Plant Physiol. (1999) 121: 97-112
Stress-Induced Legume Root Nodule Senescence. Physiological,
Biochemical, and Structural Alterations1
Manuel A. Matamoros,
Lisa M. Baird,
Pedro R. Escuredo,
David A. Dalton,
Frank R. Minchin,
Iñaki Iturbe-Ormaetxe,
Maria
C. Rubio,
Jose F. Moran,
Anthony J. Gordon, and
Manuel Becana*
Departamento de Nutrición Vegetal, Estación
Experimental de Aula Dei, Consejo Superior de Investigaciones
Científicas, Apdo 202, 50080 Zaragoza, Spain (M.A.M., P.R.E.,
I.I.-O., M.C.R., J.F.M., M.B.); Biology Department, University of San
Diego, San Diego, California 92110 (L.M.B.); Biology Department, Reed
College, Portland, Oregon 97202 (D.A.D.); and Institute of Grassland
and Environmental Research, Plas Gogerddan, Aberystwyth SY23 3EB,
United Kingdom (F.R.M., A.J.G.)
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ABSTRACT |
Nitrate-fed and dark-stressed bean
(Phaseolus vulgaris) and pea (Pisum
sativum) plants were used to study nodule
senescence. In bean, 1 d of nitrate treatment caused a partially
reversible decline in nitrogenase activity and an increase in
O2 diffusion resistance, but minimal changes in carbon
metabolites, antioxidants, and other biochemical parameters, indicating
that the initial decrease in nitrogenase activity was due to
O2 limitation. In pea, 1 d of dark treatment led to a
96% decline in nitrogenase activity and sucrose, indicating sugar
deprivation as the primary cause of activity loss. In later stages of
senescence (4 d of nitrate or 2-4 d of dark treatment), nodules showed
accumulation of oxidized proteins and general ultrastructural
deterioration. The major thiol tripeptides of untreated nodules were
homoglutathione (72%) in bean and glutathione (89%) in pea. These
predominant thiols declined by approximately 93% after 4 d of
nitrate or dark treatment, but the loss of thiol content can be only
ascribed in part to limited synthesis by 
-glutamylcysteinyl,
homoglutathione, and glutathione synthetases. Ascorbate peroxidase was
immunolocalized primarily in the infected and parenchyma (inner cortex)
nodule cells, with large decreases in senescent tissue. Ferritin was almost undetectable in untreated bean nodules, but accumulated in the
plastids and amyloplasts of uninfected interstitial and parenchyma
cells following 2 or 4 d of nitrate treatment, probably as a
response to oxidative stress.
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INTRODUCTION |
Legume N2 fixation is particularly sensitive
to environmental perturbations, including defoliation, water deficit,
continuous darkness, and nitrate fertilization (Vance et al., 1979
;
Witty et al., 1986; Layzell et al., 1990). In most types of stress, the
initial decrease of nitrogenase activity is associated with a decline
in the O2 concentration reaching the infected
cells and bacteroids (Witty et al., 1986; Carroll et al., 1987; Layzell et al., 1990; Escuredo et al., 1996). Prolongation of stress induces premature nodule senescence, which shares some features with natural senescence (nodule aging), such as the loss of N2
fixation, the increase in lytic activities, and the formation of green
pigments from leghemoglobin (Lb) (Pfeiffer et al., 1983; Sarath et al., 1986). This stress-induced senescence has been linked to the enhanced production of oxidants and the lowering of antioxidant defenses (Escuredo et al., 1996; Gogorcena et al., 1997). Oxidants include inorganic (H2O2) and
organic (lipid) peroxides as well as "catalytic iron," the fraction
of iron in plant tissues capable of catalyzing the generation of
hydroxyl radicals through Fenton reactions (Becana et al., 1998).
A major antioxidant mechanism operating in the nodule cytosol is the
ascorbate-GSH cycle, which results ultimately in the detoxification of
H2O2 at the expense of
NAD(P)H. The pathway involves the concerted action of four enzymes:
ascorbate peroxidase (APX), dehydroascorbate reductase (DR),
monodehydroascorbate reductase (MR), and glutathione reductase (GR),
and requires a continuous supply of ascorbate, thiols, and reduced
pyridine nucleotides (Dalton et al., 1986, 1992). The initial enzyme of
the pathway, APX, may account for up to 1% of the total soluble
protein of nodules (Dalton et al., 1998). The thiol tripeptide GSH
(Glu-Cys-Gly) also participates in the removal of peroxides through
the ascorbate-GSH cycle, but it performs additional roles in plants,
such as the transport and storage of sulfur, the control of redox
status, and the detoxification of heavy metals (Rennenberg, 1995; May et al., 1998). The synthesis of GSH involves two ATP-dependent reactions catalyzed by the enzymes -glutamylcysteinyl synthetase (ECS) and GSH synthetase (GSHS). Thiol tripeptides are particularly abundant in the leaves, roots, and seeds of legumes, where a thiol tripeptide homolog homoglutathione (hGSH; Glu-Cys-
Ala), may be
present in addition to or instead of GSH (Klapheck, 1988). Apparently,
a specific hGSH synthetase (hGSHS) catalyzes the second step of hGSH
synthesis in the leaves of some legumes (Macnicol, 1987; Klapheck et
al., 1988). It is not known whether hGSH and hGSHS are present in the
nodules.
An entirely different antioxidant mechanism involving the sequestration
of catalytic iron by ferritin may also operate in the nodules. Plant
ferritins are composed of 24 subunits and can store up to 4,500 atoms
of iron in a safe, nontoxic form (Briat and Lobréaux, 1997). In
nodules, ferritin may also act as an iron reservoir for nitrogenase,
Lb, and other iron-proteins, since ferritin protein increases early in
nodulation and then declines concomitantly with the increase in
nitrogenase activity, heme, and non-heme iron (Ragland and Theil,
1993). However, little is known about ferritin in senescent nodules.
This information may be of considerable interest because nodules are
extremely rich in iron (Ragland and Theil, 1993) and this may become
available (catalytic iron) for Fenton reactions by proteolysis during
nodule senescence, either natural or induced by stress (Becana et al., 1998).
Many other important alterations related to oxygen, carbon, and
nitrogen metabolism occur in nodules during senescence, but the precise
sequence of these biochemical changes is far from clear (Carroll et
al., 1987; Layzell et al., 1990; Gordon et al., 1997). There is also a
paucity of information regarding the structural changes involved in
stress-induced nodule senescence. Light and electron microscopy studies
have been carried out with nodules of detopped alfalfa (Vance et al.,
1979), dark-treated soybean (Cohen et al., 1986), and nitrate-treated
lupine (Lorenzo et al., 1990). Nevertheless, legume symbioses differ in
their tolerance to stress, and at least some of these differences may
be related to the growth pattern (indeterminate versus determinate) of
nodules (Sprent, 1980). Comparison of the inhibitory effects of stress on other legumes may provide insight into the mechanisms underlying stress tolerance and nodule senescence.
The general objective of the present study was to ascertain the time
course of events leading to the loss of function and structural
deterioration of nodules following stress application to the plant. The
specific objective was to gain further information on the role of some
important antioxidants (APX, thiols, and ferritin) in the protection of
legume N2 fixation against the noxious effects of
peroxides, free radicals, and catalytic iron. Data presented in this
paper are intended to complement two previous reports (Escuredo et al.,
1996; Gogorcena et al., 1997) in providing an orthogonal comparison of
nitrate- and dark-stress-induced senescence in determinate and
indeterminate nodules.
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MATERIALS AND METHODS |
Plants and Treatments
Nodulated bean (Phaseolus vulgaris L. cv Contender × Rhizobium leguminosarum bv phaseoli 3622) and
pea (Pisum sativum L. cv Lincoln × R. leguminosarum bv viciae NLV8) plants were grown in a
perlite:vermiculite mixture (2:1) in controlled-environment chambers as
described by Gogorcena et al. (1997). When bean had reached the
late-vegetative stage (30-32 d), pots were divided at random into
three groups receiving 10 mM potassium nitrate for 1, 2, or 4 d, and one group of controls receiving N-free
nutrient solution and harvested on the 3rd d. Likewise, when pea plants had reached the late-vegetative stage (34-36 d), pots were divided at
random into three groups placed in the dark (with otherwise identical
conditions) for 1, 2, or 4 d, and one group of controls kept in
the light and harvested on the 3rd d. These control plant harvests were
arranged so that the maximum age difference between treated and control
plants was 2 d. The same protocol was used to produce dark-treated
bean and nitrate-treated pea plants so as to obtain nodules for
microscopic studies. Nodules to be used for light and electron
microscopic analyses were fixed immediately upon detachment, as
described under "Light Microscopy Studies." Nodules to be used for
biochemical analyses were flash-frozen in liquid
N2 and stored at 
80°C for later analysis
(within 4-5 weeks).
Nodule Activity and Carbon Metabolites
Nitrogenase activity and root respiration of intact, undisturbed
plants were measured simultaneously using a flow-through gas system
(Minchin et al., 1983) housed in a controlled-environment chamber. Root
systems were sealed in the growth pots and allowed to stabilize for
18 h. In vivo nitrogenase activity was measured as
H2 evolution (Witty and Minchin, 1998) using
electrochemical H2 sensors (City Technology,
Portsmouth, UK) and respiratory CO2 production
was measured using an IR gas analyzer. Measurements were made in air
for 5 min and then in a gas stream of 79% (v/v) Ar/21% (v/v)
O2. Following exposure to
Ar/O2, steady-state conditions were reached after
60 to 80 min and the external O2 concentration was then increased over the range of 21% to 60% (8.55-24.54 mmol O2 L
1) in steps of 5% or
10%. Each increase in O2 took 5 to 6 min and was
followed by a 20- to 25-min equilibration period. These data were used
to calculate the oxygen diffusion barrier (ODB) resistance and carbon
costs of nitrogenase, as described in Escuredo et al. (1996).
Based on the maximum H2 production under
Ar/O2, the electron allocation coefficient for
N2 was 0.67 for control pea nodules and 0.73 for
control bean nodules. The hup-negative genotype of the two
Rhizobium strains used in these studies was confirmed by
Southern-blot analysis of EcoRI-digested total DNA using a hup-specific DNA probe prepared by dioxigenin labeling of
cosmid pAL618 containing the entire hup gene cluster from
Rhizobium leguminosarum bv viciae strain UPM791
(Leyva et al., 1990).
Lb was determined by a method based on the fluorescence emitted by the
tetrapyrrol ring after removal of iron by hot, saturated oxalic acid
(LaRue and Child, 1979) using myoglobin (horse skeletal muscle,
Calbiochem) as the standard. Protein of the nodule cytosol and
bacteroids was determined with a commercial dye (Bio-Rad) using
crystalline BSA (Sigma) as the standard. Total lipids were extracted
from nodules at room temperature essentially as described by Bligh and
Dyer (1959).
Carbohydrates were extracted from 0.2 g of nodules with 10 mL of
boiling 80% (v/v) ethanol. The ethanol-soluble extracts were dried in vacuo at 40°C and the soluble compounds redissolved in 4 mL
of water. The samples were centrifuged at 20,000g for 10 min, and the contents of Glc, Fru, and Suc were determined
spectrophotometrically at 340 nm using enzymatic assays coupled to the
formation of NADH (González et al., 1995). Starch was extracted
from the ethanol-insoluble residue and quantified as the Glc released
following digestion with amyloglucosidase (MacRae, 1971).
Catalase and Enzymes of the Ascorbate-GSH Cycle
Antioxidant enzymes were extracted at 4°C from 0.5 g (bean)
or 0.25 g (pea) of nodules with a mortar and pestle. Catalase and
APX were extracted with 10 mL (bean) or 5 mL (pea) of 50 mM potassium phosphate buffer (pH 7.0) containing 0.5% (w/v) PVP-10. DR,
GR, and MR were extracted with 5 mL (bean) or 2.5 mL (pea) of 50 mM potassium phosphate buffer (pH 7.8) containing 1% (w/v) PVP-10, 0.2 mM Na2EDTA, and 10 mM -mercaptoethanol. The homogenate was filtered through
one layer of Miracloth (Calbiochem) and centrifuged at
15,000g for 20 min.
Catalase activity was assayed by following the decomposition of
H2O2 at 240 nm (Aebi,
1984). APX and DR activities were determined by measuring the oxidation
of ascorbate at 290 nm (Asada, 1984) and the formation of ascorbate at
265 nm (Nakano and Asada, 1981), respectively. MR and GR activities
were assayed by monitoring the oxidation of NADH (Dalton et al., 1992)
and NADPH (Dalton et al., 1986) at 340 nm, respectively.
All activities were measured at 25°C in 1-mL reaction mixtures within
the linear range. Measurements were made with a spectrophotometer (Lambda-16, Perkin-Cetus) during the first 1.5 to 3 min with no lag
period (except for APX, which was measured after a lag of 40 s).
Assays were made using sample volumes ranging from 10 (catalase) to 100 µL (GR). Where appropriate, controls made by omitting or boiling
extracts were run in parallel to correct for nonenzymatic rates, and
buffers and reagents were treated with Chelex resin to avoid
contamination by trace amounts of transition metal ions.
Thiols and Thiol Synthetase Activities
GSH and hGSH were extracted from 50 mg of nodules with 500 µL of
200 mM methanesulfonic acid containing 0.5 mM
diethylenetriaminepentaacetic acid. After centrifugation at
13,000g for 5 min, the supernatant was derivatized at pH 8.0 with 2 mM monobromobimane (Fahey and Newton,
1987). The bimane derivatives of GSH and hGSH were separated and
quantified by HPLC (Waters) using an analytical
C18 column (3.9 × 150 mm; 4 µm; Nova-Pak,
Waters) and a 15% (v/v) methanol/0.25% (v/v) acetic acid (pH
3.5) solvent at a constant flow of 1 mL min1.
Detection was by fluorescence (model 474 scanning fluorescence detector, Waters) with excitation at 380 nm and emission at 480 nm.
Standards of GSH (Sigma) and hGSH (obtained by chemical synthesis at
the University of Nebraska, Lincoln) were processed identically to the
samples. The proportion of thiol tripeptides present in the oxidized
form was determined by an enzymatic recycling procedure using yeast GR
to reduce the disulfide forms and vinylpyridine as a thiol blocking
agent (Griffith, 1980; Law et al., 1983).
The extraction and assays of ECS, GSHS, and hGSHS activities were
performed by modification of previous protocols, based on the HPLC
separation of synthesized Glu-Cys, GSH, and hGSH after
derivatization with monobromobimane (Hell and Bergmann, 1988; Kocsy et
al., 1996).
Other Metabolites
Ascorbate was extracted from 0.25 g of nodules with 5 mL of
5% (w/v) metaphosphoric acid and quantified by a method based on the
ascorbate-dependent reduction of iron (III) to iron (II). Formation of
the complex between iron (II) with 2,2
-dipyridyl was measured at 525 nm (Law et al., 1983).
Pyridine nucleotides were extracted from 30 mg of nodules with 2× 0.5 mL of 0.1 M NaOH (NADH and NADPH) or with 2× 0.5 mL of 5%
(w/v) TCA (NAD+ and
NADP+) at room temperature. After thorough
homogenization for 90 to 120 s in an Eppendorf tube, the extracts
were boiled for 6 min, cooled on ice, and centrifuged at
13,000g for 6 min at room temperature. The supernatant (25 µL for bean and 50 µL for pea) was made up to 100 µL with NaOH or
TCA, and the nucleotides were quantified by the enzymatic cycling
method of Matsumura and Miyachi (1980).
Oxidant Damage
Lipid peroxides were extracted from 0.5 g of nodules with 5 mL of 5% (w/v) metaphosphoric acid and 100 µL of 2% (w/v in
ethanol) butyl hydroxytoluene (Minotti and Aust, 1987). The extract was filtered through one layer of Miracloth and centrifuged at
15,000g for 20 min. An aliquot of the supernatant was
reacted with thiobarbituric acid at low pH and 95°C and cooled to
room temperature. The resulting thiobarbituric acid-malondialdehyde
adduct was extracted with 1-butanol and quantified by HPLC as described
in detail by Iturbe-Ormaetxe et al. (1998).
Protein carbonyl content was measured by derivatization with
2,4-dinitrophenyl-hydrazine as indicated by Levine et al. (1990) with
some modifications. Proteins were extracted from 0.5 g of nodules
with 5 mL of 100 mM potassium phosphate (pH 7.0), 0.1% (v/v) Triton X-100, 1 mM Na2EDTA, and
2.5 µg each of aprotinin and leupeptin to prevent proteolysis of
oxidized proteins during sample preparation. After precipitation of
possible contaminating nucleic acids in the samples with 1% (w/v)
streptomycin sulfate, an aliquot of 0.8 mL of the extracts was reacted
with 0.2 mL of 20 mM dinitrophenylhydrazine (in 2 M HCl), and another aliquot (control) with 0.2 mL of 2 M HCl for 1 h, with vigorous shaking every 10 to 15 min. Proteins were then precipitated with 10% (w/v) TCA, and the
pellet was washed four times with 1:1 (v/v) ethanol:ethyl acetate.
Precipitated proteins were solubilized in 6 M guanidine-HCl (pH 4.5) by incubation for 30 min with shaking. The insoluble material
was removed by centrifugation, and the absorbance of the hydrazones
(derivatized carbonyls) was measured at 370 nm (Levine et al., 1990).
To obtain more accurate results, the amount of protein to be analyzed
for carbonyl content was adjusted to 0.5 mg in all samples.
Light Microscopy Studies
Prior to immunodetection of relevant proteins at the light or
electron microscope levels, bean and pea nodule extracts were subjected
to western analysis (Cresswell et al., 1992). Polyclonal antibodies
used in this study were raised in rabbits against nitrogenase (universal antibody to the ferroprotein of nitrogenase; courtesy of Dr.
Paul Ludden, University of Wisconsin, Madison), APX (soybean nodule cytosol; Dalton et al., 1993, 1998), and ferritin (soybean seeds; courtesy of Dr. Elizabeth Theil, Children's Hospital Oakland Research Institute, Oakland, CA).
Nodules to be used for immunodetection of nitrogenase and APX were
fixed overnight in 2% (v/v) paraformaldehyde, 1.25%
(v/v) glutaraldehyde, and 50 mM PIPES (pH 7.2).
Fixed samples were dehydrated in an ethanol series, embedded in LR
White resin, and cut into 1-µm sections with a microtome (Reichert
Ultracut R, Leica, Deerfield, IL). For light microscopy
immunogold staining, slides were incubated overnight at room
temperature in a 1:50 dilution of antibodies raised against nitrogenase
or APX. For both antigens, the slides were incubated for 1 h with
secondary antibody consisting of affinity-purified goat anti-rabbit IgG
conjugated to colloidal gold (Auroprobe, Amersham) at a 1:40 dilution.
Silver enhancement solution (InsenSEM, Amersham) was applied at room
temperature and monitored for sufficient development (15-25 min)
following the protocols of the manufacturer. All sections were
counterstained with 0.5% (w/v) safranin in water for 60 s.
Representative nodule sections were photographed with a microscope
(Laborlux S, Leica) equipped with a camera (FE-2, Nikon).
For light microscopy immunofluorescence staining of APX, the secondary
antibody consisted of affinity-purified goat anti-rabbit antibodies
conjugated to a Cy3 fluorophore (excitation at 550 nm and emission at
570 nm; Jackson ImmunoResearch Laboratories, West Grove, PA) at a
dilution of 1:300 for 30 min. Sections were viewed with the light
microscope equipped with a rhodamine filter.
Electron Microscopy Studies
Ultrastructural studies and immunolocalization of APX and ferritin
at the electron microscopy level were carried out as indicated in
Dalton et al. (1993), except that the APX and ferritin antibodies were
used at dilutions of 1:500 and 1:25, respectively, and grids were
counterstained only with uranyl acetate. Nodule sections were obtained
with a Reichert Ultracut E microtome, and representative micrographs
for all treatments were taken with a transmission electron microscope
(model 900T, Zeiss) at 80 kV.
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RESULTS |
Nitrogenase Activity and Related Parameters
Bean plants were treated with 10 mM nitrate for up to
4 d to progressively induce nodule senescence, which was monitored
by measuring general markers of metabolic activity (Table
I). One day of nitrate application was
sufficient to inhibit the in vivo nitrogenase activity by 73% and to
increase the ODB resistance by 3-fold. At this stage, the total root
respiration and carbon cost of nitrogenase did not vary, but there were
significant increases in Lb and total soluble protein of nodules (Table
I). After 2 d with nitrate, the carbon cost of nitrogenase
increased 1.7-fold, whereas Lb and soluble protein returned to control
values. After 4 d there was a further increase in the carbon cost
of nitrogenase up to 2.4-fold and decreases of 53% and 31% in Lb and
soluble protein, respectively, relative to the control. An increase in rhizosphere O2 produced a recovery of nitrogenase
activity at all stages of the nitrate treatment. Maximum activities at
increased O2 concentrations relative to those at
21% O2 were 82%, 211%, 148%, and 130%
after 0, 1, 2, and 4 d, respectively (Table I).
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Table I.
Some general markers of metabolic activity in
senescing nodules of bean treated with nitrate and of pea treated with
darkness for 0, 1, 2, and 4 d
Means (n = 4-8) were compared by one-way analysis of
variance and the Duncan's range test. For each parameter and legume
species, means denoted by the same letter do not differ significantly
at P < 0.05.
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Dark treatment of pea plants led to a drastic inhibitory effect on
nodule activity. After only 1 d of dark, in vivo nitrogenase activity was almost abolished, whereas total root respiration declined
by 68% and the ODB resistance was enhanced by 10-fold (Table I).
However, there was no effect on the carbon cost of nitrogenase, and Lb
and soluble protein only decreased by 29% and 14%, respectively.
Prolongation of dark stress for 1 more d lowered Lb content by an
additional 63% but had no effect on total soluble protein. There were
no further significant changes after 4 d of darkness, except for
total root respiration, which decreased by 86% relative to control
(Table I). Increasing rhizosphere O2
concentrations above 21% caused a slight rise in nitrogenase activity
in pea after 1 d of dark, but did not induce any recovery of
activity after 2 and 4 d of continuous darkness.
Carbohydrates
Nitrate had considerably less effect on nodule carbohydrates than
dark stress. Treatment of bean plants with nitrate led to a progressive
decline in Glc, Suc, and starch, whereas Fru increased by 35% after
1 d and declined to 60% of the control value after 2 or 4 d
(Table II). The contents of Glc and Suc
decreased by 30% to 40% after 1 d of nitrate application, 47%
to 57% after 2 d, and 75% after 4 d. Starch was similarly
affected after 1 and 2 d with nitrate, but remained at 48% of
control after 4 d (Table II).
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Table II.
Carbohydrate content in senescing nodules of bean
treated with nitrate and of pea treated with darkness for 0, 1, 2, and
4 d
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In contrast, nodules of pea plants dark-treated for 1 d had lost
97% of their Suc content, along with 69% of Glc, 53% of Fru, and
74% of starch (Table II). The contents of Glc and starch remained essentially constant at this low level following prolongation of the
dark treatment to 2 or 4 d, whereas Fru declined by 85% relative
to control and Suc virtually disappeared after 4 d (Table II).
Antioxidant Enzymes
Nitrate application for 1 d had only a minor effect on
antioxidant enzyme activities from bean nodules, with the exception of
GR activity, which increased by 41% (Table
III). Following 2 d with nitrate,
only APX activity had decreased significantly with respect to control,
but after 4 d there was a general decline in activities of all
enzymes of the ascorbate-GSH cycle. The decreases ranged from
approximately 20% for DR and GR to 63% for APX. Catalase activity
responded quite differently, with no variation after 2 d of
nitrate supply and a 52% decline after 4 d (Table III).
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Table III.
Antioxidant enzyme activities in senescing nodules
of bean treated with nitrate and of pea treated with darkness for 0, 1, 2, and 4 d
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Likewise, after 1 d of dark stress, pea nodules did not show
apparent changes in antioxidant activities, but after 2 d there were decreases of 26% to 32% in APX and GR, and catalase activity increased by 31% (Table III). Following 4 d of dark treatment, there were major declines in the activities of the enzymes of the
ascorbate-GSH cycle: 40% to 50% for APX, GR, and MR, and 82% for DR.
In contrast, catalase activity exceeded the control value by 44%
(Table III).
Antioxidant Metabolites and Nucleotides
Some improvements of an HPLC method based on the derivatization of
thiols to form highly fluorescent adducts (Fahey and Newton, 1987)
allowed us to quantify separately GSH and hGSH in nodules (Table
IV). Control (untreated) bean nodules
contained 103 nmol GSH and 262 nmol hGSH g1
fresh weight, and pea nodules contained 698 nmol GSH and 90 nmol hGSH
g1 fresh weight. Therefore, GSH was the major
thiol (89%) in pea nodules, whereas hGSH predominated (72%) in bean
nodules. Assuming uniform distribution and 85% water content in
nodules, the total thiol tripeptide (GSH + hGSH) concentrations in bean
and pea nodules were estimated to be 0.4 and 0.9 mM,
respectively.
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Table IV.
Antioxidant metabolites and nucleotides in
senescing nodules of bean treated with nitrate and of pea treated with
darkness for 0, 1, 2, and 4 d
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As occurred with antioxidant enzyme activities, nitrate application for
1 or 2 d had only a very limited effect on the content of
antioxidant metabolites and pyridine nucleotides in bean nodules (Table
IV). After 1 d with nitrate, there were only minor or moderate changes in GSH (37% decrease), ascorbate (20% increase), and NADH and
NADP+ (18% decrease). The contents of other
nucleotides and hGSH remained essentially constant, as did the
proportion of the disulfide forms of GSH and hGSH. Following 2 d
with nitrate, both GSH and hGSH decreased by 28%, but this decrease
was not due to simple thiol oxidation, since 98% of total thiols were
still present in the reduced form. Only after 4 d was there a
significant but modest increase in the proportion of the disulfide
forms from 2% to 5%. At this stage, GSH and hGSH had declined by 64%
and 85%, respectively, and pyridine nucleotides, except NADPH, by 39%
to 56%. In contrast, the ascorbate content of nodules treated with
nitrate for 4 d was identical to that of untreated nodules (Table
IV).
The effect of prolonged darkness on pea nodule antioxidants and
nucleotides was also moderate after 1 d, with decreases in the
range of 26% to 37% for ascorbate, NAD+, and
NADH (Table IV). After 2 d, all parameters except the hGSH content
experienced substantial declines ranging from approximately 24% for
NADPH to 62% for NADH. In addition, the proportion of oxidized thiols
rose significantly, from 14% to 20%. After 4 d, ascorbate and
hGSH declined by approximately 60% and GSH by 92%. The nodule content
of pyridine nucleotides also decreased markedly, albeit to a lower
extent for NAD(H) than for NADP(H). Nevertheless, the
NAD+ to NADH and the NADP+
to NADPH ratios were kept at approximately 3 and 1, respectively, throughout the dark treatment.
Synthesis of GSH and hGSH
Because the nitrate and dark treatments led to major
declines of GSH and hGSH in nodules that could not be accounted
for by oxidation to the disulfide forms (Table IV), we decided to
investigate whether there is GSH and hGSH synthesis in nodules and, if
so, whether this can be impaired by the stress treatments. Optimization of previous HPLC methods enabled us to assay all the enzyme activities required for the synthesis of GSH and hGSH in nodules (Table
V), indicating that there is genuine
synthesis of thiol tripeptides in the nodule tissue. The first
committed step for the synthesis of both tripeptides is catalyzed by
ECS, and this activity was similar in control (untreated) bean and
pea nodules. The second step is thought to be catalyzed by specific
synthetases, either GSHS or hGSHS. Both activities were present in bean
and pea nodules, but hGSHS activity predominated in bean nodules,
whereas GSHS activity predominated in pea. This is in agreement with
the major thiol tripeptides found in the respective nodules (Table IV), which strongly suggests that the relative abundance of GSH and hGSH is
determined at least in part by the corresponding thiol tripeptide
synthetase activities.
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Table V.
Activities of enzymes involved in GSH and hGSH
synthesis in senescing nodules of bean treated with nitrate and of pea
treated with darkness for 0, 2, and 4 d
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Nitrate supply of bean plants for 2 d led to decreases of 40% in
ECS activity and 60% in GSHS activity, but had no effect on hGSHS
activity; prolongation of treatment up to 4 d led to a decline of
50% to 60% for all three activities of nodules (Table V). Placement
of pea plants in the dark for 2 d caused decreases of 40% in GSHS
and hGSHS activities of nodules; after 4 d, these activities had
decreased by 50% to 60%, whereas the ECS activity of nodules had
increased by 55% relative to control (Table V).
Oxidant Damage
The contents of lipid peroxides (malondialdehyde) and oxidatively
modified proteins (carbonyl groups) were used as markers of free
radical damage in nodules. The response of both parameters during
senescence was, however, distinctly different (Table
VI). Nitrate treatment of bean or dark
treatment of pea for 4 d caused decreases of 39% to 48% in the
content of lipid peroxides of nodules. In contrast, both nitrate and
prolonged darkness increased the amount of oxidized proteins in nodules
by approximately 30% (Table VI).
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Table VI.
Oxidant damage of lipids and proteins in senescing
nodules of bean treated with nitrate and of pea treated with darkness
for 0, 1, 2, and 4 d
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Immunolocalization of Nitrogenase and APX
Immunogold localization of nitrogenase with silver enhancement and
dark-field light microscopy indicated the presence of abundant protein
in the infected cells of control (untreated) bean (Fig. 1A) and pea (data not shown) nodules.
Little or no labeling occurred in nodules that had been exposed to
either of the stress treatments for 4 d (Fig. 1B). Using the same
technique but with bright-field light microscopy, APX protein was found
to be localized predominantly in the endodermis and adjacent cell
layers of the nodule parenchyma (inner cortex), as well as in the
infected zone of control bean nodules (Fig.
2A). Using immunofluorescence with the
secondary antibody conjugated to the fluorophore Cy3, a similar
distribution of APX protein was noticed in control pea nodules (Fig.
2C). Treatment with either nitrate or dark led to a substantial
decrease in labeling intensity in both bean and pea nodules (Fig. 2, B
and D), in agreement with the observed declines in enzyme activity
(Table III). As expected, bean and pea nodule sections in which rabbit
normal serum was used in place of the primary antibody showed only very
sparse background labeling.
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| Figure 1.
Immunolocalization of nitrogenase in bean nodules
using dark-field light microscopy. A, Control (untreated) nodules.
Bright spots in the infected zone of this section indicate the presence
of nitrogenase. B, Nodule after 4 d with nitrate. Note the almost
complete lack of staining in this section. NC, Nodule cortex; NP,
nodule parenchyma; INF, infected zone. Bars = 100 µm.
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| Figure 2.
Immunolocalization of APX in nodules of bean (A
and B, bright-field immunogold detection) and pea (C and D,
immunofluorescent Cy3 detection). A and C, Control (untreated) nodules
showing strong labeling in the infected zone and a prominent band in
the nodule parenchyma. B, Nodules after 4 d with nitrate. D,
Nodules after 4 d of dark stress. B and D show a marked decrease
in labeling in both the infected zone and nodule parenchyma. V,
Vascular bundle. Other symbols are as in the legend to Figure 2.
Bars = 100 µm.
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The pattern of immunolocalization of APX at the electron microscopy
level was similar to that observed at the light microscopy level. No
label was evident in negative controls (Fig.
3, A and B), whereas strong labeling was
present in the cytosol of the parenchyma and infected cells of control
bean and pea nodules (Fig. 3, C and D). Label was also noted in the
cytosol of interstitial cells (Fig. 4B),
over the symbiosomes, and occasionally over the mitochondria and
bacteroids (Fig. 4C). Immunolabeling of APX decreased in bean and pea
nodule tissue with nitrate and dark stress. The location of labeling
was not affected by either of the two treatments.
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| Figure 3.
Immunogold localization of APX in control
(untreated) nodules. A and B, Negative controls (normal rabbit
antiserum in place of APX antibody) of bean and pea nodules,
respectively, showing no label in the cytosol or organelles of infected
cells. C and D, Bean and pea nodule sections, respectively, showing APX
label over the cytosol of infected cells. b, Bacteroid;
w, cell wall; m, mitochondrion; p, plastid. The bar in A = 0.2 µm for
all panels.
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| Figure 4.
Ultrastructural changes and APX localization in
bean nodules after nitrate and dark treatments. A, Negative control
showing no label in the cytosol, organelles, or symbiosome of a control
(untreated) nodule. B and C, Control nodules showing APX localization
in uninfected and infected cells. Label is seen frequently in the
cytosol of both uninfected (B, arrowhead) and infected (C, arrowhead)
cells, over the symbiosome, and occasionally over mitochondria (C,
double arrowhead) and bacteroids (C, asterisk). D, Detail of infected
cell of the nodule after 2 d of nitrate treatment. Cells and
bacteroids are similar in appearance to controls. E, Detail of infected
cell of the nodule after 4 d with nitrate showing the absence of
symbiosome membrane, but little change in bacteroids or in
poly--hydroxybutyrate granules. F, Detail of infected cells of the
nodule after 1 d of dark stress showing disruption of the
cytoplasm (arrowhead). Symbiosome membranes are disrupted in both of
the cells shown. G, Infected cells of nodule after 2 d of dark
stress showing lesions in the symbiosome membrane and the cytoplasmic
disruption observed throughout the nodule. H, Nodule following 4 d
of dark stress showing general disruption of infected and uninfected
cells. However, most bacteroids remain intact. m, Mitochondrion; p,
plastid; px, peroxisome; and u, uninfected cell. Bars = 0.4 µm.
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Ultrastructural Studies
Ultrathin sections of representative nodules from nitrate-treated
bean and dark-treated pea were also examined by electron microscopy to
follow the structural changes occurring during senescence, and
complement the physiological and biochemical data. In addition, nodules
from dark-treated bean and nitrate-treated pea were processed in
parallel to allow for orthogonal comparisons.
Infected cells of control (untreated) bean nodules were densely packed
with symbiosomes, each enclosing one to four bacteroids. Bacteroids
contained abundant poly--hydroxybutyrate granules, and organelles of
the infected cell, including mitochondria and plastids, were confined
to the periphery of the cell. Bean nodules treated for 1 or 2 d
with nitrate essentially had features similar to control nodules (Fig.
4D). After 4 d of nitrate treatment, significant disruption of the
host cytoplasm and of symbiosome membranes was observed; however, there
were no evident changes in bacteroids or in poly--hydroxybutyrate
content (Fig. 4E).
Dark stress resulted in observable changes in bean nodules after only
1 d of treatment (Fig. 4F). Many infected cells, particularly toward the interior of the nodule, showed disrupted cytoplasm and
discontinuities in the symbiosome membrane. After 2 d of dark, breaks in the symbiosome membrane were more frequently observed, and
cytoplasmic breakdown of infected cells was evident (Fig. 4G). Cell
disruption was observed throughout the infected zone of the nodule
after 4 d of dark treatment and no intact symbiosomes were
evident; however, the integrity of bacteroids was not affected and
poly--hydroxybutyrate granules were comparable in appearance to
those observed in controls (Fig. 4H).
Control pea nodules contained abundant symbiosomes within the dense
host cytoplasm. No poly--hydroxybutyrate was observed within the
bacteroids (Fig. 5, A-C). Cell organelles were located at the
periphery of infected cells, and profiles of the rough ER and Golgi
apparatus were often apparent. One day of dark resulted in minimal
changes in infected cells (Fig. 5C).
Following 2 d of dark treatment, cytoplasmic disruption was
widespread, including cytoplasmic breakdown and lesions in the
symbiosome membranes (Fig. 5D). Bacteroids ranged in appearance from
those showing slight cytoplasmic disruption to others showing extensive
breakdown. After 4 d of dark, bacteroids were often misshapen,
with disrupted cytosplasm and symbiosome membranes (Fig. 5E). The host
cell cytoplasm was not discernible and host cell organelles were rarely
observed.
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| Figure 5.
Ultrastructural changes and APX localization in
pea nodules after nitrate and dark treatments. A, Negative control
showing no label in the cytosol, wall, or symbiosome of an untreated
nodule. B, Infected cell of untreated nodule showing distribution of
APX label over the cytosol and symbiosome. Label was also observed in
uninfected cells and occasionally over mitochondria. C, Infected cell
of nodule after 1 d of dark stress showing only minor disruption
of symbiosome membrane (arrowhead). D, Low-magnification micrograph of
cell after 2 d of dark stress showing the extent of cytoplasmic
disruption and bacteroids with various degrees of breakdown. E,
Infected cell of the nodule after 4 d of dark stress showing
disruption of symbiosome membranes and misshapen, disrupted bacteroids.
F and G, Infected cells of nodules after 2 d with nitrate showing
some cell plasmolysis (F, arrowhead), discontinuity in the symbiosome
membrane, and misshapen bacteroids (G, arrowhead). H, Infected cell of
the nodule after 4 d with nitrate showing the extensive
cytoplasmic and bacteroid degeneration characteristic of this stage. A,
B, C, E, F, and H, Bars = 0.4 µm. D and G, Bars = 2 µm.
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One day of nitrate treatment had little effect on the appearance of
infected cells in pea nodules. Cell plasmolysis and discontinuities in
the symbiosome membrane were apparent after 2 d with nitrate (Fig.
5F). Bacteroids with irregular membranes were occasionally observed
(Fig. 5G). Cytoplasmic and bacteroid degeneration was extensive
following 4 d of nitrate treatment, and cellular detail was
difficult to discern (Fig. 5H).
Ferritin Localization
Preliminary immunoblots of ferritin showed that the antibody
raised to soybean seed ferritin recognized a single protein band of 28 kD, characteristic of ferritin subunits, in bean nodule extracts. As
expected, a positive control consisting of a soluble extract of tobacco
leaves overexpressing soybean ferritin produced an identical band (Fig.
6). The immunoblots also revealed that ferritin accumulated in bean nodules in response to nitrate treatment (Fig. 6).
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| Figure 6.
Western analysis of ferritin in bean nodules using
chemiluminiscence detection. Lanes 0, 1, 2, and 4 were loaded with
soluble extracts (20 µg protein) from nodules treated with nitrate
for 0, 1, 2 and 4 d. Two extracts obtained independently were
loaded for each treatment. Lane T was loaded with a soluble extract (20 µg protein) of transgenic tobacco leaves overexpressing soybean
ferritin (Van Wuytswinkel et al., 1998), which served as a positive
control. In all lanes a single band at 28 kD, characteristic of
ferritin subunits, was observed.
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The soybean antibody, however, exhibited very poor reactivity with pea
ferritin (data not shown). Consequently, ferritin was only
immunolocalized in bean nodules. Control (untreated) bean nodule
sections showed little labeling for ferritin (Fig.
7A), but this was clearly visible after 2 or 4 d of nitrate treatment (Fig. 7, B-D), confirming blot
analysis. After 2 d with nitrate, ferritin was localized in the
plastids and amyloplasts of uninfected and infected cells, and
occasionally over the bacteroids. The heaviest labeling was observed in
the amyloplasts of the uninfected interstitial cells and of the
parenchyma cells (Fig. 7, B and C). In some sections, scattered arrays
or small clusters of ferritin particles could be seen in the
amyloplasts without the assistance of gold labeling (Fig. 7B). After
2 d (and especially after 4 d) of nitrate treatment, large
ferritin aggregates were easily observed in the amyloplasts (Fig. 7, C
and D). Quite often, these large deposits did not show any
immunolabeling, although it was observed in plastids or amyloplasts
within the same nodule sections (Fig. 7C), indicating that the antibody
was recognizing isolated ferritin particles and smaller ferritin
deposits.
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| Figure 7.
Electron micrographs of bean nodules showing
ferritin localization. A, Control (untreated) nodule section showing
scant deposition of gold label over infected and uninfected cells. B,
Detail of parenchyma cell after 2 d with nitrate showing ferritin
localization in plastids and amyloplasts. A linear array of ferritin
particles (arrowhead) is visible in the amyloplast. C, Detail of
parenchyma cell after 2 d with nitrate showing large ferritin
deposits not labeled with gold (arrowhead) as well as disperse labeling
of ferritin particles (double arrowhead) in the amyloplast. D,
Amyloplast from a parenchyma cell after 4 d with nitrate. A large
aggregate of ferritin (arrowhead) is present, but did not label with
gold. b, Bacteroid; p, plastid; ps, peribacteroid space; s, starch
grain; v, vacuole; w, cell wall. Bars = 0.4 µm.
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DISCUSSION |
In this work we have obtained simultaneously an array of
physiological, biochemical, and structural data for bean and pea nodules induced to senesce by treating the plants with nitrate or
prolonged darkness. This allowed us to monitor the progress of nodule
senescence in a controlled manner and to discern more readily between
the reversible and irreversible stages of stress application.
Nitrate-Induced Legume Nodule Senescence
Nitrate had a two-stage effect on bean nodules. In the first stage
(1-2 d), there were major declines in the in vivo nitrogenase activity
and increases in the ODB resistance and carbon cost of nitrogenase, but
only moderate or no changes in the nodule content of carbohydrates,
antioxidants, and pyridine nucleotides; furthermore, there were
significant increases in Lb, soluble protein, and ascorbate. At this
early stage, there were no detectable changes in the nodules at the
ultrastructural level. Therefore, it would appear that the decrease of
nitrogenase activity after 1 to 2 d may be attributed to
O2 limitation at the ODB level and not to
biochemical factors such as degradation of nitrogenase or Lb, oxidative
damage of cell components, or sugar deprivation of host cells or
bacteroids. However, the causal relationship between nitrogenase
activity and ODB operation cannot be fully examined by the present
data, and it is possible that nitrogenase activity is decreased by an as yet unknown mechanism that results in closure of the ODB.
O2 limitation through the ODB was confirmed by
the partial recovery of nitrogenase activity in nitrate-treated plants
upon increasing the concentration of rhizospheric
O2. This recovery, and the absence of structural
and biochemical damage, indicate that at this early stage the effect of
nitrate was still reversible. In contrast, in the second stage (4 d),
there was an almost complete loss of both nitrogenase activity and
protein, along with a marked decline in Lb, soluble protein, sugars,
antioxidants (except ascorbate), and nucleotides. There was also a
general deterioration of the nodule ultrastructure, in particular of
symbiosome membranes, and an accumulation of oxidized proteins.
This stage of nitrate inhibition would therefore appear to be
essentially irreversible. However, these conclusions await final
verification through recovery experiments.
The effects of nitrate on bean and pea nodules can be compared with the
limited information available for other legumes. In lupine and
clusterbean nodules, nitrogenase activity (assayed in both cases with a
closed system) was inhibited by 35% after 3 to 5 d with 20 mM nitrate, and structural degradation of lupine nodules
was only recognizable after 10 d (Lorenzo et al., 1990; Swaraj et
al., 1993). In bean and pea nodules, nitrogenase activity (assayed in
both cases with a flow-through gas system) was inhibited by
approximately 85% after only 2 d with 10 mM nitrate.
Structural data revealed differences in the progression of senescence
between pea and bean nodules. In pea nodules the plasmolysis of host
cells and the disruption of symbiosome membranes and bacteroids were already evident after 2 d, while in bean nodules, nitrate had little effect on the shape or poly--hydroxybutyrate content of bacteroids even after 4 d. These data suggest that pea nodules are
particularly sensitive to nitrate.
Dark-Induced Legume Nodule Senescence
Prolonged darkness severely affected pea nodule metabolism. After
only 1 d of dark there were major effects on total root respiration, ODB resistance, and carbon costs of nitrogenase. In
addition there were moderate decreases in Lb and in some antioxidants and nucleotides; however, the most affected parameters were by far the
in vivo nitrogenase activity and the nodule Suc content, which
decreased by 97%. The almost complete depletion of Suc, the major
carbon and energy source for host cell metabolism, along with the
substantial decreases in other carbohydrates, is the most likely cause
for the limitation of nitrogenase activity in dark-treated pea. This
conclusion is supported by immunoblots of nitrogenase showing no loss
in the protein after 1 d of dark in both bean (Gogorcena et al.,
1997) and pea (data not shown), and is also consistent with the finding
that isolated bacteroids from dark-treated soybean retained 50% of the
initial nitrogenase activity (Sarath et al., 1986) and fully recovered
upon addition of succinate (Carroll et al., 1987). After 2 d of
dark, pea nodules showed drastic decreases in Lb and carbohydrates,
moderate decreases of many antioxidants, accumulation of modified
proteins, and evident symptoms of structural deterioration. After
4 d of dark, there was a general collapse of metabolism and
extensive structural damage.
There are some significant differences in the response of other legume
nodules to dark stress. Thus, inhibition of nitrogenase activity was
much less dramatic in soybean (Sarath et al., 1986) and clusterbean
(Swaraj et al., 1994) than it was in bean (Gogorcena et al., 1997) and
pea (this work). Also, 2 d of dark had no significant effect on Lb
or total soluble protein in nodules of soybean (Pfeiffer et al., 1983;
Gordon et al., 1993), clusterbean (Swaraj et al., 1994), or bean
(Gogorcena et al., 1997). However, in pea nodules, the same treatment
caused a 74% decline in Lb but only a 14% decline in total soluble
protein, which is indicative of a relatively high sensitivity of pea Lb
to degradation compared with other cytosolic proteins. This conclusion
is consistent with an early observation that exposure of pea plants to
3 d of dark was sufficient to induce greening of nodules and
breakdown of 50% of total heme (Roponen, 1970). Because the pathway
for Lb degradation in vivo is largely unknown, it can only be
speculated that the rapid loss of pea Lb in dark-stressed nodules is
due to a particularly rapid activation or decompartmentation of
proteases located in the infected cells that display a high affinity
for Lb, especially at the acidic intracellular pH of senescing nodules
(Pladys et al., 1991).
Thiols, APX, and Ferritin in Senescent Legume Nodules
Three important antioxidants, GSH and APX, which are critical for
the operation of the ascorbate-GSH cycle, and ferritin, which is
critical for the control of the cellular concentration of catalytic
iron, have been studied in further detail to extend earlier studies on
the mechanism of stress-induced nodule senescence.
An enzymatic method (Griffith, 1980) was previously employed to
estimate the concentration of GSH in nodules. Because this method could
not distinguish between GSH and hGSH (Klapheck, 1988), and because it
was uncertain whether hGSH was present in nodules, we estimated the
concentration of total tripeptides in pea and bean nodules as 0.9 mM (Escuredo et al., 1996; Gogorcena et al., 1997). Using
HPLC, we found in this work that hGSH is present in nodules at variable
concentrations: 0.12 mM GSH and 0.31 mM hGSH
for bean nodules, and 0.82 mM GSH and 0.11 mM
hGSH for pea nodules. Thus, the enzymatic method tends to overestimate
the thiol content in plant tissues having primarily hGSH, because the
reaction of yeast GR with hGSH is faster than with GSH (Klapheck, 1988). For pea nodules, which mainly contain GSH, both methods yielded
similar results.
An obvious question raised in this work is why hGSH is the major thiol
in bean nodules. Although this cannot be answered at present, our
results show that the relative abundance of GSH and hGSH in nodules is
likely to be dictated by the presence of specific tripeptide
synthetases: hGSHS in bean nodules and GSHS in pea nodules. This
conclusion is reinforced by the partial purification of distinct
enzymes from pea (GSHS) and mungbean (hGSHS) leaves (Macnicol, 1987),
which contain only GSH and hGSH, respectively (Klapheck, 1988). The
fact that hGSH is the only thiol tripeptide present in the leaves of
some legumes also implies that the corresponding chloroplasts have a
functional ascorbate-hGSH pathway to avoid photooxidative damage, and
that hGSH and GSH share at least some antioxidative role in vivo.
The drastic decreases of the major thiols after 4 d of treatment
(85% hGSH in bean nodules and 92% GSH in pea nodules) cannot be
accounted for by oxidation to the disulfide forms or by the 50% to
60% decline in GSHS and hGSHS activities. Nor can it be ascribed to a
limitation of ECS activity, either directly (the activity only
declined by 60% in bean nodules and increased moderately in pea
nodules) or through the availability of Cys (the content of this
substrate decreased by 50%-60% in both bean and pea nodules after
4 d of treatment). The lack of evidence for a marked inhibition of
thiol synthesis, together with the extensive Lb degradation and
oxidative reactions taking place in stressed nodules that are manifest
by a loss of Lb heme and accumulation of oxidized proteins, may provide
a clue for the large decline in GSH and hGSH. These thiols may be
consumed by nonenzymatic reactions with activated oxygen species or by
enzymatic degradation, with the possible formation of mixed disulfides
between thiols and proteins (Rennenberg, 1995). These aspects of thiol
catabolism in nodules, and in plants in general, remain virtually
unexplored.
Another antioxidant, APX, is critical for the disposal of
H2O2 in nodules. The APX
protein was predominantly located to the parenchyma and infected zone,
confirming a previous report in which APX protein and mRNA were shown
to be enhanced in the parenchyma and infected cells of alfalfa nodules
(Dalton et al., 1998). In the same study, APX protein was also found to
be increased in the nodule parenchyma and infected zone of several
determinate nodules (Dalton et al., 1998). Our electron microscopy
studies corroborate the heterogeneous distribution of APX within
nodules and indicate that the protein is very abundant in the cytosol of infected cells. The observation of occasional labeling of APX in the
mitochondria would explain the detection of enzyme activity in purified
mitochondria from soybean nodules (Dalton et al., 1993). The activity
and content of APX protein largely decreased in senescent nodules,
which may cause a lowering in protection against
H2O2 generated by the
respiratory activity of the nodule parenchyma and infected cells.
Finally, we conducted studies to localize ferritin in bean nodules and
to determine the changes in ferritin content during senescence.
Ferritin was barely detectable in control nodules but accumulated in
nodules treated with nitrate for 2 or 4 d. The very low content of
ferritin protein found in mature, untreated bean nodules is in
agreement with the observation that ferritin accumulates in young
soybean nodules but declines in mature nodules, when iron storage is
apparently no longer required (Bergersen, 1963; Ragland and Theil,
1993). Ferritin was predominantly found in the plastids and amyloplasts
of interstitial cells in the infected zone and in the parenchyma cells.
The subcellular location of ferritin in bean nodules is fully
consistent with other reports showing ferritin particles in plastids
and amyloplasts of soybean, alfalfa, and lupine nodules (Bergersen,
1963; Lucas et al., 1998), in amyloplasts of soybean cell cultures
(Briat and Lobréaux, 1997), and in chloroplasts and other
plastids of several plants (Seckback, 1982).
The scattered arrays and large deposits of ferritin particles observed
after 2 or 4 d with nitrate closely resemble, respectively, the
F-2 and F-3 types described by Seckback (1982). The latter category is
defined as paracrystalline ferritin arrangements (sometimes including
small zones of crystalline structure), such as those observed in
plastids of iron-treated Xanthium without the assistance of
gold labeling. Our finding that the paracrystalline deposits of
ferritin-like material did not label as densely as would be expected
with the soybean antibody requires further investigation, but similar
ferritin structures, clearly immunolabeled, accumulate in the cortex of
senescing soybean and lupine nodules (Lucas et al., 1998).
Ferritin synthesis in plants is regulated by iron and is induced by
various adverse conditions, including iron overload, which lead to
oxidative stress (Briat and Lobréaux, 1997). Recent experiments with de-rooted maize plantlets have shown that
H2O2 induces ferritin mRNA
accumulation in the presence of low iron concentrations and that this
effect is prevented by pretreatment of plantlets with antioxidants,
indicating that the induction of ferritin gene expression in this
system requires an oxidative step (Briat and Lobréaux, 1997).
Based on these observations, our results showing ferritin protein
accumulation in senescent bean nodules may be interpreted as a response
to oxidative stress. This oxidative stress is evidenced by the
accumulation of damaged proteins in nitrate-treated bean nodules and is
probably a consequence of the lowering of antioxidant activities and
the release of catalytic iron from proteins (Becana et al., 1998).
Further work is needed to establish the mechanism for ferritin
induction in nitrate-treated bean nodules and to determine whether a
similar phenomenon occurs in other legume nodules under different types
of stress.
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FOOTNOTES |
1
This work was supported by grant nos. PB95-0091
and PB98-0522 from the Dirección General de Enseñanza
Superior e Investigación Científica (Ministry of
Education and Culture, Spain) to M.B., and by fellowships from the
Gobierno Vasco (M.A.M.), the European Union (I.I.-O.), and the Ministry
of Education and Culture (P.R.E., M.C.R., J.F.M.).
*
Corresponding author; e-mail becana{at}eead.csic.es; fax
34-976-575620.
Received April 16, 1999;
accepted June 2, 1999.
| |
ACKNOWLEDGMENTS |
We thank Shannon Joyner (Reed College), Caron James (Institute
of Grassland and Environmental Research), and Gloria Rodríguez (Estación Experimental de Aula Dei) for technical assistance. We
are considerably indebted to Tomas Ruiz-Argüeso (Universidad Politécnica de Madrid) for the Southern analysis of
hup genes in the two Rhizobium strains used in
this study, to Paul Ludden (University of Wisconsin, Madison)
for providing nitrogenase antibody, to Elizabeth Theil (Children's
Hospital Oakland Research Institute, Oakland, CA) and
Jean-François Briat (Université de Montpellier II,
France) for providing ferritin antibodies and help with western analysis, and to Chris Davitt and Valerie Lynch-Holm (Washington State
University, Pullman) for sectioning and embedding samples for
light microscopy.
| |
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Abstract
Introduction