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Plant Physiol. (1998) 118: 1067-1078
Localized Changes in Peroxidase Activity Accompany Hydrogen
Peroxide Generation during the Development of a Nonhost Hypersensitive
Reaction in Lettuce1
Charles S. Bestwick*, 2,
Ian R. Brown, and
John W. Mansfield
Department of Biological Sciences, Wye College, University of
London, Wye, Kent TN25 5AH, United Kingdom
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ABSTRACT |
Peroxidase
activity was characterized in lettuce (Lactuca sativa
L.) leaf tissue. Changes in the activity and distribution of the enzyme
were examined during the development of a nonhost hypersensitive
reaction (HR) induced by Pseudomonas syringae (P. s.) pv phaseolicola and in response to an
hrp mutant of the bacterium. Assays of activity in
tissue extracts revealed pH optima of 4.5, 6.0, 5.5 to 6.0, and 6.0 to
6.5 for the substrates tetramethylbenzidine, guaiacol, caffeic acid,
and chlorogenic acid, respectively. Inoculation with water or with
wild-type or hrp mutant strains of P. s.
pv phaseolicola caused an initial decline in total
peroxidase activity; subsequent increases depended on the hydrogen
donor used in the assay. Guaiacol peroxidase recovered more rapidly in
tissues undergoing the HR, whereas changes in tetramethylbenzidine
peroxidase were generally similar in the two interactions. In contrast,
increases in chlorogenic acid peroxidase were significantly higher in
tissues inoculated with the hrp mutant. During the HR,
increased levels of Mn2+/2,4-dichlorophenol-stimulated NADH
and NADPH oxidase activities, characteristic of certain peroxidases,
were found in intercellular fluids and closely matched the accumulation
of H2O2 in the apoplast. Histochemical analysis
of peroxidase distribution by electron microscopy revealed a striking,
highly localized increase in activity within the endomembrane system
and cell wall at the sites of bacterial attachment. However, no clear
differences in peroxidase location were observed in tissue challenged
by the wild-type strain or the hrp mutant. Our results
highlight the significance of the subcellular control of oxidative
reactions leading to the generation of reactive oxygen species, cell
wall alterations, and the HR.
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INTRODUCTION |
ROS, a collective term for radicals and other nonradical but
reactive species derived from oxygen, have been implicated in numerous
developmental and adaptive responses in both animal and plant cells
(Dypbukt et al., 1994 ; de Marco and Roubelakis-Angelakis, 1996; Lamb
and Dixon, 1997). In plants, the increased production of both the
superoxide radical and H2O2
is a common feature of defense responses to challenge by microbial
pathogens and elicitors (Lamb and Dixon, 1997). It has been proposed
that a rapid increase in either intra- or extracellular
H2O2 is involved in the
induction and/or execution of the HR (Levine et al., 1994; Low and
Merida, 1996; Bestwick et al., 1997).
The HR represents the rapid and localized death of plant cells in
response to challenge with an avirulent pathogen and is observed in
most examples of race-specific resistance and in many examples of
nonhost resistance (Mansfield et al., 1997). ROS may induce lipid
peroxidation, which has been detected as an early event in some HRs and
may also damage DNA directly and modify or inactivate proteins, leading
to a loss of cell viability (Ádám et al., 1989, 1995; Croft
et al., 1990). Of potential importance is the reaction of
H2O2 with
Fe2+/Cu+ to form the highly
reactive hydroxyl radical (Halliwell and Gutteridge, 1989). In addition
to direct participation in the destruction of the host cell, ROS
themselves or the direct products of their action may also induce more
complex programmed cell death pathways in plants. Some investigators
have described the HR as resembling the process of apoptosis, the
principal manifestation of programmed cell death in many animal
cell types (Dangl et al., 1996; Morel and Dangl, 1997).
There is considerable evidence that
H2O2 has a wider role in
resistance reactions, since it is required for cross-linking plant cell
wall components as part of structural defense reactions and may also
regulate gene expression associated with antioxidant defenses,
phytoalexin biosynthesis, and the development of systemic acquired
resistance (Lamb and Dixon, 1997). The efficacy and scope of
H2O2 action is dependent on
the intensity and longevity of its production. Plasma membrane-bound,
neutrophil-like NADPH oxidase activity has been implicated in
H2O2 production in many
plant species (Low and Merida, 1996; Lamb and Dixon, 1997). However, peroxidase activity, which is an important component of plant stress
responses, may also regulate the level of ROS (Wojtaszek, 1997).
Plant peroxidases are monomeric, heme-containing proteins that are
usually glycosylated. Isoperoxidases, arising from the transcription of
different genes or from posttranslational modification, are widely
distributed within both the intra- and extracellular environment
(Rathmell and Sequeira, 1974; Campa, 1991; Ievenish, 1992; Jackson and
Ricardo, 1994). Peroxidases are active in the H2O2-dependent
polymerization of hydroxycinnamyl alcohols (monolignols) during the
final stages of lignin biosynthesis (Monties, 1989). Increases in
peroxidase activity during incompatible plant-pathogen/elicitor interactions are often associated with a progressive incorporation of
phenolic compounds within the cell wall (Fink et al., 1991; Graham and Graham, 1991; Reimers et al., 1992; Milosevic and
Slusarenko, 1996). Peroxidase also catalyzes rapid,
H2O2-dependent
cross-linking of cell wall proteins such as the Hyp-rich glycoproteins
and Pro-rich proteins, as well as cross-links between other wall
components (Iiyama et al., 1994). The reinforcement of the wall reduces
susceptibility to wall-degrading enzymes, possibly restricts diffusion
of pathogen-derived toxins to the host, and, in the case of some fungal
pathogens, acts as a mechanical barrier to physical penetration toward
the protoplast (Aist and Gold, 1987; Brisson et al., 1994).
Peroxidase activity might be expected to reduce the level of ROS by
metabolizing H2O2, but
peroxidase is also capable of various "oxidase" reactions leading
to H2O2 generation. For
example, the oxidation of IAA, NADPH, NADH, certain phenols, and thiols
in vitro has been shown to produce
H2O2 (Pedreño et al.,
1990; Vianello and Macri, 1991; Pichorner et al., 1992; Jiang and
Miles, 1993). Both intra- and extracellular peroxidases may be involved
in these reactions (Vianello and Macri, 1991). However, to our
knowledge, the nature of the in vivo reductant for peroxidase-catalyzed
generation of H2O2 has not
yet been identified (Bolwell et al., 1995; Wojtaszek, 1997).
Here we examine changes in activity of peroxidase with reference to
phenol oxidation and H2O2
generation in the apoplasm and symplasm of lettuce (Lactuca
sativa L.) leaves challenged with wild-type and nonpathogenic
hrp mutant strains of Pseudomonas syringae
(P. s.) pv phaseolicola. Both strains caused
localized wall alterations and the formation of paramural deposits or
papillae that contain peroxidase substrates such as Hyp-rich
glycoproteins and phenolic compounds, but only the wild type induced a
confluent HR (Bestwick et al., 1995, 1997). By comparative studies it
is therefore possible to identify responses specific to the HR. In addition to biochemical analyses, cytochemistry using DAB as a substrate was used to identify spatial changes in peroxidase activity and to relate such changes to the timing and location of
H2O2 production, papilla
formation, and the development of the HR.
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MATERIALS AND METHODS |
Plant Inoculation
Alternate panels in leaves of 4-week-old lettuce (Lactuca
sativa L.) plants were inoculated with suspensions of
108 bacteria mL 1 in
sterile distilled water as previously described (Bestwick et al.,
1995). Inoculated areas were excised from leaves using new razor
blades. In some experiments 10 leaf discs, 1.4 cm in diameter, were
removed from the inoculation sites with a cork borer. The tissue pieces
were weighed and placed in perforated aluminum foil bags, which were
then plunged into liquid nitrogen and stored at  80°C for no longer
than 24 h following excision.
Extraction of Enzymes from Leaf Tissue
Frozen leaf tissue was added to a mortar containing liquid
nitrogen. The nitrogen was allowed to evaporate and the tissue was
ground to a fine powder, which was then transferred to a mortar chilled
on ice containing 200 mg of coarse sand and extraction buffer (4°C)
containing 50 mM phosphate buffer
(Na2HPO4/NaH2PO4), pH 6.8, and 1% (w/v) sodium metabisulphite. Homogenization was for 1 min with a ratio of 6 mL of buffer:1g fresh weight leaf tissue, and
then polyvinylpolypyrrolidone was gently mixed into the grindate (0.5 g
polyvinylpolypyrrolidone:1 g fresh weight tissue). The grindate was
filtered through a layer of Miracloth (Calbiochem), and the filtrate
was centrifuged at 20,000g for 25 min at 4°C. Supernatants
were desalted by loading onto spin columns of Sephadex G-15 (Pharmacia)
or they were concentrated and desalted by ultrafiltration using 10,000 Mr cutoff microconcentrators (Amicon,
Beverly, MA). Finally, supernatants were aliquoted, glycerol was added
to give a final concentration of 10% (v/v), and the samples were
snap-frozen in liquid nitrogen and stored at 80°C. Protein
concentrations were determined by the method of Bradford (1976).
Extraction of Intercellular Fluid
Intercellular fluid was extracted essentially according to the
method of Rathmell and Sequeira (1974). Ten discs, 1.4 cm in diameter,
were removed from lettuce leaves using a cork borer. The leaves were
gently washed in sterile water at 4°C for 3 min and were then blotted
dry and placed in 3 mL of 10 mM phosphate buffer
(Na2HPO4/NaH2PO4)
at 4°C, pH 6.8, containing 0.1% (w/v) sodium metabisulphite. Discs
were infiltrated under vacuum (100 KPa) for a period of 3 min, with
the vacuum broken and reinstated every 30 s to achieve effective
infiltration of the tissue. The infiltrated leaf discs were then washed
in sterile water, dried by blotting on filter paper, and transferred to
the barrel of a 5-mL syringe that had been placed inside a centrifuge
tube. The discs were centrifuged at 500g for 10 min at
4°C. The resulting fluid expressed (consistently 80 µL) was
used as the source of extracellular enzymes.
Enzyme Assays
All assays were performed in 1-mL volumes at 25°C in a
UV/visible light spectrophotometer (model SP8-100, Pye Unicam,
Cambridge, UK). Increases in absorbance were recorded at selected
wavelengths depending on the substrate. For measurement of guaiacol
peroxidase activity (A470) the assay
contained 800 µL of 10 mM guaiacol in 50 mM
potassium phosphate buffer, pH 6.0, 10 to 80 µL of extracted supernatant or intercellular fluid, 20 to 90 µL of sterile water, and
100 µL of 35 mM
H2O2. TMB peroxidase
activity (A654) was measured with a minor
modification to the method of Imberty et al. (1984). The assay
contained 50 µL of 20 mM TMB dissolved in absolute
ethanol, 800 µL of sodium phosphate-citric acid buffer (90 mM Na2HPO4 and 55 mM citric acid), pH 4.5, 0 to 45 µL of water, 5 to 50 µL of total soluble extract or intercellular fluid, and 100 µL of
35 mM H2O2.
Assays of chlorogenic acid peroxidase
(A410) and caffeic acid peroxidase
(A450) were conducted essentially as described by Mäder et al. (1977). The chlorogenic acid peroxidase assays comprised 800 µL of 50 mM potassium phosphate buffer, pH
6.5, 50 µL of 80 mM chlorogenic acid, 100 µL of 35 mM H2O2, 10 to
50 µL of extracted protein, and 0 to 40 µL of water. The caffeic acid peroxidase assay contained 800 µL of McIlvaine buffer (0.043 M citric acid and 0.114 M
Na2HPO4), pH 5.5, 50 µL
of 80 mM caffeic acid, 100 µL of 35 mM
H2O2, 10 to 50 µL of
extract/intercellular fluid, and 0 to 40 µL of water. Peroxidase
assays were initiated by the addition of
H2O2.
NADH and NADPH oxidase activities in supernatants and intercellular
fluid were measured by following the decrease in
A340 nm according to the method
of Mäder and Amberg-Fisher (1982). For time-course studies the
assay contained 45 mM Mes, pH 6.0, 0.15 mM
NADPH or NADH, 0.1 mM MnCl2, 1 mM 2,4-dichlorophenol, and a suitable quantity of
supernatant or intercellular fluid.
Glc-6-P dehydrogenase activity was measured as described by Reimers et
al. (1992). The reaction mixture in a volume of 1 mL contained 100 µM Tricine buffer (adjusted to pH 8.0 with 0.1 M NaOH), 2 µM MgCl2, 6 mM Glc-6-P, 0.6 mM NADP+,
and 25 µL of diluted supernatant or intercellular fluid, and the
increase in A340 was monitored.
The effect of protein concentration on activity was assessed for all
enzymes studied. All peroxidase activities measured at their optimal pH
showed linear increases in activity with increasing protein
concentrations, suggesting that there were no effects of
inhibitors/effectors within the supernatants. Where appropriate, the pH
optima and the effect of substrate concentration on activity were also
determined. Buffers were prepared as described by Dawson et al. (1989).
To determine the efficiency of extraction of enzymes, the pellets
resulting from the 20,000g centrifugation were re-extracted with buffer containing 0.1% (w/v) Triton X-100; no significant further
release of peroxidase or NADH/NADPH oxidase activity was detected.
Recovery experiments were conducted by adding 25 µL of a 10% (w/v)
preparation of horseradish peroxidase (Sigma) to the extraction buffer.
Approximately 91% of activity was recovered under the optimized
extraction conditions.
Supernatants from sonicated bacteria were analyzed for peroxidase
activity, but even in the presence of 80 µg of protein from the
supernatants, no activity was detected.
Peroxidase Localization
The protocol used was a modification of that of Sexton and Hall
(1978). Samples of lettuce leaf panels were cut under fixative into 1- to 3-mm2 pieces, which were then fixed in a
mixture of 1% (v/v) glutaraldehyde/1% (w/v) paraformaldehyde at pH
7.0 in buffer A (50 mM sodium cacodylate) for 45 min at
room temperature. Following fixation, samples were washed twice for 10 min in buffer A and transferred for 30 min into 50 mM
potassium phosphate buffer at pH 7.8 (buffer B) or at pH 6.0 (buffer
C). Following washing, samples were transferred to 0.5 mg
mL1 DAB and 5 mM
H2O2 dissolved in either
buffer B or C. To prevent autooxidation of DAB, the staining medium was
freshly prepared and staining was undertaken in dim light. Optimal
times for staining were determined by incubation for 10, 15, 30, 60, and 90 min.
Samples were washed twice for 10 min in buffer B or C, postfixed in 1%
(w/v) osmium tetroxide in buffer A for 45 min, and washed twice for 10 min in buffer A and twice for 10 min in water. Subsequent dehydration
was undertaken in a graded-ethanol series. Following two 10-min washes
in 100% ethanol, samples were transferred to propylene oxide for two
washes of 10 min each. Samples were then embedded in
Epon-araldite resin following progressive incubation in
propylene oxide/resin mixtures, with the ratio of propylene oxide to
resin decreased as follows: 3:1 for 15 min, 2:1 for 12 h or
overnight, 1:1 for 30 min, 1:2 for 30 min, and 1:3 for 30 min.
Following incubation in resin alone for 12 h, samples were transferred to fresh resin for 4 h. Finally, samples were placed in molds containing fresh resin and polymerized at 60°C for 48 h. Blocks were sectioned (70-90 nm) using a diamond knife (Diatome, Bienne, Switzerland), mounted on uncoated copper grids (300-mesh, Agar
Aids, Bishops Stortford, UK), and examined without further treatment or
stained with uranyl acetate and lead citrate.
To inhibit all enzyme activity following washing in buffer B or C,
selected leaf samples were heated at 95°C for 15 min. To determine
the H2O2 dependency of
staining, H2O2 was omitted
from the staining medium. Furthermore, because
H2O2 may be generated in
planta, samples were preincubated in buffer B or C containing 20 µg
mL1 catalase (commercial preparation from
bovine liver; Sigma) and then placed in a staining medium containing
catalase from which H2O2
had been omitted. To inhibit endogenous catalase activity, samples were
incubated for 30 min in 20 µM ATZ in buffer B
or C prior to staining. During staining, 20 µM ATZ was
also included in the staining medium. To inhibit peroxidase, catalase,
and Cyt oxidase activity, samples were incubated in 5 mM
KCN for 30 min prior to staining, and cyanide was included in the
staining medium.
H2O2 Localization
H2O2 production was
assessed cytochemically via determination of cerium perhydroxide
formation after reaction of CeCl3 with endogenous
H2O2 as previously
described (Bestwick et al., 1997). Sites of positive staining, given
categories 1 to 3 (lowest to highest intensity of staining) in the cell
wall were recorded as described by Bestwick et al. (1997), and an index
of cerium perhydroxide deposition was calculated using the formula:
([percentage of sites in category 1 × 1] + [percentage of
sites in category 2 × 2] + [percentage of sites in category
3 × 3])  3, giving a maximum possible score of 100.
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RESULTS |
Lettuce HR
The development of the HR induced in lettuce by P. s.
pv phaseolicola has already been described in detail
(Bestwick et al., 1995). Following an induction time of approximately
2.5 h, visible signs of cell collapse are observed by 5 h
after inoculation, and the lesion is brown and dessicated by 16 h.
Analysis of cell plasmolysis indicated that irreversible membrane
damage is detectable in approximately 10% of cells as early as 3 h after inoculation (i.e. 0.5 h postinduction time). The
progressive development of membrane damage is followed by increased
electrolyte leakage, which coincides with increasing ultrastructural
evidence of cell decompartmentation. The hrpD mutant induces
only a transient glazing of the leaf surface, occurring between 5 and
12 h after inoculation, but ultrastructural and plasmolysis
studies demonstrated that approximately 10% of cells collapsed in this
interaction within the first 24 h after inoculation.
Peroxidase Activity in Noninoculated Tissue
Given the broad substrate (hydrogen donor) utilization by
peroxidase, we assayed activity in the presence of both natural (caffeic and chlorogenic acid) and artificial (guaiacol and TMB) hydrogen donors. Both caffeic and chlorogenic acids have been described
as abundant phenolic compounds in lettuce tissues (Bennett et al.,
1996). NADH and NADPH were also added to supernatants, because they
represent potential in vivo reductants for peroxidase-catalyzed H2O2 generation (Vianello
and Macri, 1991).
Different substrates were used, and significant differences in pH
optima were found as follows: TMB, 4.5; guaiacol, 6.0; caffeic acid,
5.5 to 6.0; and chlorogenic acid, 6.0 to 6.5 (Fig.
1). The addition of the catalase
inhibitor ATZ (20 µM) to prevent depletion of
H2O2 from the assay medium
did not influence pH profiles. No activity was detected following
denaturation of extracts for 5 min in a boiling water bath. The
omission of H2O2 or the
inclusion of 20 µg mL1 catalase abolished the oxidation
of guaiacol, TMB, and caffeic acid. A low rate of heat-denaturable,
catalase-insensitive oxidation of chlorogenic acid was observed and a
correction was made for this rate. All peroxidase activities were
totally inhibited by 20 µM KCN.
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| Figure 1.
pH profiles of soluble peroxidase-oxidase
activities within 20,000g lettuce leaf supernatants.
Effect of pH on peroxidase activity with different substrates: A,
guaiacol; B, TMB; C, caffeic acid; D, chlorogenic acid. E and F,
Effects of pH on the oxidation of NADH and NADPH. Buffers used were Mes
( ), citric acid-sodium phosphate ( ), Mops ( ), N-2
hydroxyethylpiperazine-N-3-propane sulfuric acid ( ),
borate-HCl ( ), Tris-maleate ( ), and potassium phosphate ( ).
Data are the means ± SE of a minimum of four
experiments, each consisting of two replicates.
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The rates of heat-denaturable NADPH and NADH oxidation by leaf extracts
were determined (Fig. 1, E and F). Because numerous enzymes may be
responsible for this oxidation, we attempted to stimulate peroxidases
specifically by addition of Mn2+ and monophenols,
as demonstrated by Vianello and Macri (1991). A combination of 0.1 mM MnCl2 and 1 mM
2,4-dichlorophenol produced maximal, heat-sensitive stimulation of both
NADH and NADPH oxidation, both of which had optima between 5.5 and 6.5. Oxidation was accompanied by catalase-sensitive tetraguaiacol
formation, indicating the production of
H2O2.
Changes in Total Peroxidase, NADH, and NADPH Oxidase Activities
after Inoculation
Time courses were conducted with both the "artificial" and the
"natural" substrates and, as shown in Figure
2, certain common alterations to activity
were observed at their pH optima. First, there was a rapid decline in
activity following all inoculations. A detailed time course using
guaiacol as the substrate (Fig. 2, A and B) revealed that this decline
occurred by approximately 15 min after infiltration and represented a
striking decrease to only 11% to 15% of the activity in uninoculated
tissue. Activity gradually recovered to preinoculation levels but
increased further only in response to bacterial challenge. The timing
of recovery and the extent of the increase depended on the hydrogen
donor used. Guaiacol peroxidase activity recovered more rapidly in
tissues undergoing the HR and eventually reached levels considerably in excess of those in tissues responding to the hrp mutant.
However, increases in TMB peroxidase were similar in response to both
wild-type and mutant bacteria. Increases in chlorogenic acid and
caffeic acid peroxidase activity were significantly higher in tissues inoculated with the hrp mutant relative to those undergoing
the HR. Both caffeic acid and chlorogenic acid peroxidase activity in
water-inoculated tissues recovered to preinoculation levels more
rapidly than that of extracts from tissues inoculated with bacteria.
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| Figure 2.
Changes in peroxidase activity within inoculated
lettuce leaves. Effects of inoculation with wild-type () and
hrp D mutant () strains of P. s. pv
phaseolicola, or sterile water () were examined.
Activities of guaiacol (A-C) and TMB peroxidase (D) are presented as a
function of total soluble protein (A, B, and D) or leaf fresh weight
(C). Changes in caffeic acid peroxidase activities (E) and chlorogenic
acid (F) are presented as a function of protein concentration. Data are
means ± SE of activities from three replicate
supernatants; similar trends were found in repeated experiments. Note
that tissues desiccated during the HR. fwt, Fresh weight.
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Changes in NADPH oxidase activity in the presence of 0.1 mM
MnCl2 and 1 mM 2,4-dichlorophenol are
presented in Figure 3. A profile similar
to that of peroxidase activity was determined: an initial decline
followed by recovery, which was far more rapid in tissues undergoing
the HR.
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| Figure 3.
Changes in NADPH oxidase activity within
inoculated lettuce leaves. Effect of inoculation with wild-type ()
and hrp D mutant () strains of P. s.
pv phaseolicola or sterile water () on the rate of
2,4-dichlorophenol/Mn2+-stimulated NADPH oxidation by
lettuce leaf extracts. A and B, Rates of NADPH oxidation
mg1 protein and g1 fresh leaf weight,
respectively. Data are means ± SE of three
replicates; duplicate experiments demonstrated a similar trend in both
NADPH and NADH oxidation. Note that tissues desiccated during the HR.
fwt, Fresh weight.
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Changes in Extracellular Peroxidase, NADH, and NADPH Oxidase
Activities following Inoculation
Within the first 5 h of the HR, no activity of the marker
enzyme Glu-6-P dehydrogenase could be detected in the intercellular fractions, implying no appreciable symplastic contamination of the
apoplast (Fig. 4A). However, after 5 h the extensive cell collapse occurring during the HR resulted in
varying levels of contamination. Time-course studies of enzyme
activities were therefore confined to the first 5 h after
inoculation.
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| Figure 4.
Changes in peroxidase and oxidase activities
determined in intercellular fluids. A, Activity of the cytoplasmic
marker enzyme Glc-6-P dehydrogenase within homogenates () and
intercellular fluids () extracted from lettuce leaves inoculated
with wild-type P. s. pv phaseolicola.
Changes in the activities of guaiacol and TMB peroxidase (B and C) NADH
oxidase (D) following inoculation with wild-type () or hrp
D mutant () of P. s. pv
phaseolicola or sterile water () are shown. Analysis
of NADPH oxidation by intercellular fluids revealed a trend similar to
that of NADH oxidation. Data are based on the intercellular fluids
recovered from three leaf discs. Means ± SE of four
replicates are given; the trends observed were confirmed in repeated
experiments.
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No peroxidase activity was detected using natural hydrogen donors, but
considerable H2O2-dependent
activity was found in the presence of guaiacol and TMB (Fig. 4, B and
C). Following an initial decline, guaiacol peroxidase activity
increased in tissue challenged with both the hrp mutant and
the wild-type strain, the rate and magnitude of increase being far
greater in tissues undergoing the HR. With TMB, no decline in
peroxidase activity was observed after bacterial infiltration, but
bimodal increases occurred in response to both strains, with a much
greater increase occurring during the HR (Fig. 4D). The peroxidases
recovered in intercellular fractions would be expected to represent a
subset of isoforms contributing to total soluble peroxidase activity.
An increase in Mn2+/2,4-dichlorophenol-stimulated
NADH and NADPH oxidase activity was observed by 1 h after
inoculation with both bacterial strains. Between 3 and 5 h a
second striking increase in activity occurred in response only to
wild-type bacteria (Fig. 4D).
Localization of Peroxidase Activity
Noninoculated Leaves
Following fixation in paraformaldehyde/glutaraldehyde, peroxidase
was detected by the formation of osmium black after treatment of
sections with DAB/H2O2 at
pH 6.0 or 7.8. Staining was first observed after 15 min of incubation
in DAB/H2O2 at pH 6.0 and after 30 min at pH 7.8, and in both cases optimal stain intensity was
reached after 60 min. Despite the short fixation times used (45 min),
cell ultrastructure was remarkably well preserved. When present,
peroxidase activity was usually observed in the middle lamella and on
the extracellular face of the cell wall (Fig.
5).
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| Figure 5.
Cytochemical localization of peroxidase activity
in uninoculated tissue. Note that staining, indicated by the formation
of electron-dense deposits (arrows), is mainly located on the
extracellular face of the plant cell wall and in the middle lamella.
Bar = 0.5 µm. IS, Intercellular space; C, chloroplast.
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Inoculated Leaves
For the time-course studies, tissues were stained at pH 6.0. Between 3 and 8 h after inoculation, a localized increase in peroxidase activity, indicated by the presence of electron-dense deposits, was observed in cell walls at the sites of both wild-type and
hrp mutant attachment and, therefore, to sites of papilla development (Fig. 6A). Coincident with
increased wall activity, peroxidase was also detected within the
nuclear envelope, the ER, the Golgi apparatus, and variously sized
vesicles within the challenged cells (Fig. 6, B and C). High levels of
peroxidase were also identified within the material surrounding and
encapsulating both strains of bacteria on the epidermal and mesophyll
cell walls (Fig. 6, B and C).
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| Figure 6.
Peroxidase activity in the cell wall and cytoplasm
of mesophyll cells after inoculation with P. s. pv
phaseolicola. A, Low magnification (bar = 2 µm)
showing the extension of electron-dense staining (small arrows) from
sites of bacterial attachment (wild-type strain) onto the plant cell
wall 5 h after inoculation. Note that the cell wall away from the
bacteria is unstained (large arrow). The large papilla (asterisk) has
patchy staining that is more distinct at higher magnification in B. Bar = 0.5 µm. B, Staining for peroxidase can be seen to extend
into the material surrounding bacteria, and activity is also associated
with the rough ER (arrows) and small vesicles in the dense plant cell
cytoplasm at reaction sites. C, Reaction to the hrpD
mutant 8 h after inoculation includes increased peroxidase
activity around attached bacteria, within the plant cell wall, and
within small vesicles (arrow). B, Bacterium; IS, intercellular space;
CV, central vacuole; M, mitochondrion.
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In cells showing signs of collapse during the HR, peroxidase activity
was not greater than that detected in cells with little evidence of
disruption (Fig. 7, A and B).
Condensation of the cytoplasm to form an electron-dense osmiophilic
"corpse" during the terminal phases of the HR prevented further
observations on cytoplasmic peroxidase in this interaction. The
detection of peroxidase activity in the cell wall also became
increasingly inconsistent and it was necessary to include ATZ to detect
peroxidase reliably within the walls of collapsing cells. By contrast,
high levels of peroxidase activity were maintained at sites of
hrp mutant attachment (Fig. 6C).
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| Figure 7.
Peroxidase localization in cells collapsing during
the HR and controls showing the specificity of staining for enzyme
activity. A and B, Sites undergoing the HR in response to the wild-type
strain 5 and 8 h after inoculation, respectively. Note that
activity (arrows) is detected in the cell wall, ER, and small vesicles.
Staining within the wall is less intense than observed at earlier
stages of the plant cell's response (Fig. 6). Bars = 1 and 2 µm
in A and B, respectively. C and D, Sections from sites 3 and 8 h
after inoculation with the wild-type strain used as controls in which 5 mM KCN was added to the staining medium to inhibit
peroxidase activity. The sites examined were directly comparable to
those shown in Figure 6, in which the similar electron density of
organelles such as mitochondria, which lack peroxidase activity,
contrasts with the strongly stained plant cell wall in the absence of
KCN. Bars = 0.5 and 2 µm for C and D, respectively. B,
Bacterium; IS, intercellular space; CW, cell wall; CV, central vacuole;
C, chloroplast.
|
|
Controls
Catalase may be detected using DAB as a substrate because of its
"peroxidative" activity, which is reportedly favored by fixation in
glutaraldehyde (Lewis, 1977). Although considerable ATZ-sensitive staining of peroxisomes and the surrounding cytoplasm was observed following incubation at pH 7.8, no such staining was apparent at pH
6.0. In addition, at pH 6.0, ATZ did not affect the distribution of DAB
oxide during the first 8 h after inoculation but was required to
observe oxide formation within the cell walls of collapsed tissues
during the later stages of the HR. There have also been reports that
DAB may serve as a substrate for polyphenol oxidase, but no staining
could be attributed to this enzyme, since thylakoid membranes were not
stained and the polyphenol oxidase inhibitor diethyldithiocarbamate
failed to prevent staining. All staining was heat labile and was
verified as being associated with the activity of a heme-containing
protein by its inhibition with cyanide (Fig. 7). The
H2O2 dependence of DAB
oxide formation was confirmed when the omission of
H2O2 from the staining
medium totally abolished activity in noninoculated tissues; however,
during the HR the inclusion of catalase was required to
prevent staining completely.
H2O2
The distribution of peroxidase activity was closely compared with
the accumulation of H2O2
detected by the catalase-sensitive formation of cerium perhydroxides at
reaction sites after incubation with CeCl3. As
previously reported by Bestwick et al. (1997), the main site of
H2O2 accumulation was
apoplastic, with staining present in both the cell wall and the
papillae (Fig. 8A). We also observed a
previously unreported, less frequent, and nonlocalized accumulation of
cerium perhydroxides within the symplasm of cells undergoing the HR
(Fig. 8B). The more widespread accumulation of cerium perhydroxides was
first apparent 8 h after inoculation and was not necessarily
associated with visible cellular disruption.
View larger version (116K):
[in this window]
[in a new window]
| Figure 8.
Localization of H2O2
accumulation in lettuce cells adjacent to wild-type P. s. pv phaseolicola 5 h after inoculation.
A, Electron-dense deposits of cerium perhydroxides extending from the
site of attachment of bacteria into the surrounding plant cell wall
(arrows). Bar = 1 µm. B, Detection of
H2O2 in cells undergoing the HR. Deposits of
cerium perhydroxide (arrows) are present in both the cell wall and in
the degenerating cytoplasm. Bar = 0.5 µm. B, Bacterium; IS,
intercellular space; CW, cell wall; CV, central vacuole; C,
chloroplast.
|
|
An index based on the frequency and intensity of perhydroxide formation
in the apoplasm was devised based on the scoring system reported by
Bestwick et al. (1997). Time-course studies using the index to
summarize staining (Fig. 9) revealed that
high levels of H2O2 were
maintained within the apoplast during the HR, with peak
H2O2 production coinciding
with the onset of membrane damage (Bestwick et al., 1995). Treatment
with ATZ led to the maintenance of a high cerium perhydroxide index
throughout the time course of the HR, indicating that
H2O2 continued to be
produced but was scavenged by catalase, probably as it was released
from degrading peroxisomes. Unfortunately, it was not possible to
devise an index for cytoplasmic
H2O2
production/accumulation because the increased electron density
associated with the condensation of the cytoplasm at later times,
representing the terminal phase of cell death, often obscured
perhydroxide formation.
View larger version (20K):
[in this window]
[in a new window]
| Figure 9.
H2O2 accumulation during
the HR. Tissues were stained with CeCl3 and processed for
electron microscopy, and an intensity index for
H2O2 production was compiled as described in
``Materials and Methods'' (maximum score, 100). Tissues inoculated
with wild-type (circles) or hrpD mutant (squares) of
P. s. pv phaseolicola were analyzed in
the presence (open symbols) or absence (solid symbols) of the catalase
inhibitor ATZ. Data are from analysis of a minimum of 30 interaction
sites at each time after inoculation. Key stages of cellular responses
are marked by numbers: 1, completion of the induction time; 2, 30% of
mesophyll cells fail to plasmolyze; 3, 10% of cells fail to plasmolyze
in response to the hrp mutant; 4, 70% of cells fail to
plasmolyze and 50% have collapsed during the HR.
|
|
| |
DISCUSSION |
Both the activity and distribution of peroxidase were altered
within lettuce cells following the inoculation of leaves with wild-type
and hrp bacteria. The changes observed
indicate three levels of response: (a) nonspecific changes associated
with inoculation; (b) changes associated with general defense
responses; and (c) responses that have relevance to the development of
the HR. Such complexity is perhaps predictable given the
multifunctional role that isoforms of peroxidase may play in plant
stress responses.
The initial decline in peroxidase activity is a response to inoculation
because it was induced by water infiltration and bacterial treatment.
Such decreases may arise from an inhibition of enzyme synthesis or
activity or from relocalization to an environment such as the cell
wall, which might be recalcitrant to buffer extraction. Subsequent
increases in both apoplastic and symplastic peroxidase activities to
levels above those found in noninoculated leaves were, however, found
only in tissues challenged by bacteria. In general, increases in total
and apoplastic peroxidase activities both correlated well with the
microscopic detection of DAB oxide formation in cells challenged by
bacteria; however, there were significant differences in the changes in
activity recorded using various hydrogen donors as substrates. There
appear to be differences in the substrate specificities of the
isoenzymes that make up the total activities measured, which may
reflect the different nature of the resistant responses induced by
wild-type and hrp strains.
Cytochemical localization and assays of intercellular fluids
demonstrated that early increases in activity were mainly confined to
the apoplasm and, in particular, to sites of bacterial attachment to
lettuce cell walls. A rapid synthesis and site-directed secretion of
the enzyme is probable, because the increase in apoplastic peroxidase
activity coincided with or occurred following the appearance of DAB
oxide within the endomembrane system. The addition of heme to the
apoprotein moiety may occur within the ER; therefore, the synthesis of
isoperoxidases may first become detectable by cytochemistry during this
stage of processing (Birecka et al., 1975; Mendgen, 1975; Ebermann and
Stich, 1986). The heavily stained vesicles in the cytoplasm of cells
challenged by bacteria may be involved in peroxidase secretion, as
suggested by van Huystee (1987). Pre-existing symplastic isoperoxidases
may also be relocated to the wall, and it is also possible that a
component of the increased wall activity represents the activation of
pre-existing wall-bound peroxidase.
The sites of increased peroxidase activity in the cell wall are also
regions of papilla development. Ca2+, which has
been implicated in the deposition of callose (a major component of
papillae), is known to regulate peroxidase secretion and may therefore
act as a local signal for site-specific structural alterations (Sticher
et al., 1981; Aist and Gold, 1987). It seems likely that peroxidase is
responsible for cross-linking the phenolic compounds and Hyp-rich
glycoproteins found within papillae and also for the modification of
the cell wall at reaction sites (Bestwick et al., 1995). Such reactions
are H2O2 dependent, and
increased concentrations of
H2O2, as revealed by
catalase-sensitive cerium perhydroxide precipitation, were also
localized to sites of papilla formation.
Clearly, there is no precise relationship between peroxidase levels and
H2O2 accumulation. The
markedly lower levels of
H2O2 in the interaction
with the hrp mutant contrasts sharply with the generally
high level of peroxidase in this interaction, as determined both
cytochemically and in assays using "natural" phenolic substrates.
Nevertheless, our data demonstrate the production of
H2O2 in response to the
hrp mutant, with the ROS available (although at low levels)
for cross-linking reactions mediated by peroxidase. It remains to be
determined whether cross-linking reactions are effectively restrained
by the lower level of H2O2 prevalent within the cell wall and papillae adjacent to the
hrp mutant, or whether an excess of
H2O2 is produced during the interaction with
the wild-type strain (Bestwick et al., 1997). The extent to which
cross-linking occurs may be resolved by examining the protoplasting
efficiency of tissues in both interactions and by the susceptibility of
wall-bound phenolic compounds to removal by mild saponification
(Brisson et al., 1994; Bennett et al., 1996).
The origin of the H2O2
produced during defense reactions in lettuce is intriguing. The largely
apoplastic location of cerium perhydroxide formation suggests that
either the plasma membrane or the cell wall is the primary site of the
superoxide/H2O2 generator. Isolated plant protoplasts are capable of generating elictor-induced oxidative bursts, and a membrane-associated, neutrophil-like NADPH oxidase complex is thought to be responsible for
superoxide/H2O2 generation
within elicitor/microbe-stimulated soybean, potato, parsley, and rose
cells (Lamb and Dixon, 1997). However, in cotton challenged with
Xanthomonas, superoxide generation has been linked to the
apoplastic NADH oxidase activity of a peroxidase-like enzyme, and a
cell wall peroxidase has also been implicated in the generation of an
elicitor-induced oxidative burst in bean cells (Bolwell et al., 1995;
Martinez et al., 1997). The potential for ROS production by systems
other than membrane-bound NADPH oxidase is indicated from experiments
in which cell wall preparations from a number of species have been
shown to generate oxidative bursts when challenged with elicitors or
digested with cell wall-degrading enzymes (Ishii, 1987; Kiba et al.,
1997). In tobacco there is evidence that multiple oxidase systems may
operate and that individual contributions to
H2O2 production may vary
between interactions/elicitors (Allan and Fluhr, 1997).
In lettuce low concentrations of the neutrophil NADPH oxidase inhibitor
diphenylene iodonium significantly reduced
H2O2 levels detected by
staining with CeCl3, and the in vivo production
of H2O2 was also cyanide
and azide sensitive, suggesting the involvement of both a
neutrophil-like NADPH oxidase and peroxidase in ROS generation
(Bestwick et al., 1997). Recently, Frahry and Schopfer (1998)
demonstrated that the oxidase activity of horseradish peroxidase is
also reduced by diphenylene iodonium, indicating that the inhibitor should no longer be considered a reliable marker for neutrophil-like NADPH oxidase in plants. We have now shown that heightened apoplastic Mn2+/2,4-dichlorophenol stimulated NADH and NADPH
oxidase activity, which is characteristic of certain peroxidases
(Vianello and Macri, 1991), coincides with peak
H2O2 production during the
HR.
Overall, our results suggest that certain isoforms of peroxidase with
NAD(P)H oxidase activity are a major source of ROS production in
lettuce. We have used NADPH and NADH as substrates for peroxidase-based H2O2 generation, and
although the former may not be physiologically relevant in the
apoplast, NADH may be supplied to the cell wall via a malate
oxaloacetate shuttle across the plasma membrane involving a wall-bound
malate dehydrogenase (Gross et al., 1977). Additional isoforms of
peroxidase that may be strongly bound to the cell wall (either
ionically or covalently) and may not have been extracted may also
contribute to H2O2
generation in the apoplast. The isolation of peroxidase isoforms and
the characterization of their enzymic activities is needed to confirm
their proposed role in the HR.
Electron microscopy has demonstrated that the incorporation of new
material into cell walls and papilla formation often occur in the
absence of the HR (Bestwick et al., 1995; Brown et al., 1995). The
extent to which oxidative processes are necessary for and controlled in
these reactions may determine the continued viability of lettuce cells.
It is possible that an imbalance in peroxidase-oxidase activity during
the HR contributes to an increase in
H2O2 and, therefore, the
cerium perhydroxide precipitation observed in our experiments. For
example, although increases in soluble peroxidase activity within
tissues challenged by wild-type and hrp mutant strains are
initially similar, the
Mn2+/2,4-dichlorophenol-stimulated NADH/NADPH
oxidase activity is primarily enhanced in response to challenge with
wild-type bacteria. Significantly, during the HR, increases in
extracellular, peroxidase-like NADH/NADPH oxidase activity preceded
major increases in extracellular peroxidase activity detected in
intercellular fluids. Peroxidase activity driven by
H2O2 generated either by
peroxidase isoforms or other sources (such as a neutrophil-like NADPH
oxidase complex) may subsequently reduce the level of free
H2O2.
In conclusion, our experiments demonstrate that resistance responses,
including ROS production, are highly localized and tightly controlled
within sites in contact with the invading microorganism. The
localization observed highlights the potential importance of specific
isoforms of peroxidase and reactions occurring in the apoplasm in
mediating both the HR and other processes of resistance.
| |
FOOTNOTES |
1
This work was supported by grants from the
Biotechnology and Biological Sciences Research Council (UK) and the
European Union Biotechnology Framework IV program.
2
Present address: Division of Micronutrient and
Lipid Metabolism, Rowett Research Institute, Greenburn Road, Bucksburn,
Aberdeen, AB21 9SB, UK.
*
Corresponding author; e-mail csb{at}rri.sari.ac.uk; fax
44-1-224-716687.
Received April 20, 1998;
accepted August 17, 1998.
| |
ABBREVIATIONS |
Abbreviations:
AZT, 3-amino-1,2,4-triazole.
DAB, 3 3-diaminobenzidine.
HR, hypersensitive reaction.
ROS, reactive
oxygen species.
TMB, tetramethylbenzidine.
| |
ACKNOWLEDGMENTS |
We wish to thank Shelagh Reardon for assistance with electron
microscopy and Atilla Ádám for valuable discussion about
the role of peroxidase in the HR.
| |
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Top
Abstract
Introduction