Plant Cell Physiol. 49(12): 1879–1886 (2008)
doi:10.1093/pcp/pcn170, available online at www.pcp.oxfordjournals.org
ß The Author 2008. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: [email protected]
Imaging of NPQ and ROS Formation in Tobacco Leaves: Heat Inactivation
of the Water–Water Cycle Prevents Down-Regulation of PSII
Éva Hideg
1
2
1,
*, Péter B. Kós
1
and Ulrich Schreiber
2
Institute of Plant Biology, Biological Research Center, Szeged, 6701 Hungary
Julius-von-Sachs Institut für Biowissenschaften, Universität Würzburg, D-97070 Germany
Non-photochemical chlorophyll fluorescence quenching
(NPQ) plays a major role in the protection of the
photosynthetic apparatus against damage by excess light,
which is closely linked to the production of reactive oxygen
species (ROS). The effect of a short heat treatment on
NPQ and ROS production was studied with detached
tobacco leaves by fluorescence imaging of chlorophyll and
of the ROS sensor dye HO-1889NH. NPQ was stimulated
`3-fold by 3 min pre-treatment at 448C, in parallel with
suppression of CO2 uptake, while no ROS formation could
be detected. In contrast, after 3 min pre-treatment at 468C,
NPQ was suppressed and ROS formation was indicated by
quenching of HO-1889NH fluorescence. After 3 min pretreatment at 468C and above, partial inactivation of
ascorbate peroxidase and light-driven accumulation of
H2O2 was also observed. These data are discussed as
evidence for a decisive role of the Mehler ascorbate
peroxidase or water–water cycle in the formation of the
NPQ that reflects down-regulation of PSII.
Keywords: Fluorescence imaging — Heat stress-reactive
oxygen species (ROS) — Mehler ascorbate peroxidase
(MAP) cycle — PAM fluorometry — Photosynthetic
electron transport.
Abbreviations: APX, ascorbate peroxidase (EC
1.11.1.11); DAB, diamino benzidine tetrahydrochloride;
Fm, maximum fluorescence yield in the dark-acclimated
state; Fo, minimum fluorescence yield in the darkacclimated state; Fm0 , maximum fluorescence yield in the
light-acclimated state; Fv/Fm, maximum photochemical
yield of PSII in the dark-acclimated state; HO-1889NH,
3-(N-dansyl)aminomethyl-2,2,5,5-tetramethyl-2,5-dihydro1H-pyrrole; LED, light-emitting diode; MAP cycle, Mehler
ascorbate peroxidase cycle; MDA, monodehydroascorbate;
MDAR, monodehydroascorbate reductase; NPQ, nonphotochemical quenching; PAM, pulse amplitude modulation; PAR, photosynthetically active radiation; ROS,
reactive oxygen species; Y(II), photochemical yield.
Introduction
Chlorophyll fluorescence allows deep insights into the
complex process of photosynthesis in vivo (see review in
Papageorgiou and Govindjee 2004). In particular, pulse
amplitude modulation (PAM) fluorometry and fluorescence
quenching analysis by the saturation pulse method have
proven useful for non-invasive assessment of the efficiency
of photosynthetic energy conversion (Schreiber 2004).
Measurements of non-photochemical quenching, mostly
expressed by the parameter NPQ (Bilger and Björkman
1990), have allowed the mechanisms that protect plants
from damage by excess light to be studied (for reviews, see,
for example, Demmig-Adams and Adams 1992, Osmond
1994, Horton et al. 1996, Müller et al. 2001). There is
general consensus that NPQ reflects the dissipation of
excess excitation energy in the form of harmless heat (downregulation of PSII), thus protecting the plant from the
damaging effects of reactive oxygen species (ROS). The
majority of studies on NPQ have been focused on the
underlying molecular mechanisms, emphasizing the pivotal
roles of the trans-thylakoid proton gradient (pH) and
zeaxanthin. Less attention has been paid to the question of
what kind of electron flow is responsible for the formation
of the pH that drives violaxanthin de-epoxidation and
renders zeaxanthin an efficient quencher, when CO2dependent electron flow is limiting. This happens during
the first minute of illumination after dark adaptation, when
Calvin–Benson cycle enzymes are not yet light activated or
in stressed leaves when these enzymes are damaged
(Schreiber and Bilger 1987).
Whenever CO2 is not available, molecular oxygen may
substitute as electron acceptor, either in photorespiration
(Osmond 1981) or in the Mehler ascorbate peroxidase
(MAP) cycle (Nakano and Asada 1981, Asada and Badger
1984, Asada and Takahashi 1987). Electron flow associated
with photorespiration is characterized by a higher ATP/
NADPH ratio than CO2 fixation and, hence, does not
qualify for NPQ generation. On the other hand, no ATP is
*Corresponding author: E-mail, [email protected]; Fax, þ36-62-433-434.
1879
NPQ and ROS imaging in heat-treated leaves
consumed in the MAP cycle, so that in the intact system
even low rates of this cycle can build up an appreciable
pH (Hormann et al. 1993, Hormann et al. 1994). Hence, it
has been proposed that O2-dependent electron flow in the
MAP cycle is mainly responsible for generation of the NPQ
associated with the down-regulation of PSII (Schreiber and
Neubauer 1990, Neubauer and Yamamoto 1992, Schreiber
et al. 1995). One turnover of this cycle (transient formation
of one H2O2) consumes 8 quanta and leads to the
translocation of at least 16 Hþ from the stroma to the
lumen. As all H2O2 that is formed via O2 reduction is
eventually re-reduced to H2O and as the electrons for the
reduction of O2– and H2O2 originate from the splitting of a
stoichiometric amount of H2O in PSII, the name ‘water–
water cycle’ was coined for this sequence of reactions
(Asada 1999).
A central role for the water–water cycle in generating
NPQ in intact leaves has been questioned by Heber and
co-workers (Wu et al. 1991, Kobayashi and Heber 1994,
Heber 2002), who favor cyclic PSI electron flow instead,
with the water–water cycle only serving a ‘poising’ function
to relieve the intersystem electron transport chain from
‘over-reduction’. This view has been supported by
Munekage et al. (2002) who explained the suppression of
NPQ in a ‘proton gradient-deficient’ Arabidopsis mutant
(PGR5) by a defect in the ferredoxin-dependent cyclic PSI
pathway. These authors, however, explicitly do not rule out
the possibility ‘that O2 reduction is mediated by the PGR5dependent pathway’. The main reason why this important
question is not clarified yet is the fact that to date no
methods are available to quantify reliably the fluxes of the
water–water cycle and cyclic PSI in illuminated intact
leaves. Both reactions can be identified only indirectly, via
the NPQ and the ‘scattering change’ associated with pH
formation.
Recently we introduced a new PAM fluorescence
imaging system, which allows parallel imaging of NPQ
and ROS formation via fluorescence quenching of the ROS
sensor dye HO-1889NH (Hideg and Schreiber 2007). In this
previous study, the performance of the new system was
demonstrated after leaf infiltration with methyl viologen in
order to induce ROS formation upon illumination. In the
present study, a mild, short heat treatment is applied for the
same purpose, as a stress treatment that may also occur in
the natural environment. Schreiber and Klughammer (2008)
very recently showed that with increasing treatment
temperatures NPQ was first substantially stimulated (at
44–468C in outdoor-grown rose leaves) but then was
strongly suppressed at only 28C higher temperature. In
this study, the heat-induced suppression of NPQ was
assumed to be caused by an inhibition of the water–water
cycle, while the heat-induced stimulation of NPQ was
explained by inhibition of CO2 fixation (Bilger et al. 1987).
If indeed both the Calvin–Benson cycle and the water–water
cycle would be inactivated, the suppression of NPQ should
be correlated with ROS formation. It is the aim of the
present study to check on this hypothesis by parallel
imaging of NPQ and fluorescence quenching of a ROS
sensor.
Results
Figure 1 shows that while at up to 428C there was only
little inhibition, 3 min exposure to 448C caused about 75%
inhibition of CO2 uptake capacity. In leaves exposed to
468C, CO2 uptake was completely blocked.
With the same leaf material, chlorophyll fluorescence
parameters were imaged after 3 min heat treatment
(Figs. 2, 3). By exposing only half the areas of the leaves
to the heat, any heat-induced change in photosynthetic
parameters can be readily distinguished by comparison with
the untreated half. Fig. 2 shows that, in agreement with the
CO2 uptake data in Fig. 1, the 428C pre-treatment did not
have any effect on Fv/Fm and the photochemical yield
[Y(II)], both of which displayed a very homogenous
distribution over the imaged area. The non-photochemical
quenching parameter, NPQ, showed some heterogeneity
along the leaf veins, which was more pronounced in the
untreated half. Distinct changes were induced by treatment
at 448C, which were particularly pronounced in Y(II) (large
decrease) and NPQ (large increase), whereas Fv/Fm was just
marginally decreased. It may be noted that heating not only
affected the heated half of the leaf, but also spread to some
extent into the untreated half. Exposure to 468C resulted in
further decreases of Fv/Fm and Y(II), and almost complete
suppression of NPQ. In contrast to the leaf treated at 448C,
there is a sharp border between the untreated and treated
halves of the leaf, which is characterized by very high NPQ
10.0
Photosynthesis (µmol CO2 m−2 s−1)
1880
untr
40°C
42°C
44°C
46°C
7.5
5.0
2.5
0.0
0
250
500
PAR (µmol
750
1000
m−2 s−1)
Fig. 1 Photosynthetic carbon dioxide uptake in tobacco leaves
pre-treated for 3 min at the indicated temperatures (n ¼ 4).
NPQ and ROS imaging in heat-treated leaves
(deep blue, equivalent to 1.8–1.9). As temperatures in this
border area are uncertain, these areas were not included in
data averaging.
For a more detailed analysis, the dark–light induction
kinetics of Y(II) and NPQ in terms of dependence on the
A
Fv/Fm
0.8
Fv/Fm
Y(II)
B
0.0
Y(II)
C
2.0
NPQ
NPO
0.0
Fig. 2 Images of various chlorophyll fluorescence parameters of
32 24 mm center parts of detached tobacco leaves. The lower
halves of the imaged areas were exposed to 3 min heat treatment at
the indicated temperatures, while the upper parts were untreated.
Images are color coded according to the patterns shown next to the
images. The following parameters derived by the saturation pulse
method are shown: A, Fv/Fm; B, Y(II) at 75 mmol m–2 s–1 PAR; C,
NPQ at 75 mmol m–2 s–1 PAR.
pre-treatment temperature are presented in Fig. 3A and B,
respectively. At the applied moderate light intensity of
75 mmol m–2 s–1 photosynthetically active radiation (PAR),
in untreated control samples the kinetics reflect the
activation of CO2 fixation in the Calvin–Benson cycle
(Schreiber et al. 1986): during the first minute after onset of
illumination, Y(II) decreases, whereas NPQ increases.
Subsequently, Y(II) rises to a high level in the steady
state, whereas NPQ drops to a low steady-state level. These
transients have been interpreted as the ‘fingerprint’ of an
energizing, non-assimilatory electron flow, which after dark
adaptation activates Calvin–Benson cycle enzymes and thus
primes CO2 fixation (Schreiber and Bilger 1987). This
interpretation is supported by the present data.
Both measurements of CO2 uptake and chlorophyll
fluorescence parameters show a clearly defined critical
threshold between 42 and 448C treatment temperatures,
below which CO2 uptake and fluorescence induction
kinetics essentially are not affected and above which CO2
uptake is inhibited, while the characteristic increase of Y(II)
and decrease of NPQ are suppressed. The physiological
state after 3 min pre-treatment at 448C is of special interest,
because it is characterized by high NPQ in the light, which
reflects down-regulation of PSII and thus protection of the
plant from photoinhibitory damage (for reviews, see
Demmig-Adams and Adams 1992, Horton et al. 1996,
Müller et al. 2001). The question is, what kind of protoncoupled electron transport is responsible for this high NPQ
(see Introduction and Discussion). In this context, it is
important that treatment at just 28C higher temperature
(3 min at 468C) causes complete suppression of NPQ.
Hence, there is another clearly defined critical threshold
between 44 and 468C, which reflects the inactivation of the
protective NPQ-generating reaction and thus should help to
elucidate the identity of these reactions.
B
1.5
unt
38°C
40°C
42°C
44°C
46°C
48°C
0.6
NPQ
Y(II), Photochemical yield
A
1881
0.3
1.0
0.5
0.0
0.0
0
50
100
150
Time (s)
200
250
300
0
50
100
150
200
250
300
Time (s)
Fig. 3 Effect of heat treatment on dark–light induction kinetics of chlorophyll fluorescence parameters in tobacco leaves. (A) Effective
photochemical quantum yield Y(II); (B) non-photochemical quenching NPQ. Illumination at 75 mmol m–2 s–1 PAR. Treatments and
measurements are as described for Fig. 2 (n ¼ 3).
A
NPQ and ROS imaging in heat-treated leaves
B
1.0
C
D
ROS sensor
fluorescence
(a.u.)
0.0
Fig. 4 Fluorescence images of 18 24 mm tobacco leaf segments
infiltrated through two pinholes with a 2 mM solution of HO1889NH after 3 min heat exposure to 38 and 468C, as indicated in
the figure. Images were taken before (A, C) and after (B, D)
exposure to 25 mmol m–2 s–1 PAR for 30 min. Fluorescence
intensities are color coded according to the pattern shown next
to the images.
If, as has been suggested before, the water–water cycle
is responsible for generation of the protective NPQ
(Schreiber and Neubauer 1990, Schreiber et al. 1995,
Asada 1999), the abrupt suppression of NPQ by 3 min
treatment at temperatures above 448C should be due to the
inactivation of one or several of the enzymes involved in this
cycle, namely superoxide dismutase (SOD), ascorbate
peroxidase (APX) and monodehydroascorbate reductase
(MDAR), so that ROS can accumulate upon illumination.
The following experiments were designed to check the
validity of this hypothesis. ROS were detected on the basis
of fluorescence quenching of a sensor probe (HO-1889NH),
a solution of which was fed via pinholes into untreated and
heat-pre-treated leaf halves, as illustrated in Fig. 4. For
optimal differentiation between heated and untreated leaf
halves, a relatively long illumination time (30 min) at
relatively low intensity (25 mmol m–2 s–1) proved optimal.
Fig. 4A and C shows fluorescence images before, and
Fig. 4B and D after illumination. While generally no lightinduced quenching was observed in the untreated leaf
halves or in leaves treated at lower temperatures (Fig. 4A,
B), HO-1889NH fluorescence decreased markedly in the
leaf half exposed to 468C (Fig. 4D), which clearly reflects
formation of ROS. A quantitative analysis based on such
images is presented in Fig. 5, which shows that the threshold
temperature for ROS detection was between 44 and 468C.
Because HO-1889NH is not reactive to H2O2, this species
was detected on the basis of observing brown coloring in
leaves treated with DAB (diamino benzidine tetrahydrochloride). This was most pronounced in leaves preexposed to 488C and only slightly noticeable in response to
468C (Fig. 6).
HO-1889NH fluorescence (% of untreated)
1882
100
80
*
60
*
40
20
0
untr
38
40
42
44
46
48
Pre-treatment temperature (˚C)
Fig. 5 Effect of 3 min heat treatment at 38–488C on relative
intensity of HO-1889NH fluorescence in infiltrated tobacco leaves.
Fluorescence quenching indicates light-induced ROS production.
Fluorescence intensities were averaged from approximatelty 1 cm–2
areas of the HO-1889NH-infiltrated leaf segments (from four
different images like the ones shown in Fig. 4) before and after
exposure to 25 mmol m–2 s–1 PAR for 30 min. Changes in average
intensities are shown as a percentage of mean fluorescence
intensity before illumination. Error bars represent standard deviations. Values significantly (P50.05, in a paired t-test) different from
the activity measured in untreated samples are marked with an
asterisk.
Fig. 7 demonstrates a strong temperature sensitivity of
APX. Enzyme activity was decreased by approximately
30% even by the relatively short, 3 min heat treatments, if
these were performed at 468C or higher temperatures. This
was due to a loss of function and not to any loss of APX
protein, as illustrated by the protein gel immunoblot in
Fig. 8. This experiment was carried out using heat-treated
pea leaves, because commercially available antibodies were
not reactive with tobacco APX.
Discussion
NPQ is induced whenever pH formation exceeds the
consumption of pH in ATP synthesis for CO2 fixation
(and photorespiration). In Fig. 1 it was demonstrated that
CO2 uptake in tobacco leaves can be inhibited by a 3 min
heat treatment at 448C. The same treatment resulted in
43-fold stimulation of NPQ by 3 min heat treatment at
448C and almost complete suppression of NPQ by 3 min
heat treatment at 468C (Figs. 2, 3). The same critical
treatment temperature was also observed for the occurrence
of light-induced fluorescence quenching of the ROS sensor
dye HO-1889NH (Figs. 4, 5), for DAB coloration (Fig. 6)
and for inactivation of APX (Fig. 7). This is the first direct
demonstration of ROS formation in leaves under in vivo
conditions. It shows that ROS are formed and can cause
APX (mM mg protein−1 S−1)
NPQ and ROS imaging in heat-treated leaves
38°C
40°C
42°C
44°C
46°C
1883
30
*
*
20
10
48°C
0
Fig. 6 DAB staining for hydrogen peroxide in cuttings from
tobacco leaves pre-exposed to heat for 3 min. Leaves were kept
under 25 mmol m–2 s–1 PAR for 2 h in the DAB solution before
chlorophyll was removed.
damage as soon as the NPQ-generating process is inhibited,
which can be induced by heat inactivation of the APX.
These findings suggest that the ROS-scavenging and NPQgenerating processes are closely linked. Hence, the
presented data support the hypothesis that the MAP or
water–water cycle is responsible for formation of the pH
that induces NPQ in leaves and thus provides 2-fold
protection of the plant against damage by ROS and
excess radiation (Asada 1999).
A close link between NPQ formation and ROS
scavenging has already been established. Previous work
with intact and class D spinach chloroplasts has shown that
for pH and NPQ formation the presence of molecular
oxygen (apparent Km of 60 mM) and ascorbate (apparent
Km of 7 mM) is essential (Schreiber and Neubauer 1990,
Schreiber et al. 1995). Furthermore, it was previously shown
that it is the reduction of the H2O2 formed via the Mehler
reaction and superoxide dismutation, involving enzymatic
activity of APX and MDAR (Asada and Takahashi 1987,
Asada 1999), which is mainly responsible for energydependent NPQ (Schreiber et al. 1991). In agreement with
these findings, we now show that the suppression of room
temperature NPQ by heat pre-treatment indeed occurs
together with accumulation of H2O2 (Fig. 6) and a
suppression of APX activity (Figs. 7, 8). Therefore, it
appears likely that it is the heat inactivation of the
chloroplast APX which causes the suppression of NPQ
depicted in Figs. 2 and 3, the formation of ROS
demonstrated in Figs. 4 and 5, and the accumulation of
H2O2 shown in Fig. 6.
Upon heating of leaves, electron donation to the
intersystem electron transport chain in the dark is
reportedly stimulated (Schreiber et al. 1976, Havaux
1996). The resulting acceleration of P700þ re-reduction
untr
38
40
42
44
46
48
Pre-treatment temperature (°C)
Fig. 7 Activity of the ascorbate peroxide enzyme in extracts made
from tobacco leaves after a 3 min exposure to heat as detailed in
Materials and Methods. Symbols correspond to mean values, and
error bars show standard deviations (n ¼ 4). Values significantly
(P50.05, in a paired t-test) different from the activity measured in
untreated samples are marked with an asterisk.
Untr
42°C
46°C
48°C
Fig. 8 Protein gel blot analysis of ascorbate peroxidase content of
heat-pre-treated pea leaves (see Materials and Methods for details).
following oxidation by far-red light and the NPQ generated
by far-red illumination have been discussed as evidence for
a role for cyclic PSI in the down-regulation of PSII reflected
in NPQ (Bukhov et al. 1999). Hence, it could be argued that
the stimulation of NPQ after 3 min pre-treatment at 448C
demonstrated in Figs. 2 and 3 is due to stimulation of cyclic
PSI. Such a role for cyclic PSI electron flow in NPQ
formation was also suggested by Heber and co-workers (Wu
et al. 1991, Kobayashi and Heber 1994, Heber 2002). The
following points, however, argue against this interpretation.
(i) Far-red light-induced electron flow does not necessarily
correspond to cyclic PSI, particularly if a sufficiently large
electron pool at the donor side and molecular oxygen at the
acceptor side are available. (ii) PSI is known to display
relatively high heat stability and heat-induced acceleration
of P700 re-reduction was still observed after 15 min at 518C
(Havaux 1996), whereas in our experiments the lightinduced NPQ was almost completely suppressed after
3 min treatment at 468C (Figs. 2, 3). (iii) Reising (1994)
showed by simultaneous measurements of pulse-modulated
fluorescence and photoacoustics of tobacco leaves that
almost 30% of linear electron flow persisted after inactivation of CO2 fixation at 448C. As this linear flux was not
reflected by a net photobaric signal, the underlying O2
1884
NPQ and ROS imaging in heat-treated leaves
evolution must have been balanced by an equivalent O2
uptake. The latter was also reflected by strong stroma
alkalization (Reising and Schreiber 1992). In agreement
with the data of the present study, both linear flux and
stroma alkalization were suppressed above 458C. (iv) A
recent domain model of Albertsson (2001) suggests that
cyclic PSI is restricted to the stroma lamellae, whereas the
PSI located in the margin region is mainly responsible for
linear electron transport. To our knowledge, there is no
evidence for a control of PSII, most of which is located in
the stacked grana region, via the pH formed in the stroma
lamellae.
The observed suppression of NPQ in heat pre-treated
tobacco leaves strongly resembles that in the PRG5 mutant
of Arabidopsis (Munekage et al. 2002). While Munekage
et al. favored a role for PGR5 in cyclic electron flow around
PSI, they did not exclude the possibility that the activity of
the water–water cycle was also affected by the mutation and
suggested that the PGR5 mutation may lead to pleiotropic
effects on both cyclic PSI and the water–water cycle. They
also showed that removal of O2 leads to strong limitation at
the PSI acceptor side, which argues for a substantial O2dependent electron flow, i.e. the reduction of O2 and
monodehydroascorbate (MDA). Such electron flow can
protect the photosynthetic apparatus in two ways: first, by
relieving the PSI acceptor side, thus preventing PSI
photoinhibition (Sonoike and Terashima 1994); and,
secondly, by pH/NPQ formation and the resulting
down-regulation of PSII, with a direct photoprotective
effect on PSII as well as an indirect photoprotective effect
on PSI due to decreased ‘electron pressure’. In the pro/
contra discussion of cyclic PSI vs. the water–water cycle it is
often overlooked that cyclic PSI cannot relieve the PSI
acceptor side as much as a reduction of O2 and MDA,
because during the cyclic flow every electron taken up at the
acceptor side is fed into the donor side, from where it is
returned to the acceptor side again. The indirect effect of
cyclic PSI via pH/NPQ formation, however, is well known
to depend on ‘poising’ by O2-dependent electron flow. This
suggests that in any case cyclic PSI is functionally linked to
the water–water cycle. The decisive question remains what
are the relative fluxes of these two processes. For a
regulatory function, the fluxes do not need to be high.
Analogous heat treatment studies with Arabidopsis wild
type and the PRG5 mutant may contribute to a better
understanding of these processes.
In conclusion, the presented data emphasize the 2-fold
protective function of the water–water cycle, as previously
outlined by Asada (1999): first, whenever O2 is reduced by
PSI, the generated ROS is scavenged and in this way
enzyme damage is prevented. Secondly, in the course of this
scavenging process an exceptionally efficient electron
acceptor, MDA, is formed, reduction of which by linear
electron transport generates the NPQ that leads to downregulation of PSII and thus to protection against damage by
excess radiation. In the heat-pre-treated leaves, even partial
inactivation of APX, a key component of the water–water
cycle, leads to both ROS production and suppressed NPQ.
The source of the detected ROS is probably heterogeneous,
partly originating in the inefficient water–water cycle and
partly in PSII, which is not down-regulated in the absence
of NPQ.
Materials and Methods
Tobacco (Nicotiana tabaccum, L.) and garden pea (Pisum
sativum) plants were grown in the greenhouse at 22–248C with a
natural photoperiod and daytime irradiation maxima around
220–250 mmol m–2 s–1 PAR. With the exception of protein
immunoblots, all experiments were carried out using tobacco
leaves. The youngest fully expanded leaves of 3-week-old tobacco
plants were detached and one half of the leaf was immersed in
water heated up to one of the following temperatures for 3 min: 38,
40, 42, 44, 46 or 488C in dim (2–5 mmol m–2 s–1 PAR) green light.
After this relatively mild heat treatment and drying of the surfaces,
the leaves were used in photosynthesis or ROS measurements.
Non-immersed halves of the leaves served as controls. Leaf stems
were kept wet through all treatments and measurements.
ROS were measured with a newly developed imaging
apparatus capable of detecting changes in the fluorescence of the
ROS sensor 3-(N-dansyl)aminomethyl-2,2,5,5-tetramethyl-2,5dihydro-1H-pyrrole (HO-1889NH; Kálai et al 1998) as described
earlier (Hideg and Schreiber 2007). Briefly, the infiltrating solution
containing 2 mM HO-1889NH in a water : ethanol (99 : 1, v/v)
solution was forced into the middle layer of leaf tissue using a
plastic syringe without a needle, through a pinhole made with a
sharp pin, as described earlier (Hideg et al 2002). Images of HO1889NH fluorescence were taken before and after exposing the leaf
to low intensity (25 mmol m–2 s–1) PAR for 30 min. PAR was
provided from a KL-1500 lamp (DMP, Geneva, Switzerland) via
fiberoptics. ROS production was assessed as lowering of HO1889NH fluorescence during this period. Results are shown as
color-coded images of the ROS sensor fluorescence. Fluorescence
intensities were also averaged from areas of approximately 1–1.5
cm–2 where infiltration was relatively uniform. HO-1889NH
trapping experiments were repeated three times.
Hydrogen peroxide production was visualized with DAB
staining (Thordal-Christensen et al. 1997). In these experiments,
segments were cut from the leaves after the indicated heat pretreatments, then floated on 1 mM DAB solution for 2 h under
25 mmol m–2 s–1 PAR. This was followed by removing chlorophyll
from the leaf cuttings by ethanol (658C, 2 h) to visualize brown
staining characteristic of the presence of H2O2. These experiments
served as illustrations only; H2O2 production was not quantified.
Rates of CO2 assimilation in the leaves were determined by a
portable infrared gas analyser (LI-6400, LI-COR Biosciences,
Lincoln, NE, USA) using reference air with 400–450 mmol mol–1
CO2, corresponding to the growth conditions. To record light
response curves, 6 cm2 areas of untreated or heat-pre-treated leaves
were enclosed in the analyzer’s leaf chamber and irradiated
through a transparent window with a KL-1500 lamp (DMP).
Incident PAR was measured using the gas analyzer’s built-in
sensor. PAR was increased from 0 to 1,000 mmol m–2 s–1, stepwise.
NPQ and ROS imaging in heat-treated leaves
After keeping the leaf at the chosen PAR level for 5 min, net
photosynthesis rates were measured and expressed as rates of CO2
uptake (mmol CO2 m–2 s–1). Experiments were repeated four times,
using four different leaves undergoing the same temperature
exposure before measuring CO2 uptake. Values represent means
of these repetitions with standard deviations.
Chlorophyll fluorescence was imaged at room temperature,
using the MINI-version of the Imaging-PAM (Heinz Walz GmbH,
Effeltrich, Germany), with which areas up to 24 32 mm can be
assessed. This instrument employs the same blue power lightemitting diodes (LEDs) for pulse modulated measuring light,
continuous actinic illumination and saturation pulses. Fo, the
minimal fluorescence yield of dark-acclimated samples, is imaged
at low frequency of pulse-modulated measuring light, while images
of the maximal fluorescence yield, Fm, are obtained with the help of
a saturation pulse. Based on Fo and Fm, the images of maximal,
dark-acclimated PSII quantum yield, Fv/Fm, are derived. With
illuminated samples, the maximal fluorescence yield, Fm0 , is nonphotochemically quenched with respect to Fm (Schreiber et al.
1986). The effective PSII quantum yield of illuminated samples is
calculated from the expression Y(II) ¼ (Fm0 – F)/Fm0 (Genty et al.
1989). Non-photochemical quenching is quantified by the parameter NPQ ¼ (Fm – Fm0 )/Fm0 (Bilger and Björkman 1990) For
assessment of heat-induced changes in photosynthesis, dark–light
induction curves were recorded: first Fo and Fm images were
measured with dark-acclimated samples, from which an Fv/Fm
image was derived. This was followed by 3 min exposure at 75 mmol
m–2 s–1 PAR, with repetitive measurements of Fo and Fm0 images
every 20 s, from which automatically images of Y(II) and NPQ
were calculated by the ImagingWin software. Kinetics of Y(II)
decrease and NPQ build-up were constructed by averaging data
from corresponding areas of interest in the images. The data points
of these induction curves are averages of three repetitions using
new heat-pre-treated leaves, with error bars corresponding to
standard deviations. Results are also shown as color-coded images
of Fv/Fm, Y(II) and NPQ after the 3 min illumination.
Due to the non-invasive nature of PAM fluorescence
measurements, APX could be extracted afterwards from the
same leaves. Following heat treatment and fluorescence measurements, soluble proteins, including enzymes, were extracted from
the tobacco leaves in ice-cold buffer (10 ml g–1 leaf FW) containing
50 mM potassium phosphate (pH 7.0), 1 mM EDTA, 50 mM NaCl,
1% polyvinylpyrrolidone and 1 mM ascorbate. Extracts were
centrifuged at 10 000g for 20 min at 48C and supernatants were
stored at 808C until use. APX activity was measured spectrophotometrically, according to Nakano and Asada (1981), in
50 mM potassium phosphate buffer (pH 7.0) in the presence of
0.5 mM ascorbate and 10 mM H2O2. APX activity was calculated
from the decrease in the 290 nm absorption of ascorbate, using
2.8 mM–1 cm–1 as the molar extinction coefficient. Measurements
were corrected for the direct oxidation of ascorbate by H2O2,
which was 510% of the enzyme-related absorbance change. APX
activity data are given as mM H2O2 consumed per mg total protein
per min; values are averages of four repetitions using new leaves,
and error bars represent standard deviations.
APX protein contents were determined using Western blots.
Due to the reactivity of commercially available antibodies, this
measurement was carried out using garden pea leaves. After
applying the same heat treatments to whole pea leaves as to the
halves of tobacco leaves, the former were weighed and homogenized in 1.5 ml reaction tubes with an equal amount of extraction
buffer [50 mM Tris–HCl, pH 7.2, containing 1 mM phenylmethylsulfonyl fluoride (PMSF)]. The samples were centrifuged at 48C for
1885
45 min at 18,000g to remove the insoluble fraction. The protein
content of the supernatant was determined by Bradford assay
using Bradford Reagent (Sigma-Aldrich Gmbh, Germany) and
bovine serum albumin (BSA) as standard. From each sample, 15 mg
of total soluble protein was separated by 12% SDS–PAGE, then
transferred to a nitrocellulose membrane (G. Kisker GbR, Szeged,
Hungary) and the APX content was probed with anti-APX
antibody (Agrisera, Vännäs, Sweden).
Funding
The Hungarian Research Fund (grant Nos. OTKA
T49438, OTKA-NKTH K67597).
Acknowledgments
We thank Professors Tamás Kálai and Kálmán Hideg (Pécs
University, Department of Organic and Medicinal Chemistry,
Hungary) for the fluorescent ROS sensor HO-1889NH.
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(Received September 5, 2008; Accepted October 31, 2008)