of Plant Physiology
journal homepage: www.elsevier.de/jplph
Characterization of photosystem II photochemistry in transgenic tobacco plants
with lowered Rubisco activase content
Bin Cai, Aihong Zhang, Zhipan Yang, Qingtao Lu, Xiaogang Wen, Congming Lu ∗
Photosynthesis Research Center, Key Laboratory of Photobiology, Institute of Botany, Chinese Academy of Sciences, Beijing 100093, PR China
a r t i c l e
i n f o
Article history:
Received 15 March 2010
Received in revised form 18 May 2010
Accepted 19 May 2010
Keywords:
CO2 assimilation
Photosystem II
Rubisco activase
Thermoluminescence
Tobacco (Nicotiana tabacum)
a b s t r a c t
Rubisco activase plays an important role in the regulation of CO2 assimilation. However, it is unknown
how activase regulates photosystem II (PSII) photochemistry. To investigate the effects of Rubisco activase on PSII photochemistry, we obtained transgenic tobacco (Nicotiana tabacum) plants with 50% (i7),
25% (i28), and 5% (i46) activase levels as compared to wild type plants by using a gene encoding tobacco
activase for the RNAi construct. Both CO2 assimilation and PSII activity were significantly reduced only in
transgenic i28 and i46 plants, suggesting that activase deficiency led to decreased PSII activity. Flashinduced fluorescence kinetics indicated that activase deficiency resulted in a slow electron transfer
between QA (primary quinine electron acceptor of PSII) and QB (secondary quinone electron acceptor of PSII). Thermoluminescence measurements revealed that activase deficiency induced a shift of
S2 QA − and S2 QB − recombinations to higher temperatures in parallel, and a decrease in the intensities
of the thermoluminescence emissions. Activase deficiency also dampened the period-four oscillation of
the thermoluminescence B-band. Protein gel blot analysis showed that activase deficiency resulted in
a significant decrease in the content of D1, D2, CP43, CP47, and PsbO proteins. Transmission electron
microscopy analysis demonstrated that activase deficiency induced a significant decrease in the number of grana stacks per chloroplast and discs per grana stack. Our results suggest that activase plays an
important role in maintaining PSII function and chloroplast development.
© 2010 Elsevier GmbH. All rights reserved.
Introduction
Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco, EC
4.1.1.39) is a key regulatory enzyme of photosynthetic carbon
assimilation, which catalyzes the first step of both the photosynthetic carbon assimilation and photorespiratory pathways
(Woodrow and Berry, 1988). Prior to acting as a catalyst, Rubisco
must be activated to become catalytically competent in the photosynthetic carbon reduction cycle by the binding of an ‘activator’
CO2 and Mg2+ to a lysine residue within Rubisco’s catalytic sites
(Miziorko and Lorimer, 1983). The Rubisco activation process is catalyzed by the chloroplast enzyme Rubisco activase (Portis, 1990).
Rubisco activase does not appear to affect Rubisco activation per
se, but it allows uninhibited access to the active site for binding of
Abbreviations: DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea; Fv /Fm , the
maximal efficiency of PSII photochemistry in the dark-adapted state; PSI, photosystem I; PSII, photosystem II; QA , primary quinine electron acceptor of PSII; QB ,
secondary quinone electron acceptor of PSII; Rubisco, ribulose-1,5-bisphosphate
carboxylase/oxygenase.
∗ Corresponding author. Tel.: +86 10 6283 6554; fax: +86 10 6259 5516.
E-mail address: [email protected] (C. Lu).
0176-1617/$ – see front matter © 2010 Elsevier GmbH. All rights reserved.
doi:10.1016/j.jplph.2010.05.004
the activators CO2 and Mg2+ (Portis, 1990, 1992). Rubisco activase
requires ATP for activity, and is inhibited by ADP (Streusand and
Portis, 1987; Robinson and Portis, 1989). It is thought that Rubisco
activase controls the release of ribulose-1,5-bisphosphate (RuBP)
and other inhibitors from Rubisco, and thus regulates its activation
state (Brooks and Portis, 1988; Portis, 1990). To date, the precise
mechanism by which activase acts remains unclear, but most likely
involves an interaction between activase and Rubisco–sugar phosphate complexes (Portis, 2003; Portis et al., 2008).
The essential role of activase in regulating photosynthesis is
illustrated by the rca mutant of Arabidopsis that lacks this protein
and is unable to maintain sufficient activation of Rubisco to grow
under ambient CO2 concentrations (Somerville et al., 1982; Salvucci
et al., 1985, 1986). To investigate the role of activase in regulating
photosynthesis, a series of studies have examined the effects of different activase contents on CO2 assimilation rate using transgenic
plants with reduced levels of activase compared to wild type plants
(Mate et al., 1993, 1996; Jiang et al., 1994; Eckardt et al., 1997; Jin
et al., 2006). Although the relatively large amount of protein per
leaf is allocated to activase (Fukayama et al., 1996), the level of
activase does not proportionally affect CO2 assimilation rate. Only
when activase contents were decreased to about 10–35% of that of
the wild type plants was there a significant effect on CO2 assimila-
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B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
tion rate (Mate et al., 1993; Jiang et al., 1994; Eckardt et al., 1997;
von Caemmerer et al., 2005; Jin et al., 2006).
Previous studies on the role of activase in photosynthesis have
been focused on the regulation of CO2 assimilation (Jiang et al.,
1994; Mate et al., 1996; Eckardt et al., 1997; von Caemmerer et
al., 2005; Jin et al., 2006). However, to fully examine the control
that activase exerts on photosynthesis, there is a need to investigate how activase affects photosystem II (PSII) photochemistry and
chloroplast development. In this study, we investigated the role of
activase in regulating PSII photochemistry and chloroplast development in transgenic tobacco plants with reduced activase levels.
Our results revealed that activase deficiency led to decreased PSII
activity and abnormal chloroplast development.
Materials and methods
Vectors and plant transformation
The partial coding region for the tobacco Rubisco activase gene
(NTU35111, gi: 1006834) was cloned into the pKANNIBAL vector between the XhoI–KpnI sites in sense orientation and the
BamHI–ClaI sites in antisense orientation (Wesley et al., 2001). The
primers used were: 5 -GAT CTC GAG TTA TCT TGG GTA TCT GGG-3
and 5 -CAC GGT ACC GGA CCC TGG CAT TCT CTT-3 (irca antisense
primers) and 5 -TTC GGA TCC TTA TCT TGG GTA TCT GGG-3 and
5 -GCG ATC GAT GGA CCC TGG CAT TCT CTT-3 (irca sense primers).
The construct made in pKANNIBAL was subcloned as NotI fragment
into pART27, then introduced into Agrobacterium tumefaciens strain
LBA4404 by tri-parental mating (An, 1987). Nicotiana tabaccum
(Wisconsin 38) were transformed by the standard Agrobacteriummediated transformation (Horsh et al., 1985). Regenerated plants
were transplanted into sterilized soil and grown in a greenhouse at
27/20 ◦ C (day/night), with maximum PPFD of 1000 ␮mol m−2 s−1 ,
and a photoperiod of 12/12 h light/dark. T1 plants obtained by selfpollination of T0 plants were used in this study.
Plant growth
Three independent lines of transgenic tobacco plants (i7, i28,
and i46) were used in the present study. The seeds of these transgenic plants were allowed to germinate on agar in the presence of
50 ␮g l−1 kanamycin. The seeds of wild type plants were allowed
to germinate an agar in the absence of kanamycin. After growth for
2 weeks, plants were transferred to soil. The transplanted plants
were then grown for 8 weeks in a growth chamber at 26 ± 1 ◦ C
with PPFD of 120 ␮mol m−2 s−1 , a relative humidity of 75–80%, and
a photoperiod of 12/12 h light/dark. Plants were watered daily and
twice weekly with a complete nutrient solution. All the measurements on physiological and biochemical parameters were carried
out on the youngest fully expanded leaves.
Transmission electron microscopy
For transmission electron microscopy processing, the tobacco
leaves from 8-week old plants were collected. The leaf tissue was
cut into small pieces and fixed in 1.25% (v/v) glutaraldehyde and
1.25% (v/v) paraformaldehyde in phosphate buffer for 4 h at 4 ◦ C.
After fixation, tissues were rinsed and post-fixed for 45 min in 1%
(v/v) OS O4 overnight at 4 ◦ C. After rinsing in phosphate buffer, the
samples were dehydrated in an ethanol series, infiltrated with a
graded series of epoxy resin in epoxy propane, and embedded in
Eponaraldite (Agar Aids, Bishop’s Stortford, UK). After 12 h in pure
resin, followed by a change of fresh resin for 4 h, the samples were
polymerized at 60 ◦ C for 48 h. Ultrathin sections (70 nm thick) were
obtained with a Leica ultramicrotome, stained with uranyl acetate,
pH 5.0, and lead citrate. Electron micrographs were viewed with
a transmission electron microscope JEM-1230 TEM (JEOL) at an
accelerating voltage of 80 kV.
Analysis of gas exchange
Gas exchange analysis was performed on a fully expanded
attached leaf of tobacco seedlings using an open system (Ciras-1,
PP systems, UK). Leaf CO2 assimilation rate was made at the atmospheric CO2 concentration of 360 ␮l l−1 and at a temperature 25 ◦ C
with a relative humidity 80% and growth light (120 ␮mol m−2 s−1 ).
Measurements of chlorophyll fluorescence
Chlorophyll fluorescence was measured using a PAM 2000
portable chlorophyll fluorometer connected by a leaf-clip holder
with a trifurcated fiber optic cable (Heinz Walz, Germany). The
measurements were performed on the attached leaves of tobacco
seedlings. Before each measurement, leaves were dark adapted for
20 min. After that, the Fo was measured by a red light (650 nm) with
very low intensity (0.8 ␮mol m−2 s−1 ). A saturating pulse of white
light (8000 ␮mol m−2 s−1 for 0.8 s) was applied to determine the
maximum fluorescence at closed photosystem II (PSII) centers in
the dark-adapted state (Fm ). The maximal efficiency of PSII photochemistry in the dark-adapted state, i.e. Fv /Fm , was calculated as
(Fm − Fo )/Fm .
The decay of chlorophyll a fluorescence yield after a singleturnover flash was measured with a double-modulation fluorescence fluorometer (model FL-200, Photon Systems Instruments,
Brno, Czech Republic) (Trtilek et al., 1997). The instrument contained red LEDs for both actinic (20 ␮s) and measuring (2.5 ␮s)
flashes, and was used in the time range of 100 ␮s to 100 s. With
this type of measurement, it is important to avoid distortion of the
relaxation kinetics due to the actinic effect of measuring flashes.
This was carefully checked, and the intensity of the measuring
flashes was set at a value that was low enough to avoid reduction
of QA (primary quinine electron acceptor of PSII) in the presence of
3-(3 ,4 -dichlorophenyl)-1,1-dimethylurea (DCMU).
Semi-quantitative RT-PCR analysis
Thermoluminescence(TL) measurements
Total RNA was extracted from 0.1 g fresh leaves using the Trizol reagent (Invitrogen Carlsbad, USA). After DNase I treatment to
remove any residual genomic DNA contamination, 2 ␮g of total
RNA from each sample was used to synthesize first-strand cDNA
in a 20 ␮l total volume (SuperScript Pre-amplification System,
Promega, USA). The transcript levels in 8-week old plants were
examined by semi-quantitative RT-PCR using irca sense primers,
and amplified PCR products were collected and analyzed following different numbers (20 cycles, 30 cycles) of amplification cycles.
The RT-PCR reactions were repeated twice, with identical results
obtained. The expression level of tobacco actin was used as an
internal control, as described previously (Ding et al., 2009).
TL measurements of leaves were performed with the thermoluminescence extension of the Double-Modulated Fluorometer
FL2000-S/F, consisting of Thermoregulator TR2000 (Photon Systems Instruments, Brno, Czech Republic). After 30 min dark
adaptation at 30 ◦ C, the cells were cooled to 0 ◦ C and illuminated
with one or multiple number of single-turnover flashes. Then the
cells were warmed up to 60 ◦ C at a heating rate of 1 ◦ C s−1 and the TL
light emission was measured during the heating. To detect periodfour oscillation of the B-band, leaves were illuminated with a series
of single-turnover flashes. For S2 QA − recombination studies, leaves
were measured in the presence of DCMU (20 ␮M) before the flash
B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
1459
illumination. The nomenclature of Vass and Govindjee (1996) was
used for characterization of the flash-induced TL glow peaks.
Electron transport activities
Photosynthetic electron transport activities were determined
as described previously (Yang et al., 2008). The youngest fully
expanded leaves were harvested and homogenized in a medium
containing 0.4 M sucrose, 50 mM tricine (pH 7.6). The homogenate
was filtrated through 16 layers of gauze and the filtrate was then
centrifuged at 500 × g for 3 min to remove large debris. The supernatant was further centrifuged at 3000 × g for 10 min. The pellet
was washed twice by the buffer (50 mM tricine, 10 mM NaCl, 5 mM
MgCl2 , pH 7.6) at 10,000 × g for 10 min. The resulting washed pellet
was thylakoid membranes and resuspended in the same buffer for
measuring of electron transport activities.
Electron transport activities in the thylakoid membranes were
measured with a Clark-type oxygen electrode (Hansatech, King’s
Lynn, Norfolk, UK) suspended in the medium (0.4 M sucrose, 50 mM
Tricine, 10 mM NaCl, 5 mM MgCl2 , pH 7.6) under growth light
intensity (120 ␮mol m−2 s−1 ). Photosystem I (PSI) electron transport activity was measured with 10 mM methylamine, 10 ␮M
DCMU, 1 mM sodium azide, 500 ␮M 2,6-dichlorophenol indophenol (DCPIP), 2 mM sodium ascorbate, and 1 mM methyl viologen.
PSII electron transport activity was determined using 1 mM phenylp-benzoquinone as electron acceptor.
BN-PAGE, SDS-PAGE, and immunological analyses
BN-PAGE was performed using acrylamide gels with linear
6–12% gradients, basically according to Peng et al. (2006). For
immunological analysis, proteins either from isolated thylakoid
membranes or from total leaf proteins were separated by SDS-PAGE
with 12% SDS polyacrylamide running gels containing 6 M urea
(Laemmli, 1970). After electrophoresis, proteins were transferred
electrophoretically to a polyvinylidence difluoride membrane
(Amersham Biosciences, USA). Antibodies raised against D1, D2,
CP43, CP47, PsbO, LHCII, Cyt f, PsaA and AtpA/B, Rubisco (ribulose1,5-bisphosphate carboxylase/oxygenase), and Rubisco activase
were used. Thylakoid membrane proteins were loaded with an
equal amount of chlorophyll (5 ␮g). For detection of the immobilized antibodies, a chemiluminescent substrate (Super-Signal West
Pico Chemiluminescent Substrate, USA) was used, and the bands
were exposed to X-ray film. After development, the X-ray films
were scanned with a UMax PowerLook III scanner at 300-dpi resolution and an 8-bit color depth.
Analyses of protein and chlorophyll
Chlorophyll content was determined in 80% (v/v) acetone
according to Porra et al. (1989). Soluble protein content was determined by the dye-binding assay according to Bradford (1976).
Statistical analysis
For the comparison of variance of the means, the data were
analyzed using analysis of variance (ANOVA) followed by the least
significant difference (LSD) test.
Fig. 1. (A) Protein gel blot analysis of the level of activase (RCA) and Rubisco large
subunit (RbcL) in wild type (WT) and transgenic i7, i28, and i46 tobacco plants. (B)
RT-PCR analysis in wild type (WT) and transgenic i7, i28, and i46 tobacco plants. The
tobacco actin was amplified as internal standard. (C) Phenotype of wild type (WT)
and transgenic i7, i28, and i46 tobacco plants.
transgenic lines were obtained, and some of them exhibited low
Rubisco activase levels. Three independent lines, i.e. i7, i28, and
i46 plants, were chosen to obtain progeny (T1 generation) by selffertilization. Protein gel blot analysis demonstrated that the level of
activase in transgenic i7, i28, and i46 plants was reduced to about
50%, 25% and 5% of that of wild type plants, respectively. However,
there was no significant change in the content of large subunit of
Rubisco in the transgenic tobacco plants (Fig. 1A). RT-PCR analysis showed that the mRNA levels of activase in transgenic i7, i28
and i46 plants were suppressed compared to those in wild type
plants (Fig. 1B). Compared to wild type plants, transgenic i7 plants
did not show significant morphological differences, but transgenic
i28 and i46 plants exhibited a considerable decrease in growth
(Fig. 1C). There were no differences in the contents of leaf chlorophyll between wild type and transgenic i7 plants. The contents of
leaf chlorophyll were 447.4 ± 23.1 and 447.2 ± 19.8 mg m−2 in wild
type and transgenic i7 plants, respectively. The leaves in transgenic
i28 and i46 plants appeared green-yellowish. The contents of leaf
chlorophyll in transgenic i28 and i46 plants were 391.9 ± 17.5 and
145.8 ± 13.2 mg m−2 , respectively.
CO2 assimilation
Table 1 shows CO2 assimilation rates in wild type and transgenic
plants. There was no significant change in CO2 assimilation rate
between wild type and transgenic i7 plants. The CO2 assimilation
rates were 8.7 and 8.5 ␮mol m−2 s−1 in wild type and transgenic i7
plants, respectively. However, compared to wild type plants, there
was a significant decrease in CO2 assimilation rate in transgenic i28
and i46 plants. The CO2 assimilation rates in transgenic i28 and i46
plants were 6.9 and −0.12 ␮mol m−2 s−1 , respectively.
Table 1
CO2 assimilation rate in wild type (WT) and transgenic i7, i28, and i46
tobacco plants.
Results
Generation of transgenic tobacco plants with decreased Rubisco
activase level
In this study, we generated transgenic tobacco plants with the
RNAi construct of tobacco Rubisco activase. Sixty independent
CO2 assimilation rate (␮mol m−2 s−1 )
WT
i7
i28
i46
8.71
8.52
6.90
−0.12
±
±
±
±
0.23a
0.23a
0.23b
0.20c
Values in the table are means ± SE from three independent experiments.
Values indicated with different letters were significantly different at P = 0.05.
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B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
Table 2
PSI and PSII electron transport activities in wild type (WT) and transgenic i7, i28,
and i46 tobacco plants.
PSI activity
(␮mol O2 mg−1 Chl h−1 )
WT
i7
i28
i46
236
240
243
246
±
±
±
±
4a
2a
4a
2a
PSII activity
(␮mol O2 mg−1 Chl h−1 )
122
125
95
76
±
±
±
±
2a
4a
5b
4c
Values in the table are means ± SE from three independent experiments. Values
indicated with different letters were significantly different at P = 0.05.
PSI and PSII electron transport activities
Table 2 shows PSI and PSII electron transport activities in wild
type and transgenic plants. There were no significant differences in
PSI electron transport activities between wild type and transgenic
plants. Compared to wild type plants, transgenic i7 plants showed
no changes in PSII electron transport activity, whereas transgenic
i28 and i46 plants demonstrated a significant decrease in PSII electron transport activity.
PSII photochemistry
In order to investigate how levels of activase affect PSII electron
transport, we examined Fv /Fm in wild type and transgenic plants
(Table 3). Compared to wild type plants, there were no changes in
Fv /Fm in transgenic i7 plants, while there was a decrease in Fv /Fm
in transgenic i28 and i46 plants.
The decrease in Fv /Fm in transgenic i28 and i46 plants suggests
that PSII function is affected. In order to elucidate the functional
Table 3
Maximal efficiency of PSII photochemistry (Fv /Fm )
in wild type (WT) and transgenic i7, i28, and i46
tobacco plants.
Fv /Fm
WT
i7
i28
i46
0.83
0.82
0.71
0.64
±
±
±
±
0.02a
0.04a
0.05b
0.04c
Values in the table are means ± SE from three
independent experiments. Values indicated with
different letters were significantly different at
P = 0.05.
status of the donor and acceptor sides of the PSII complex in these
transgenic plants, the kinetics of the flash-induced chlorophyll
fluorescence yield were compared between wild type plants and
transgenic plants. The relaxation of the flashed-induced increase
in variable chlorophyll fluorescence yield monitors the oxidation
of QA − , which reflects the reoxidation of QA via forward electron
transport to QB (and QB − ) and back reaction with donor side components (Crofts and Wraight, 1983; Dau, 1994). Thus, through the
measurements of the relaxation of the variable fluorescence after
single flash excitation (Trtilek et al., 1997), it is possible to obtain
simultaneous information for the donor and acceptor side modifications of PSII (Vass et al., 1999, 2002).
In the absence of DCMU, the typical modulated fluorescence
decay of wild type plants after a single saturating flash could be
resolved into three distinct phases (Fig. 2A and B, Table 4). The
modulated fluorescence intensity relaxation in wild type plants
is dominated by the fast phase (t1/2 = 286 ␮s), whose relative
Fig. 2. The flash-induced decay of modulated chlorophyll fluorescence in wild type (WT) and transgenic i7, i28, and i46 tobacco plants. (A) The curves were the actual data
of the fluorescence signals in the absence of 20 ␮M DCMU. (B) The curves were normalized relative to the total variable fluorescence in the absence of 20 ␮M DCMU. (C) The
curves were the actual data of the fluorescence signals in the presence of 20 ␮M DCMU. (D) The curves were normalized relative to the total variable fluorescence in the
presence of 20 ␮M DCMU.
B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
1461
Table 4
Parameters of decay kinetics of flash-induced variable fluorescence in wild type (WT) and transgenic i7, i28, and i46 tobacco plants.
Total amplitude (%)
Without DCMU
WT
i7
i28
i46
With DCMU
WT
i7
i28
i46
100a
100 ± 1
92 ± 3
89 ± 1
100
98 ± 2
86 ± 2
65 ± 1
Fast phase t1/2 (␮s) [A (%)]
Middle phase t1/2 (ms) [A (%)]
Slow phase t1/2 (s) [A (%)]
286 ± 39 (60.3 ± 4.9)b
298 ± 49 (59.2 ± 4.2)
386 ± 49 (52.4 ± 3.9)
410 ± 49 (48.3 ± 3.1)
8.2
8.6
11.3
15.4
±
±
±
±
0.4 (25.1 ± 1.4)
0.5 (25.0 ± 1.8)
0.5 (20.3 ± 1.6)
0.5 (19.5 ± 1.2)
7.82
7.78
6.74
5.78
±
±
±
±
0.4 (14.6 ± 0.5)
0.5 (15.7 ± 0.7)
0.4 (26.5 ± 0.5)
0.6 (31.2 ± 0.6)
– (0)
– (0)
– (0)
– (0)
120
118
173
324
±
±
±
±
10 (10.8 ± 0.2)
11 (10.8 ± 0.4)
24 (15.9 ± 0.8)
29 (21.9 ± 0.4)
1.68
1.69
1.67
1.65
±
±
±
±
0.4 (89.2 ± 1.4)
0.3 (89.2 ± 1.7)
0.4 (84.1 ± 1.3)
0.4 (78.1 ± 1.8)
The relaxation of the flashed-induced fluorescence yield was measured without or with 20 DCMU. Exponential analysis yielded either three or two phasic kinetics with
different half-times (t1/2 ) and amplitudes (A). Mean ± SE values were calculated from four independent experiments.
a
Values represent the amplitude of total variable fluorescence as a percentage of that in wild type plants.
b
Values in parentheses are relative amplitude as a percentage of total variable fluorescence obtained from wild type and different transgenic tobacco plants.
amplitude is about 60%. The contribution of the middle phase
(t1/2 = 8.2 ms) was about 25%, and that of the slow phase (t1/2 = 7.8 s)
was about 15%. Compared to wild type plants, the fluorescence
decay kinetics after a single saturating flash showed no changes
in transgenic i7 plants, but there were significant changes in transgenic i28 and i46 plants. There was a gradual decrease in the total
fluorescence amplitude in transgenic i28 and i46 plants, suggesting that there was a loss of QA reduction. In addition, there was a
significant increase in the decay half-time of the fast phase, but a
significant decrease in the amplitude of the fast phase. The t1/2 of
the fast phase increased from 286 ␮s in wild type plants to 386 and
410 ␮s in transgenic i28 and i46 plants, respectively. The amplitude of the fast phase decreased from 60% in wild type plants to
52% and 48% in transgenic i28 and i46 plants, respectively. For the
middle phase, there was a significant increase in the decay halftime and a decrease in the amplitude. The t1/2 of the middle phase
increased from 8.2 ␮s in wild type plants to 11.3 and 15.4 ␮s in
transgenic i28 and i46 plants, respectively. The amplitude of the
middle phase decreased from 25% in wild type plants to 20.3% and
19.5% in transgenic i28 and i46 plants, respectively. For the slow
phase, there was a significant decrease in the decay half-time and
a significant increase in the amplitude of the slow phase. The t1/2
of the slow phase decreased from 7.8 s in wild type plants to 6.7
and 5.8 s in transgenic i28 and i46 plants, respectively. The amplitude of the slow phase increased from 15% in wild type plants to
27% and 31% in transgenic i28 and i46 plants, respectively. These
results suggest that there was a slow down in the middle phase, but
an acceleration in the slow phase in transgenic i28 and i46 plants.
When Chl fluorescence induction kinetics is determined in the
presence of DCMU, the fluorescence relaxation reflects the reoxidation of QA − via recombination with donor side components.
Compared to wild type plants, there was no significant change in
the modulated fluorescence decay kinetics in transgenic i7 plants,
while there were significant changes in the modulated fluorescence decay kinetics in transgenic i28 and i46 plants (Fig. 2C and
D, Table 4). The analyses of these fluorescence relaxation curves
showed that, in wild type plants, the decay was composed of a fast
phase and a slow phase (Table 4). There was no significant change in
the time constant for the slow component, but its relative amplitude decreased to 84% and 78% in transgenic i28 and i46 plants,
respectively. On the other hand, there was an increase in both time
constant and the relative amplitude in these transgenic plants for
the fast phase. Its relative amplitude increased from 11% in wild
type plants to 16% and 22% in transgenic i28 and i46 plants, respectively. Also, t1/2 increased from 120 ms in wild type plants to 173
and 324 ms in transgenic i28 and i46 plants, respectively. These
results suggest that, compared to wild type plants, the changes in
the fast phase in transgenic i28 and i46 plants may arise from the
recombination reactions of QA − with PSII donor side component(s).
It should be noted that the dynamics of the state transition S1 to
S2 was generally not influenced in the range from 5% to 100% of
activase concentration in the transgenic plants relative to the wild
type plants.
TL was further used to investigate the redox properties of the
acceptor and donor sides of PSII in wild type plants and transgenic plants, as it is a useful tool to study charge stabilization and
subsequent recombination in PSII in higher plants. Recombination
of positive charges stored in the S2 and S3 oxidation states of the
water-oxidizing complex with electrons stabilized on the reduced
QA and QB acceptors of PSII results in characteristic TL emissions
(Vass and Govindjee, 1996; Inoue, 1996). The TL intensity reflects
the amount of recombining charges, whereas the peak temperature
is indicative of the energetic stabilization of the separated charge
pair; the higher the peak temperature, the greater the stabilization
(Vass et al., 1981). Illumination of a single-turnover flash with the
plant sample after a short dark adaptation induces a major TL band
called the B-band, which appears at around 30 ◦ C and arises from
S2 /S3 QB − recombination (Inoue, 1996; Demeter and Vass, 1984;
Rutherford et al., 1982). TL emission following single flash excitation of dark-adapted plants results largely from the recombination
of the S2 QB − charge pair. If electron transfer between QA and QB is
blocked by DCMU, the B-band is replaced by the so-called Q-band
arising from S2 QA − recombination at around 10 ◦ C (Rutherford et
al., 1982).
Fig. 3 shows the TL glow curves in wild type and transgenic
plants. The wild type plants exhibited TL emission maxima for
the S2 QB − and S2 QA − charge recombination at approximately 35
and 9 ◦ C, respectively. Compared to wild type plants, there was an
upshift in the peak temperatures for the S2 QB − and S2 QA − charge
recombinations in transgenic i46 plants. In transgenic i46 plants,
the peak temperatures for the S2 QB − and S2 QA − charge recombination were at approximately 39 and 14 ◦ C, respectively (Fig. 3,
Table 5
Temperature at highest emission of the TL curves in wild type (WT) and transgenic
i7, i28, and i46 tobacco plants.
Peak temperature (◦ C)
Without DCMU
WT
i7
i28
i46
34.5
34.8
35.3
37.1
±
±
±
±
a
0.1
0.2a
0.4a
0.4b
With DCMU
9.0
9.2
9.5
13.7
±
±
±
±
0.2a
0.3a
0.3a
0.2b
The measurements were performed in the absence and presence of 10 ␮M DCMU.
Mean ± SE values were calculated from four to six independent experiments.
Values indicated with different letters were significantly different at P = 0.05.
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B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
Fig. 3. Thermoluminescence glow curves in wild type (WT) and transgenic i7, i28, and i46 tobacco plants. The leaves were excited with a single flash in the absence (A) and
presence (B) of 20 ␮M DCMU.
Table 5). Since the temperature at which maximal luminescence
occurs is a function of the free energy of stabilization of the chargeseparated state, the upshifts in the peak temperatures of the S2 QB −
and S2 QA − recombination observed in transgenic i28 and i46 plants
indicate that there was an increase in the stabilities of both chargeseparated states. Moreover, since the increased stabilizations of the
S2 QB − and S2 QA − recombination occurred approximately in parallel, it seems likely that the changes in TL emission maxima for
the S2 QB − and S2 QA − were due largely to a modification of the
oxygen-evolving complex specifically affecting the charge transfer
characteristics of the S2 state.
Fig. 4 shows the oscillation of the B-band in wild type and transgenic plants as a function of the number of flashes given prior to
recording the TL curves. In wild type plants, the intensity of the
B-band oscillated with a periodicity of four, with the maximum
emission occurring on the second flash. This is the same type of
oscillation observed for whole leaf tissue (Rutherford et al., 1984).
The oscillatory pattern was not changed in transgenic i7 plants. On
the other hand, the oscillatory pattern in transgenic i28 plants was
gradually damped compared to wild type plants. In transgenic i46
plants, the period-four oscillation could almost not be observed.
Thylakoid peptide composition
Fig. 5A shows the effects of lowered activase content on thylakoid peptide composition in wild type and transgenic plants.
Compared with wild type plants, the amount of D1 protein did
not show a significant change in transgenic i7 plants, but showed a
decrease in transgenic i28 and i46 plants, which were about 86% and
74% of wild type plants, respectively. The amount of D2, CP43, and
CP47 showed a significant decrease only in transgenic i46 plants,
but not in transgenic i7 and i28 plants. There was no significant
change in the amount of PsbO protein in transgenic i7 plants, while
the amount of PsbO in transgenic i28 and i46 plants was 83% and
68% of that of wild type plant, respectively. There was a slight
increase in the level of LHCII in transgenic i46 plants. The levels of
PSI-Psa A/B proteins, Cytochrome (Cyt) f and ␤-subunit of the ATP
synthase (CF1␤) did not show significant changes between wild
type and transgenic plants.
To investigate structural alterations of thylakoid proteins,
chlorophyll–protein complexes were solubilized from thylakoid
membranes using dodecyl-␤-d-maltopyranoside (DM) and separated by blue native PAGE (BN-PAGE) (Fig. 5B). After the
first-dimensional separation in the presence of Coomassie blue G
250 dye, five major bands were resolved, apparently representing
monomeric PSI and dimeric PSII (band I), monomeric PSII (band
II), CP43 minus PSII (band III), trimeric (band IV), and monomeric
LHCII (band V) (Peng et al., 2006). The BN-PAGE analysis showed
that the monomeric PSI and dimeric PSII (bands I) decreased slightly
in transgenic i28 plants but significantly in transgenic i46 plants as
compared with wild type plants. Analyses of the two-dimensional
SDS-urea-PAGE gels after Coomassie blue staining confirmed similar results, as shown in Fig. 5A.
Chloroplast development
Fig. 4. The flash-induced oscillation of the B thermoluminescence band in the whole
leaves of wild type (WT) and transgenic i7, i28, and i46 tobacco plants. Values
represent mean of 6–8 independent measurements.
The results showed that the lowered content of activase resulted
in a decrease in the activity of PSII electron transport, so we investigated whether the lowered activase content affects thylakoid
membrane development. Fig. 6 shows electron micrographs of
chloroplasts of wild type and transgenic plants. There was no significant difference in chloroplast development between wild type
and transgenic i7 and i28 plants. However, compared to wild type
plants, transgenic i46 plants showed a significant decrease in the
number of grana stacks per chloroplast and discs per grana stack
(Table 6).
B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
1463
Fig. 5. (A) Immunological analysis of peptide composition of the thylakoid membranes (5 ␮g chlorophyll) in wild type (WT) and transgenic i7, i28, and i46 tobacco plants.
(B) BN gel analysis of thylakoid membranes (10 ␮g chlorophyll) in wild type (WT) and transgenic i7, i28, and i46 tobacco plants. (C) Two-dimensional separation of protein
complexes in the thylakoid membranes in wild type (WT) and transgenic i7, i28, and i46 tobacco plants.
Discussion
In this study, we obtained transgenic tobacco plants with
decreased levels of Rubisco activase compared to wild type plants.
Our results show that when the level of activase was decreased
to 25% of wild type plants, there was a decrease in CO2 assimilation rate (Table 1), which is similar to the results reported in
previous studies (Jiang et al., 1994; Mate et al., 1996; Eckardt et al.,
1997; Jin et al., 2006). However, there is no report on how lowered
activase content regulates photosynthetic electron transport. Our
results show that the lowered activase content had no effect on
PSI electron transport, but resulted in a significant decrease in PSII
electron transport, which is reflected in a decrease in PSII activity
when the level of activase was decreased to 25% of wild type plants
(Tables 2 and 3). We further investigated how PSII photochemistry
is regulated in these transgenic tobacco plants.
Flash-induced chlorophyll fluorescence kinetics is useful to
study the electron transfer at the donor and acceptor sides of PSII.
The double-modulation technique makes it possible to study the
reoxidation processes of QA − by both forward and back reactions
(Trtilek et al., 1997). Analyses of modulated fluorescence decay
kinetics show that a significant increase in the decay half-time of
the fast phase and a significant decrease in its amplitude in transgenic i28 and i46 plants (Fig. 2, Table 4), indicating clearly that
lowered activase content results in an inhibition of electron transfer from QA − to QB . In order to analyze whether the inhibition of
Table 6
Parameters of chloroplast development in wild type (WT) and transgenic i7, i28, and i46 tobacco plants.
Number of grana stacks per chloroplast
WT
i7
i28
i46
18.3
17.9
16.7
13.3
±
±
±
±
1.8a
2.3a
3.5a
2.1b
Number of discs per grana stack
22.3
22.1
18.1
11.6
±
±
±
±
2.1a
2.3a
2.8a
1.1b
Length of the chloroplast (␮m)
5.9
5.9
5.8
4.9
±
±
±
±
0.28a
0.36a
0.24a
0.19b
Values in the table are means ± SE from three independent experiments. To quantify the differences between chloroplasts of wild type and different transgenic plants, totally
64 chloroplast sections were analyzed from the youngest fully expanded leaves. Values indicated with different letters were significantly different at P = 0.05.
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B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
Fig. 6. Electron micrographs of chloroplasts in wild type (WT) and transgenic i7, i28, and i46 tobacco plants. To quantify the differences between wild type and different
transgenic plants chloroplasts, 64 chloroplast sections were analyzed from the youngest fully expanded leaves. Bars = 1 ␮m.
electron transfer from QA − to QB is associated with the decrease in
the apparent equilibrium constant for sharing the electron between
QA − and QB in transgenic i26 and i48 plants, we compared the t1/2 of
the slow phase in the absence or presence of DCMU. In the absence
of DCMU, the t1/2 of the slow phase in wild type plants was about
7.8 s. However, in the presence of DCMU, the t1/2 of the slow phase
decreased to about 1.7 s in wild type plants (Table 4). The slow
phase reflects the reoxidation of QA − via charge recombination with
the S2 state. In the presence of DCMU, the slow phase occurs from
the S2 QA − recombination in PSII centers. Thus, the differences in
the t1/2 indicate that, in the absence of DCMU, the recombination
should occur from the QA QB − state, which is in charge equilibrium
with the QA − QB state. Therefore, the inhibition of QA − to QB in transgenic i28 and i46 plants may be associated with the decrease in the
apparent equilibrium constant for sharing the electron between
QA − and QB , which may be due to a decreased affinity of QB binding
pocket. Indeed, we observed that there was an increase in the t1/2 of
the middle phase in the absence of DCMU in transgenic i28 and i46
plants (Table 4). This decay component is assigned to reoxidation
of QA − in the centers in which a vacant QB site has to be reoccupied
by a PQ molecule before the QA − to QB electron transfer can take
place. Thus, an increase in the t1/2 of the middle phase in transgenic
i28 and i46 plants suggests that there were modifications of the QB
niche by which PQ binding is slowed down.
In this study, we observed that the time rate constant of the
middle phase of the flash fluorescence relaxation recorded in the
presence of DCMU increased significantly in transgenic i28 and i46
plants (Table 4), suggesting that electron transport at the donor
side of PSII is affected. To investigate how electron transport at the
donor side is regulated in transgenic i28 and i46 plants, we examined the characteristics of TL in wild type and transgenic plants.
Our results show that there was a parallel increase in the stability of the S2 QA − and S2 QB − charge recombination in transgenic
i48 plants compared with wild type plants (Fig. 3, Table 5). Our
results also show that the period-four oscillation was significantly
dampened in transgenic i46 plants (Fig. 4). These results suggest
that there is an increase in the redox stability of the S2 state in
transgenic i46 plants. It has been shown that the loss of the PsbO
protein is accompanied by an increase in the stability of the S2 state
by the removal of the PsbO protein from PSII preparations in vitro
by measuring TL profiles and by monitoring deactivation kinetics
of the S states (Miyao et al., 1987; Vass et al., 1987). Moreover,
the mutant lacking the PsbO protein results in upshifts in the peak
temperatures of S2 QA − and S2 QB − recombination by determining
TL profiles and thus leads to an increase in the stability of the S2
state in a cyanobacterium Synechocystis sp. PCC 6803 (Burnap et al.,
1992). Based on these reports and our results, it is suggested that
the increased stability of the S2 state in transgenic i46 plants may
be due to the decreased PsbO protein in the thylakoid membranes.
To investigate this possibility, we examined the content of the PsbO
protein in wild type and transgenic plants. We observed that there
was a significant decrease in the content of PsbO protein in the
thylakoid membrane in transgenic i28 and i46 plants (Fig. 5). The
changes in the content of the PsbO protein in the thylakoid membranes on a chlorophyll basis in wild and transgenic plants suggest
that the increased stability of the S2 state in transgenic i46 plants
is associated with the decrease in the content of the PsbO protein
in the thylakoid membranes.
In this study, we observed the pale-green phenotype in transgenic i28 and i46 plants (Fig. 1). We thus investigated chloroplast
structure in wild type and transgenic plants. Our results show
that chloroplast development in transgenic i28 and i46 plants was
defective, with less grana compared to wild type plants (Fig. 6). It
seems that the defective nature of the chloroplast was not due to
the change in PSI function since there was no difference in PSI electron transport activity between wild type and transgenic plants
(Table 2). On the other hand, the defected chloroplast in transgenic i28 and i46 plants may be associated with the PSII function, as
B. Cai et al. / Journal of Plant Physiology 167 (2010) 1457–1465
we observed that there was a significant decrease in PSII electron
transport activity and the maximal efficiency of PSII photochemistry (Tables 2 and 3). In addition, we observed that there was a
decrease in the content of D1 and PsbO proteins in transgenic i28
and i46 plants (Fig. 5). Overall, the ultrastructural defects observed
in chloroplasts of transgenic i28 and i46 plants suggest that activase
may be required for chloroplast biogenesis.
It should be noted that the levels of the D1 protein were significantly reduced in transgenic i28 and i46 plants, while the levels of
other PSII proteins were not decreased in these transgenic plants
(Fig. 5). These results suggest that the D1 protein is more sensitive to activase deficiency as compared to other PSII proteins. The
significant decrease in D1 protein transgenic i7 and i28 plants can
be explained by aggravated photoinhibition due to the significant
decrease in CO2 assimilation rate in these transgenic plants.
In conclusion, the results in this study demonstrate that activase plays an important role not only in CO2 assimilation, but also
in PSII function. Activase deficiency led to a decrease in PSII activity that was associated with an inhibition of electron transfer at
the acceptor side and the donor side. The decrease in PSII function resulted in the defects in the development of chloroplasts.
Our results suggest that activase plays an important role in maintaining PSII function and chloroplast development. The results in
this study also suggest that there is a homoeostasis in respect to
photosynthetic electron transport and CO2 fixation upon a genetic
down regulation of Rubisco activase to below 50% of the control.
The molecular mechanisms involved in how activase regulates the
PSII function remains to be investigated further.
Acknowledgments
This study was supported by the State Key Basic Research
and Development Plan of China (2009CB118503) and the Frontier Project of the Knowledge Innovation Engineering of Chinese
Academy of Sciences (KSCX2-YW-N-042).
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