Hydrogen peroxide, nitric oxide and cytosolic ascorbate peroxidase at the crossroad between defence and cell death.

Abstract

An increase in the production of reactive oxygen species (ROS) is a typical event occurring during different stress conditions and activating conflicting responses in plants. In order to investigate the relevance of different timing and amounts of ROS production, tobacco (Nicotiana tabacum) Bright Yellow-2 (TBY-2) cells were incubated with different amounts of glucose plus glucose oxidase, for generating H(2)O(2) during time, or directly with known amounts of H(2)O(2). Data presented here indicate that, in TBY-2 cells, a difference in H(2)O(2) level is a critical point for shifting metabolic responses towards strengthening of antioxidant defences, or their depletion with consequent cell death. Timing of ROS production is also critical because it can determine programmed cell death (PCD) or necrosis. Depending on the different kinds of activated cell death, ascorbate (ASC) and glutathione (GSH) pools are altered differently. Moreover, an H(2)O(2)-dependent activation of nitric oxide synthesis is triggered only in the conditions inducing PCD. Ascorbate peroxidase (APX) has been analysed under different conditions of H(2)O(2) generation. Under a threshold value of H(2)O(2) overproduction, a transient increase in APX occurs, whereas under conditions inducing cell necrosis, the activity of APX decreases in proportion to cell death without any evident alteration in APX gene expression. Under conditions triggering PCD, the suppression of APX involves both gene expression and alteration of the kinetic characteristics of the enzyme. The changes in ASC, GSH and APX are involved in the signalling pathway leading to PCD, probably contributing to guaranteeing the cellular redox conditions required for successful PCD.

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Hydrogen peroxide, nitric oxide and cytosolic ascorbate
peroxidase at the crossroad between defence and cell death
Maria Concetta de Pinto1,†, Annalisa Paradiso1,†, Paola Leonetti2and Laura De Gara1,3,*
1Department of Plant Biology and Pathology, University of Bari, Via Orabona 4, I-70125, Bari, Italy,
2Plant Protection Institute, CNR, Via Amendola 165/A, I-70125 Bari, Italy, and
3Interdisciplinary Center for Biomedical Research (CIR) Universita ` Campus Biomedico Via Longoni 83, I-00155 Roma, Italy
Received 24 May 2006; revised 7 August 2006; accepted 10 August 2006.
*For correspondence (fax þ39 080 544 3553; e-mail degara@botanica.uniba.it).
†These authors contributed equally to this work.
Summary
An increase in the production of reactive oxygen species (ROS) is a typical event occurring during different
stress conditions and activating conflicting responses in plants. In order to investigate the relevance of
different timing and amounts of ROS production, tobacco (Nicotiana tabacum) Bright Yellow-2 (TBY-2) cells
were incubated with different amounts of glucose plus glucose oxidase, for generating H2O2during time, or
directly with known amounts of H2O2. Data presented here indicate that, in TBY-2 cells, a difference in H2O2
level is a critical point for shifting metabolic responses towards strengthening of antioxidant defences, or their
depletion with consequent cell death. Timing of ROS production is also critical because it can determine
programmed cell death (PCD) or necrosis. Depending on the different kinds of activated cell death, ascorbate
(ASC) and glutathione (GSH) pools are altered differently. Moreover, an H2O2-dependent activation of nitric
oxide synthesis is triggered only in the conditions inducing PCD. Ascorbate peroxidase (APX) has been
analysed under different conditions of H2O2generation. Under a threshold value of H2O2overproduction, a
transient increase in APX occurs, whereas under conditions inducing cell necrosis, the activity of APX
decreases in proportion to cell death without any evident alteration in APX gene expression. Under conditions
triggering PCD, the suppression of APX involves both gene expression and alteration of the kinetic
characteristics of the enzyme. The changes in ASC, GSH and APX are involved in the signalling pathway
leading to PCD, probably contributing to guaranteeing the cellular redox conditions required for successful
PCD.
Keywords: ascorbate peroxidase, programmed cell death, necrosis, hydrogen peroxide, nitric oxide, tobacco
Bright Yellow-2 cells.
Introduction
Reactive oxygen species (ROS) are produced by all aerobic
organisms as inevitable by-products of several metabolic
pathways, including electron flows in mitochondria and
chloroplasts, lipid catabolism and photorespiration in gly-
oxysomes and peroxisomes, as well as enzymatic oxyge-
nase reactions having a different cellular localization. In
order to avoid ROS toxicity, aerobic cells are provided with a
flexible set of enzymes and metabolites involved in ROS
catabolism, which often acts at the site of ROS production
(Foyer, 2004; Mittler et al., 2004; Shigeoka et al., 2002).
Despite much metabolic energy being spent on ROS
removal by plant cells, ROS are also actively produced by
cell metabolism under optimal growth conditions. In partic-
ular, hydrogen peroxide is required for biosynthesis and
developmentally related modifications of the structural
components of cell walls (Ros Barcelo ` et al., 2004). An
increasing body of data suggests that ROS also behave as
second messengers. In bacteria, yeasts and animals, altera-
tions in ROS production modify redox-sensitive transcrip-
tion factors, which activate metabolic pathways involved in
ROS-scavenging (Catani et al., 2001; Costa and Moradas-
Ferreira, 2001; Georgiou, 2002). Redox-sensitive regulation
of gene expression has also been described in plants (Mou
et al., 2003), as well as the presence of MAPK pathways and
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some phosphatases, similar to those involved in animal
ROS-dependent signalling (Apel and Hirt, 2004). Eukaryotic
organisms also exploit ROS toxicity for their defence: spe-
cific enzymatic systems producing ROS are activated by
opportune stimuli in both plant and animal cells (Desikan
et al., 1996; Diebold and Bokoch, 2005; Hu et al., 2003;
Walters, 2003).
Inplantcells,ROSoverproductionisinducedbybioticand
abioticstresses.Survivalundertheseconditionsdependson
the capability of plants to increase specific pathways
involved in ROS removal (Asada, 1999; Noctor and Foyer,
1998). Recent studies utilizing knockout and antisense plant
cell lines or organisms further underline the strong link
existing between ROS-removal enzymes and defence re-
sponses against stresses (Mittler et al., 2004). ROS overpro-
duction is also an early event in plant programmed cell
death (PCD; Dat et al., 2003; Levine et al., 1994).
During the activation of PCD, cytosolic ascorbate peroxi-
dase (APX), a pivotal enzyme for ROS removal, strongly
decreases as do ascorbate (ASC) and glutathione (GSH),
metabolites of crucial relevance for cellular redox balance
(de Pinto et al., 2002; Vacca et al., 2004). These changes in
cellular antioxidant capability probably contribute to the
ROS increase that generates the oxidative burst character-
izing PCD. However, the expression/activity of APX is also
regulated downstream by ROS (de Pinto et al., 2002),
suggesting that complicated cause–effect relations occur
between APX–ASC and ROS levels.
It is known that changes in ROS production may activate
contrasting responses: an increase or a decrease in ROS-
scavenging systems, which will guarantee cell survival or
induce cell death, respectively, according to the situation
faced by the plant (Mittler, 2002). Recently, it has been
underlined that metabolic changes triggered in order to
improve cell fitness under specific conditions are the result
of a network of different stimuli, rather than a response to a
unique stimulus (Trewavas, 2005). The signal transduction
pathways are themselves a network of reactions in which
thousands of second messengers, protein kinases and other
molecules generate an information flow that diverges,
adapts, synergizes and integrates in order to optimize the
final response (Taylor and McAinsh, 2004).
It has been reported consistently that PCD requires the
simultaneous presence of hydrogen peroxide and nitric
oxide, both in planta and in cultured cells (Delledonne et al.,
1998; de Pinto et al., 2002). Nevertheless, contrasting data
on the simultaneous requirements of the different reactive
speciesarealsopresentin theliterature.Both intobacco and
Arabidopsis cells, PCD has been activated by single treat-
ments of hydrogen peroxide or nitric oxide (Clarke et al.,
2000; Houot et al., 2001).
Despite several papers underlining the pleiotropic effects
of ROS overproduction, no study has attempted to evaluate
whether ROS generation in different amounts, or with
different rates and times, might provoke different effects
not merely due to a dose-dependent result, but as a
consequence of the activation of diverging metabolic path-
ways.
To obtain further information on the relevance of timing
and intensity of oxidative stress for PCD induction, tobacco
cultured cells were subjected to treatments giving rise to
an increase in hydrogen peroxide during different times
and in different amounts. The results obtained demon-
strate that the metabolic responses activated in cells differ
greatly depending on both the intensity of the oxidative
stress generated and the different timing of ROS produc-
tion.
Results
Hydrogen peroxide and cell death
To study the effects of different amounts and timing of H2O2
production in the responses activated by plant cells, glucose
(G) plus different concentrations of glucose oxidase (GOX)
or different amounts of H2O2were added to the culture
medium of TBY-2 cells.
In the culture medium without cells, the GGOX system
generated H2O2at a constant rate for the first 2–3 h. H2O2
remained at a constant concentration in the culture
medium for the following 2 h, after which it decreased
progressively (data not shown). When GGOX was added to
the TBY-2 cell suspension, the level of H2O2in the culture
medium was always lower than that observed when the
enzymatic system was added to the medium without cells,
probably because a certain amount of ROS was scavenged
by the cells. In cell suspension, the H2O2 reached its
maximum level 15 min after the beginning of the GGOX
treatment and then decreased (Figure 1a,c). After direct
addition of H2O2 for 15 min, the reactive species were
present in the culture medium with concentrations propor-
tional to those added (Figure 1b). At this stage of treat-
ment, the addition of 50 mM H2O2 determined H2O2
concentration in the cell suspension medium comparable
with those obtained by the highest GGOX treatment.
However, under direct H2O2 addition, the decrease in
H2O2over time was much faster than that observed under
GGOX treatment (Figure 1d).
The viability of TBY-2 cells subjected to the different
treatments was analysed (Figure 2). The lowest amount of
GOX (0.75 U ml)1) did not induce cell death, whereas when
higher GOX units were used, dose-dependent cell death was
clearly evident. The highest effect was observed after 6 h
treatment, independently from the amount of GOX used
(Figure 2a). As the GGOX system generates D-gluconolac-
tone as well as H2O2, the dependence of cell death on H2O2
was verified by the addition of catalase (CAT) to the cell
suspension a few minutes before starting the GGOX treat-
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ment. The addition of up to 30 U ml)1CAT was not able to
remove completely the H2O2 produced by GGOX, as a
certain amount of the H2O2produced remained in the cell
suspension (Figure 1a,b). In the presence of CAT, GGOX-
dependent cell death was reduced to about 80% (Figure 2a),
inagreementwiththeinabilityofCATpre-treatmenttoavoid
H2O2generation completely. The direct addition of H2O2to
the cell suspension also induced cell death with a dose-
dependent behaviour (Figure 2b). During the first 6 h of
treatment, 50 mM H2O2induced a decrease of cell viability
comparable with that caused by the highest GOX addition
(14 U ml)1). However, under direct H2O2 addition, the
intensity of cell death increased progressively throughout
the period analysed (Figure 2b). The viability of the H2O2-
treated cells was completely preserved by CAT pre-treat-
ment (Figure 2b), consistent with the very rapid and
complete removal of the added H2O2by CAT pre-treatment
(Figure 1b,d). The addition of glucose to the cell suspension
Figure 1. Presence of H2O2in the culture medium of tobacco BY-2 cells.
H2O2concentration was measured over 24 h in the cell suspension in the absence (control) or after the addition of 14 mM glucose (G) plus 0.75, 2.5, 5, 14 U ml)1
glucose oxidase (GOX) or 5–50 mM H2O2. At the times indicated, 1 ml cell suspension was taken and the concentration of H2O2in the medium determined as
described in Experimental procedures. Values represent mean (?SE) of five experiments.
(a, b)Concentrationsof H2O2after 15 mintreatment with G þ GOX or H2O2, respectively;(c, d)concentrations ofH2O2in the following24 h treatment withG þ GOX
or H2O2, respectively.
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didnotaltertheintensityofcelldeathinducedbydirectH2O2
addition (data not shown).
Some cytological markers were analysed to obtain infor-
mation on which kind of cell death (necrosis or PCD) was
induced by the different treatments. Figure 3 shows that
only the direct addition of H2O2induced cytoplasm shrink-
age, a typical marker of plant PCD, in almost 80–90% of the
Trypan blue-dyed cells (Figure 3c). No cellular shrinkage
was evident in control cells or in those treated with G plus
14 U ml)1
GOX (Figure 3a,b). DAPI-staining cells also
showed the presence of chromatin condensation and apop-
totic-like nuclei (Houot et al., 2001) only in the cells directly
treated with H2O2(Figure 3f); whereas, nuclei from control
and GGOX-treated cells had normal morphology (Fig-
ure 3d,e). Moreover, DNA laddering was evident in the cells
directly treated with H2O2but not in those subjected to
GGOX treatment (Figure 4).
Nitric oxide production
As the simultaneous production of H2O2and NO appears to
be required for the activation of PCD in TBY-2 cells (de Pinto
et al., 2002), the capability of the two different treatments
(H2O2generation by G plus 14 U ml)1GOX versus direct
50 mM H2O2addition) to induce NO production was inves-
tigated. The analysis of nitric oxide synthase (NOS) activity
indicated that the enzyme was active only in cells treated
with H2O2and not in those treated with GGOX (Figure 5a).
Production of NO was also quantified spectrofluorimetri-
cally by means of the NO-specific probe DAF-2DA (Foissner
et al., 2000). The results obtained clearly indicate that this
reactive species was produced under H2O2treatment but not
under GGOX (Figure 5b). The increase in fluorescence
occurring in the H2O2-treated cells was specifically due to
NO, as it was strongly reduced by the NO scavenger cPTIO
(a) (b) (c)
(f) (e) (d)
Figure 3. Cytological analysis
H2O2-treated cells.
Cell suspensions were treated with 14 mM glu-
cose (G) plus 14 U ml)1glucose oxidase (GOX)
or 50 mM H2O2. Aliquots of control and treated
cells were collected, stained with trypan blue or
DAPI dyes, and visualized under phase-contrast
microscopy (a–c) or fluorescence microscopy (d–
f). Pictures represent typical examples after 6 h
treatment: (a, d) control cells; (b, e) G þ GOX-
treated cells; (c, f) H2O2-treated cells. Bar, 20 lm.
Arrows and arrowheads indicate apoptotic-like
nuclei and chromatin condensation, respect-
ively.
of GGOX- or
Figure 2. Time-dependent effect of H2O2production on cell viability.
Viability of control and treated cells was measured as reported in Experimental procedures.
(a) Glucose plus 0.75, 2.5, 5, 14 U ml)1glucose oxidase; (b) 5–50 mM H2O2. Where indicated, catalase (30 U ml)1) was added to the culture medium 10 min before
treatment. The percentage (?SE) of viable cells was counted in a population of at least 1000 cells in five independent experiments.
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(Figure 5b). Interestingly, the addition of cPTIO to the H2O2-
treated cells was also able to prevent PCD (Figure 5c). The
capability of H2O2to induce NO production was also evident
when treated cells were observed under a florescent light
microscope. At the beginning of H2O2treatment all cells
produced NO, whereas after 2 h only the cells that were still
alive showed a clear NO-dependent green fluorescence
(Figure 6).
Cellular antioxidants and cytosolic ascorbate peroxidase
To analyse the cellular redox state determined by different
treatments, the ASC and GSH pools (reduced plus oxidized
forms)weredeterminedduringthefirst6 hoftreatment.The
ASC pool decreased significantly both under GGOX treat-
ment, inducing necrosis, and under H2O2addition, inducing
PCD. However, the decrease was much more severe in cells
undergoing PCD (direct addition of H2O2), with an 80% de-
crease in the first 2 h versus a 25% decrease under GGOX
treatment (Figure 7a). The two treatments caused still more
different effects on GSH. The continuous H2O2production
(GGOX treatments) induced an increase in the GSH pool,
whereas direct addition of H2O2caused a strong decrease in
the GSH pool (about 75% during the first 4 h; Figure 7b).
Nevertheless, direct addition of H2O2and GGOX treatment
induced similar decreases in the ASC and GSH redox state
(ratio between reduced and reduced plus oxidized forms).
During the first 2 h of treatment the ASC redox state de-
creased from 0.87 to 0.62 and 0.56 in GGOX- and H2O2-trea-
ted cells, respectively, and the GSH redox state from 0.98 to
0.83 and 0.81 in GGOX- and H2O2-treated cells, respectively.
The behaviour of cytosolic APX, one of the main ROS-
scavenging enzymes in plant cells, was analysed during the
different treatments. APX activity was increased transiently
when GOX was added at the lowest concentration
(0.75 U ml)1; Figure 8a). This increase was small but statis-
Figure 4. DNA analysis.
DNA was extracted from cells collected 24 h after treatment with 14 mM
glucose plus 14 U glucose oxidase or 50 mM H2O2. A representative
electrophoretic run in which 100 lg DNA was loaded in each line.
Figure 5. Nitric oxide production by tobacco BY-2 cells in response to glucose þ glucose oxidase (G þ GOX) or H2O2treatments and its effect on cell viability.
(a) Nitric oxide synthase: enzyme activity was measured in control cells and in cells treated with 50 mM H2O2or 14 mM G plus 14 U ml)1GOX 2, 4 and 6 h after
treatments. Values represent mean ? SE of five experiments.
(b) Nitric oxide production: fluorescence emission of TBY-2 cells was measured at the times indicated in cells loaded with DAF-2DA then treated with 14 mM G plus
14 U ml)1GOX or 50 mM H2O2, and control cells (loaded with DAF-2DA without any subsequent treatments). Where indicated, cell suspensions were pre-incubated
for 10 min with 500 lM carboxy-PTIO, an NO-scavenger, before treatment. Results are the mean of three independent experiments; variation in values obtained for
the same treatments was in the range 5–10%.
(c)CellviabilityofH2O2-treated cellsintheabsenceorpresence of500 lM carboxy-PTIO.Percentage(?SE) ofviablecellswascountedinapopulationofatleast1000
cells in three independent experiments.
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tically significant after 2 and 4 h treatment (Student’s t-test,
P < 0.01). On the other hand, higher amounts of GOX
determined a dose-dependent inhibition of APX activity.
Such APX inhibition induced by GGOX was attenuated over
time, as the values of APX activity after 24 h of GGOX
treatment were higher than those observed at 6 h (Fig-
ure 8a).
Figure 6. NO accumulation induced by H2O2in
tobacco BY-2 cells.
Cells were pre-loaded with DAF-2DA and treated
with 50 mM H2O2. At the times indicated, 1 ml
cell suspension was treated with 0.4% trypan
blue and examined by microscope. Representa-
tive phase-contrast and fluorescence images of
H2O2-treated cells are reported in upper and
lower panels, respectively. Bar, 20 lm.
Figure 7. Changes inascorbate(ASC)andglutathione(GSH)contentinduced
by glucose þ glucose oxidase (G þ GOX) and H2O2treatments.
Control and treated cells (14 mM G plus 14 U ml)1GOX or 50 mM H2O2) were
collectedatthetimesindicatedandusedforthedeterminationofthetotalASC
(a)orGSH(b)pools(reducedplusoxidizedforms)asreportedinExperimental
procedures. Values represent means (?SE) of three experiments.
(a)
(b)
Figure 8. Effects of glucose þ glucose oxidase (G þ GOX) or H2O2 treat-
ments on ascorbate peroxidase (APX) activity.
(a) Specific activity of APX was measured in control and treated cells (14 mM
G plus 0.75, 2.5, 5, 14 U ml)1GOX or 50 mM H2O2) during time. Values
represent means (?SE) of five experiments.
(b) Representative native PAGE of APX from tobacco BY-2 cells treated for 2 h
with 14 mM G plus 0.75 U ml)1GOX, 14 U ml)1GOX or 50 mM H2O2. Where
indicated, cell suspensions were pre-treated with catalase (30 U ml)1). For
each treatment, 300 lg total protein was loaded.
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The direct H2O2addition induced a dose-dependent APX
inhibition (with an APX activity of 1800 ? 66, 660 ? 25,
280 ? 33, 155 ? 18 at 2 h treatment with 5, 15, 30 and 50 mM
H2O2, respectively). As shown in Figure 8a, the addition of
50 mM H2O2 to the culture medium induced a prompt
decrease in enzyme activity. However, under this treatment
no recovery in APX activity was observed with time
(Figure 8a). These results were also confirmed by electroph-
oretic analyses. Figure 8b shows a representative native
PAGE of APX activity after 2 h treatment. Both cytosolic
isoforms of APX were increased slightly under treatment
with 0.75 U GOX. In contrast,the two bands were attenuated
under 14 U GOX treatment and disappeared completely
under treatment with 50 mM H2O2. CAT pre-treatment made
H2O2treatment completely inefficient at inhibiting APX. It
also remarkably reduced the inhibition of APX due to GGOX
(Figure 8b).
To verify whether the decrease in enzyme activity was
also due to changes in the kinetic characteristics of APX,
the dependence of its reaction rate on increasing ascor-
bate concentration, in the presence of a fixed concentra-
tion of hydrogen peroxide, was analysed. The kinetic
parameters of APX were investigated in both control cells
and cells treated for 15 min with GGOX (14 U ml)1GOX)
or H2O2(50 mM). As shown in Figure 9, the enzyme Vmax
of cells treated with H2O2and GOX was lower than that of
control cells. However, APX from both GGOX-treated and
control cells showed a sigmoidal dependence of rate on
ASC concentration. This means that the enzyme from
control and GGOX-treated cells showed a co-operative
pattern, having a Hill coefficient of 1.7 and 1.5, in control
and GGOX-treated cells, respectively. This is consistent
with the presence of two binding sites for ASC (Lad et al.,
2002). On the other hand, in the H2O2-treated cells, APX
lost this co-operative pattern, showing a hyperbolic
dependence of rate on ASC concentration with a Hill
coefficient close to 1 (Figure 9).
Levels of APX protein were measured by immunoblotting
using a specific monoclonal antibody (Figure 10). In cells
treated with the lowest concentration of GOX (0.75 U ml)1)
theamountofAPXproteinwasalwayshigherthanincontrol
cells. A significant decrease in the level of APX protein was
evident only after 6 h cell treatment with the highest
concentration of GOX (14 U ml)1). On the other hand, direct
addition of H2O2induced an immediate decrease in the
amount of APX protein that was completely prevented by
addition of CAT to the culture medium.
Finally, the expression of cytosolic APX (cAPX) was
investigated in cells treated with H2O2and GGOX. Gene
expression was analysed by semi-quantitative RT-PCR using
specific primers for tobacco APX. As shown in Figure 11,
addition of H2O2 to the culture medium determined a
remarkable decrease in APX gene expression, which was
progressively more evident during the treatment.
Figure 9. Kinetic analysis of cytosolic ascorbate peroxidase (cAPX) in cells treated with glucose þ glucose oxidase (G þ GOX) or H2O2.
The analysis was performed in extracts from control cells and cells treated for 15 min with 14 mM G plus 14 U ml)1GOX or 50 mM H2O2. Concentration of H2O2was
170 lM; concentration of ascorbate (ASC) varied over the range 10–700 lM. n ¼ Hill coefficient. Vmaxand Kmvalues for ASC in control and treated cells represent the
mean (?SE) of five experiments.
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Discussion
Cell death and nitric oxide requirement for programmed cell
death
The results reported here indicate that production of H2O2
over a threshold value induces cell death in TBY2 cells
(Figure 2). However, the kind of death (necrotic versus PCD)
depends strongly on the timing of H2O2production. The
prolonged production of this ROS for several hours induces
a necrotic process as, under these conditions, no cytological
PCD hallmarks are evident. On the other hand, an amount of
H2O2comparable with that inducing cell necrosis, but given
as a single pulse, triggers PCD (Figures 3 and 4). Direct H2O2
addition probably determines a situation similar to the oxi-
dative burst induced by several biotic or abiotic stresses
against which PCD is frequently activated (Overmyer et al.,
2003). However, not all the signalling pathways leading to
PCD require the generation of an oxidative burst and, under
certain situations, ROS production even seems to suppress
cell death (Torres et al., 2005). It has been reported that the
mesophyll cells of Zinnia elegans, when in vitro trans-dif-
ferentiating into tracheids, require a constant oxidative state
for PCD induction(Gomez-Ros et al., 2006).In this processof
tracheid differentiation, NO production is another crucial
event during PCD (Gabaldon et al., 2005). One noteworthy
result reported here is that only the direct addition of H2O2
activates NO synthesis, whereas the constant production of
H2O2over time has no effect on NO synthesis, when either
NOS activity or NO production is analysed (Figure 5). The
capability of the NO-scavenger cPTIO in restoring cell viab-
ility under direct addition of H2O2confirms the requirement
of the two reactive species in the signalling pathway leading
to PCD. The activation of NO biosynthesis by H2O2, probably
by a NOS-like enzyme, has also been reported in mung bean
leaves (Lum et al., 2002). Moreover, it is known that treat-
ment with NO generators is able to make ROS effects vanish
or strengthen (Delledonne et al., 2001; Orozco-Ca ´rdenas and
Ryan, 2002), further supporting the view that signalling
pathways, based on the same chemical messengers, inter-
play differently according to the different environmental or
cellular contexts. Increasing evidence suggests that NO and
ROS play an active role in regulating metabolic changes
occurring both during plant development and in responses
against stress (Foyer and Noctor, 2005; Lamattina et al.,
2003); our results support the hypothesis that, at least in
TBY-2 cells, H2O2and NO are team players in the PCD-sig-
nalling pathway when produced in a precise amount and
with precise timing.
Antioxidant involvement in cell death induced by H2O2
The metabolism of ASC and GSH plays a pivotal role in ROS
removal in plants (Mittler, 2002; Noctor and Foyer, 1998;
Shigeokaet al.,2002).IthasalsobeenreportedthatAPXand
ASC decrease remarkably as early events of the PCD process
(de Pinto et al., 2002;Vaccaet al.,2004;Figures 7 and 8). The
decrease in this antioxidant defence line has been consid-
ered part of the strategy contributing to generating the oxi-
dative burst that characterizes the PCD triggered by several
stimuli (De Gara, 2004; De Gara et al., 2003; Mittler et al.,
1998). Data reported here suggest that ASC and GSH levels,
more than their redox state, are altered as a specific signal in
cells undergoing PCD (Figure 7), as the alterations in ASC
and GSH redox state occurring under treatmentswith GGOX
or H2O2direct addition are similar. Consistent with this,
Figure 11. Levels of ascorbate peroxidase (APX) mRNA in cells treated with
glucose þ glucose oxidase (G þ GOX) or H2O2.
Total RNA was extracted at the times indicated from control cells and cells
treated with 14 mM G plus 14 U ml)1GOX or 50 mM H2O2. Semi-quantitative
RT-PCR for APX was performed as described in Experimental procedures.
Levelsof 18SrRNAs werealso determinedso thatresults could benormalized
for recovery of RNA from the initial samples.
Figure 10. Changes in the level of ascorbate peroxidase (APX) protein under
glucose þ glucose oxidase (G þ GOX) or H2O2treatment.
Immunoblotting analysis was performed using a specific APX antibody (see
Experimental procedures) in control cells and in cells treated with 14 mM G
plus 0.75 and 14 U ml)1GOX or 50 mM H2O2. Where indicated, catalase
(30 U ml)1) was added to the culture medium 10 min before treatment; 5 lg
totalproteinwasloadedineach line.Afterdensitometry,levelsofAPXprotein
intreatedcellswerecomparedwiththecontrolandcalculatedasapercentage
of alteration.Values representmean (?SE) ofthree independent experiments.
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Arabidopsis thaliana mutants, having a 10–25% wild-type
ascorbate content but a normal ASC redox state, spontane-
ously activate processes that typically occur during hyper-
sensitive PCD, such as localized cell death and the
expression of pathogenesis-related proteins (Pavet et al.,
2005).
Our results also indicate that the level and timing of H2O2
production are critical points for APX behaviour. The con-
stant production of low amounts of this reactive species,
which was ineffective in inducing cell death (0.75 U ml)1
GGOX), determines a transient, modest rise in APX (Fig-
ures 8 and 10). Such a rise is aimed at restoring redox
impairment due to ROS overproduction, occurring under
moderate GGOX treatment. The strengthening of antioxid-
ant metabolism has been reported widely in the literature as
a first line of defence against moderate oxidative stress
(Asada, 1999; Mittler, 2002; Noctor and Foyer, 1998; Shi-
geoka et al., 2002).
On the other hand, when the continuous production of
H2O2increases, a dose-dependentdecrease in APX occurs at
the level of enzyme activity and, over longer times, at the
levelof theamount of protein (Figures 8and10). Even inthis
case, APX inhibition is transient: 6 h after GGOX addition its
activity progressively increases again, probably because the
cells that are still alive become able to activate their
strategies for counteracting the residual oxidative stress.
The H2O2-dependent decrease in APX activity is not a
surprise. Although H2O2is one of the physiological sub-
strates of the enzyme, its presence over a threshold value
directly inhibits its activity by causing protein oxidation
(Hiner et al., 2000). No information is available, at least to
our knowledge, on the possibility that protein oxidation
accelerates APX turnover, which might explain the decrease
in the amount of protein, evident from Western blotting
analysis, in the absence of a decrease in its gene expression
(Figures 10 and 11). However, the site-specific oxidation of
proteins has been reported to act as a ubiquitination signal
triggering protein degradation (Iwai et al., 1998).
Ascorbate peroxidase behaves quite differently in cells
undergoing PCD due to direct addition of H2O2. In this case,
the decrease in enzyme activity appears to involve mecha-
nisms different from simple protein oxidation. In fact, a
change in the kinetic behaviour of the enzyme (from
sigmoidal to hyperbolic) occurs only under these conditions
(Figure 9). A stimulating hypothesis is that the different APX
behaviour is a consequence of NO generation. In animal
cellstheformationofS-nitrosothiolgroups,ferrousnitrosyl–
haem complexes and the nitration of specific tyrosine
residues are all well-characterized parts of the signalling
pathwaystriggeredbyNO(Hofmannet al.,2000).Aguanylyl
cyclase isoenzyme of smooth muscle is activated by the NO
binding to the haeme group located in its regulatory
domain, and the consequent increase in cGMP concentra-
tion activates a cGMP-dependent protein kinase (Hofmann
et al., 2000; Lucas et al., 2000). Moreover, NO trans-nitrosy-
lation and oxidation of specific cysteine residues has been
reported to induce conformational changes in trans-mem-
brane calcium channel proteins in neuronal cells (Nelson
et al., 2003). S-nitrosylation has also been suggested to play
a central role in the signalling pathways leading to disease
resistance in plants (Feechan et al., 2005). The fact that APX
is a haemoprotein and has a cysteine near its catalytic site
(Lad et al., 2002) makes it a putative NO target. An in vitro
inhibition of APX by NO generators, probably due to the
formation of a nitrosoferrous [Fe(II)NOþ] complex (Ferrer
and Ros Barcelo ´, 1999), has been reported previously (Clark
et al., 2000); even if the in vivo effect of NO seems to be
more complex and strongly dependent on its concentration
as well as the NO donor used and/or the presence of other
reactive species (Murgia et al., 2004a; de Pinto et al., 2002;
M.C.P. and L.D.G., unpublished data). It is worth noting that,
inthepresenceofNO,GSHcanbetransformedintoGSNO,a
powerful nitrosylation agent (Ji et al., 1999). GSNO forma-
tion might contribute to the decrease in the GSH pool
(Figure 7), also promoting APX nitrosylation, even if further
analysis is required in order to confirm this hypothesis.
Nitric oxide or, more properly, the cellular environment
determined by the simultaneous presence of NO and H2O2,
also appears to affect APX expression (Figure 11). Genera-
tion of NO in Arabidopsis plants induces a decrease in the
thylakoidal APX transcript accumulation; consistently, Ara-
bidopsis plants over- or underexpressing one thylakoidal
APX gene show increased or decreased sensitivity, respect-
ively, to both NO-induced cell death and paraquat-induced
oxidative stress (Murgia et al., 2004b; Tarantino et al., 2005).
ChangesinAPX expressioncouldalsobecorrelatedto the
observed decrease in ASC content: in several experimental
systems, a tight correlation has been reported between
changes in ASC and APX (Arrigoni, 1994; De Gara et al.,
1997). The strong decrease in ASC could be, at least in part,
due to an impairment of L-galactone-c-lactone dehydroge-
nase, the last enzyme of ASC biosynthesis (Smirnoff et al.,
2001). Accordingly, a significant decrease in the activity of
this enzyme has been observed in TBY-2 cells undergoing
heat shock-induced PCD (M.C.P. and L.D.G., unpublished
data), even if no information is yet available on a putative
role for NO in such an alteration. The different behaviour of
ASC and GSH under necrotic or PCD conditions further
underlines that, despite oxidative stress occurring in both
situations, the molecules and enzymes involved in the fine
regulation of the cellular redox balance are modulated
diversely, according to the different fates for which the cells
are programmed.
Inconclusion,thedata reportedheresuggestthatnotonly
the different kinds of reactive species generated under
environmental or developmental stimuli, but also the
amount and timing of their generation can give reasons for
the different responses triggered in plant cells. In this
792 Maria Concetta de Pinto et al.
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picture, the interplay between nitric and oxygen reactive
species plays a pivotal role in plant fitness.
Experimental procedures
Cell culture and treatments
The tobacco BY-2 cell suspension (Nicotiana tabacum L. cv. Bright
Yellow-2) was routinely propagated and cultured according to
Nagata et al. (1992). For the experiments, a stationary culture
(7 days) was diluted 5/100 (v/v). At the third day of culture, 14 mM
glucose plus glucose oxidase (Calbiochem, La Jolla, CA, USA) 0.75–
14 U ml)1or 5–50 mM H2O2was added to 30 ml cell suspension.
Where indicated, catalase (30 U ml)1) was added to the culture
medium 10 min before treatment. At the times indicated, aliquots of
cells were collected by filtration on Whatman 3MM paper for ana-
lysis.
Cell viability and nuclear morphology
Cell viability was measured as described previously using Trypan
Blue staining (de Pinto et al., 1999). For the analysis of nuclear
morphology, TBY-2 cells were stained with 4,6-diamidino-2-phen-
ylindole (DAPI), as described by de Pinto et al. (2002), and visualized
using a fluorescence microscope (DMLS, Leica, Wetzlar, Germany)
with an excitation filter of 340 nm and a barrier filter of 400 nm.
DNA laddering
Cells were collected from cell suspension and homogenized in li-
quid nitrogen. DNA was extracted by the cetyl trimethyl ammonium
bromide (CTAB) method (Vacca et al., 2004). DNAse-free RNAse
was added to a final concentration of 1 mg ml)1; DNA fragments
were separated by electrophoresis on 1.8% (w/v) agarose gel, fol-
lowed by visualization by staining with ethidium bromide.
Measurement of H2O2
H2O2was measured according to Bellincampi et al. (2000) in the
extracellular phase. Briefly, 1 ml cell culture was harvested by
centrifugation (10 000 g, 20 sec, 25?C) and the H2O2concentration
was measured in the supernatant. An aliquot of supernatant (500 ll)
was added to 500 ll assay reagent (500 lM ferrous ammonium
sulphate, 50 mM H2SO4, 200 lM xylenol orange, 200 mM sorbitol).
After 45 min incubation, the peroxide-mediated oxidation of Fe2þto
Fe3þwasdetermined by measuring the absorbance at 560 nmof the
Fe3þ–xylenol orange complex.
Measurements of nitric oxide production
Cell cultures were treated with 20 lM DAF-2DA for 1 h, then rinsed
and re-suspended in a fresh medium. Cells were then subjected to
the treatments and their fluorescence monitored at regular inter-
vals. To verify whether increases in green epifluorescence could be
attributed to NO evolution, DAF-2DA-loaded cells were incubated
for 10 min in 500 lM carboxy-PTIO, an NO-scavenger, before treat-
ment. Fluorescence intensity was measured using a Shimadzu RF-
1501 luminescence spectrophotometer at 495 nm excitation and
515 nm emission. Production of NO in the cells was observed by
fluorescence microscopy on a Leica DMLS fluorescence microscope
mountedwithanI3filter,at470 nmexcitationand515 nmemission.
Nitric oxide synthase activity
Activity of NO synthase in tobacco cells was measured by monit-
oring the conversion of L-[U-14C]arginine into L-[14C]citrulline (Rees
et al., 1995). In brief, 300 mg cells was ground to a powder under
liquid N and homogenized in ice-cold homogenization buffer
(250 mM Tris–HCl pH 7.4, 10 mM EDTA, 10 mM EGTA). The homo-
genates were centrifuged at 10 000 g for 20 min at 4?C and the
supernatant was used for analysis. Activity of NOS was determined
using the Nitric Oxide Synthase Assay Kit (Calbiochem) according
to the supplier’s recommendations. Briefly, 1 ll radioactive arginine
(approximately 64 Ci mmol)1, 1 lCi ll)1, Amersham Biosciences
Europe GMBH, Milan, Italy), was added to 9 ll cellular extracts
containing 1 lg ll)1protein and incubated for 30 min at 30?C with
40 ll mixture reaction buffer (50 mM Tris–HCl pH 7.4, 6 lM tetra-
hydrobiopterin, 2 lm flavin adenin dinucleotide, 2 lM flavin mono-
nucleotide, 10 mM NADPH, 6 mM CaCl2). The reactions were stopped
with 400 ll stop buffer (50 mM HEPES pH 5.5 with 5 mM EDTA).
Successively, 100 ll equilibrated resin, which binds arginine, was
added to the samples and then transferred to the spin cups. The
citrulline, being ionically neutral at pH 5.5, passes completely
through the column. The NOS activity was then determined by
quantifying the radioactivity in the eluate by liquid-scintillation
counting. When extracts were boiled for 10 min, no significant
conversion into L-[14C]citrulline was detected. For the negative
controls, 5 ll of the NOS inhibitor, NG-nitro-L-arginine methyl ester,
HCl were added. Rat cerebellum was used for the positive control.
Ascorbate and glutathione assay
Cells (0.5 g) were collected by filtration on Whatman 3MM paper
and homogenized with 2 vol cold 5% (w/v) metaphosphoric acid at
4?C in a porcelain mortar. The homogenate was centrifuged at
20 000 g for 15 min at 4?C, and the supernatant was collected for
analysis. Ascorbate and glutathione contents were measured as
described by de Pinto et al. (1999).
Activity and kinetics of ascorbate peroxidase
Cells were ground in liquid N and homogenized at 4?C in extraction
buffer [50 mM Tris–HCl pH 7.8, 0.05% (w/v) cysteine, 0.1% (w/v)
BSA]. The homogenate was centrifuged at 20 000 g for 15 min.
The supernatant was used for both spectrophotometric and elec-
trophoretic analyses.
The reaction catalysed by APX (L-ascorbate: hydrogen peroxide
oxidoreductase, EC 1.11.1.11, cytosolic isoenzyme) was determined
by following the H2O2-dependent oxidation of ASC as decrease
in absorbance at 290 nm, as described by Vacca et al. (2004).
The values for Vmaxand Kmwere obtained by either Michaelis–
Menten or Hill analysis using SIGMAPLOT software (SPSS Inc.,
Chicago, IL, USA). Native PAGE of APX was performed according
to de Pinto et al. (2000).
Protein determination
Assay of proteins was carried out according to Bradford (1976).
Immunoblot analysis
Cells were ground in liquid N and homogenized at 4?C in extraction
buffer [50 mM Tris–HCl pH 7.5, 0.05 % (w/v) cysteine, 0.1 % (w/v)
BSA]. The homogenate was centrifuged at 20 000 g for 15 min, and
ROS, NO and APX between defence and cell death 793
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thesupernatantwaselectrophoresedon12.5%SDS-polyacrylamide
gel under reduction conditions. Immunoblotting was carried out
using anti-cAPX monoclonal antibody (AP6 from Saji et al., 1990)
and with goat anti-mouse immunoglobulin G conjugated with
alkaline phosphatase (Promega, Madison, WI, USA) as described
previously by de Pinto et al. (2002). Densitometry values for im-
munoreactive bands were quantified using a GS-700 imaging den-
sitometer (Bio-Rad, Milan, Italy) as described by Vacca et al. (2004).
Total RNA extraction and semi-quantitative RT-PCR
Total RNA was isolated from TBY-2 cells using the RNeasy plant
minikit (Quiagen, Milan, Italy) according to the supplier’s recom-
mendations. Residual DNA was removed from the RNA samples
using a DNA-free kit (Ambion, Austin, TX, USA). Synthesis of cDNA
was performed from 2 lg total RNA with 10 lM random primers
(Amersham), utilizing an Omniscript reverse transcriptase kit (Qui-
agen) according to the supplier’s recommendations. PCR reactions
were performed with specific primers for cAPX (cAPX, D85912) and
18S rRNA (18S, AJ236016) as described by Vacca et al. (2004). 18S
rRNA was used as an internal control to normalize each sample for
variations in the amount of initial RNA. For semi-quantitative RT-
PCR, the cycle number in the linear range was determined empi-
rically. The products of PCR amplification produced a single band at
the predicted sizes of 699 and 594 bp for cAPX and 18S, respect-
ively. These were analysed on 1.5% agarose gel containing
0.5 lg ml)1ethidium bromide.
Acknowledgements
The authors wish to thank Dr Akihiro Kubo (Environmental Biology
Division, National Institute for Environmental Studies, Onogawa,
Japan) for kindly supplying ascorbate peroxidase antibody, and
Massimo Delledonne for suggestions given during the experimen-
tal work and the stimulating discussion on nitric oxide. This work
was supported financially by the Italian Ministry of Instruction,
University and Research (MIUR; PRIN 2004 No. 2004052535).
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