Journal of Experimental Botany, Vol. 47, No. 301, pp. 1111-1118,
August 1996
Journal of
Experimental
Botany
Mobilization of nitrogen reserves during regrowth of
defoliated Trifolium repens L. and identification of
potential vegetative storage proteins
Nathalie Corre 1 , Valerie Bouchart, Alain Ourry and Jean Boucaud
Laboratoire de Physiologie et de Biochimie V6ge'tales, LA INRA, Institut de Recherche en Biologie Appliqude,
Universite, F-14032 Caen Cedex, France
Received 21 November 1995; Accepted 24 April 1996
Abstract
Introduction
Although it is well established that carbon reserves
contribute to shoot regrowth of leguminous forage
species, little information is available on nitrogen
reserves except in Medicago sativa L. and Trifolium
subterraneum L. In this study, reserves were labelled
with 15 N to demonstrate the mobilization of endogenous nitrogen from roots and stolons to regrowing
leaves and new stolons during 24 d of regrowth in
white clover [Trifolium repens L.). About 55% and 70%,
respectively, of the nitrogen contents of these organs
were mobilized to support the regrowth of leaves.
During the first 6 d, nitrogen in regrowing leaves came
mainly from N reserves of organs remaining after defoliation. After these first 6 d of regrowth, most of the
shoot nitrogen was derived from exogenous nitrogen
taken up while the contribution of nitrogen reserves
decreased. After defoliation, the buffer-soluble protein
content of roots and stolons decreased by 3 2 % during
the first 6 d of regrowth. To identify putative vegetative
storage proteins, soluble proteins were separated
using SDS-PAGE or two-dimensional electrophoresis.
One protein of 17.3 kDa in stolons and two proteins of
15 kDa in roots seemed to behave as vegetative storage proteins. These three polypeptides, initially found
at high concentrations, decreased in relative abundance to a large extent during early regrowth and then
were accumulated again in roots and stolons once
normal growth was re-established.
White clover (Trifolium repens L.) is the most important
forage legume in many temperate regions of the world
(Frame and Newbould, 1986), and is normally subjected
to varying degrees of defoliation either by grazing animals
or by mechanical harvesting. Its persistence in mixed
clover/grass swards depends largely on its ability to
regrow after the removal of some or all of its photosynthetic surface. Defoliation leads to a shortage of current
photosynthate supply and the plant must draw on a
limited supply of storage material to grow new leaves
(Dankwerts and Gordon, 1989). This results in the
decline, but not really the depletion of carbohydrate levels
in all remaining parts of the plant following defoliation.
Moreover, leaf removal decreases N 2 fixation within a
few hours (Ryle et al., 1985; Gordon et al., 1990),
decreases protein and leghaemoglobin contents of nodules
and changes nodule protein complements assessed by
polyacrylamide gel electrophoresis and autoradiography
following in vivo assimilation of 35S methionine (Gordon
et al., 1990).
It is now generally accepted that regrowth after defoliation of forage legumes requires N reserves to initiate
new shoot growth because of the strong decrease of
nitrogenase activity and/or soil N uptake induced by
shoot removal. For the past 10 years, attention has been
given to nitrogen reserve mobilization estimated by pulsechase 15N labelling in Trifolium subterraneum L. (Phillips
et al., 1983; Culvenor and Simpson, 1991) and Medicago
sativa L. (Kim et al., 1991, 1993). A mean value of 50%
of total nitrogen in roots and crowns is mobilized to meet
the N demand of shoots during early regrowth. In alfalfa,
it appears that regrowth following defoliation is linked
Key words: White clover, regrowth, 15N-labelled, vegetative storage proteins, electrophoresis.
1
To wtiom correspondence should be addressed. Fax: +33 31 45 53 60. E-mail: ourry®criuc.unicaen.fr
© Oxford University Press 1996
1112 Cone et al.
to the availability of N reserves in roots rather than that
of C reserves (Ourry et al., 1994). In grass and legume
forage species, amino-N seems to be the most readily
available form of N (Ourry et al., 1988; Lefevre et al.,
1991; Hendershot and Volenec, 19936) and protein-N the
largest storage form (Ourry et al., 1989a; Culvenor and
Simpson, 1991; Kim et al., 1991) to supply shoot growth
following defoliation. More recently, specific proteins
with molecular masses of 32, 19 and 15 kDa were identified as being the main components of buffer-soluble
proteins of alfalfa taproots, and may be vegetative storage
proteins (Hendershot and Volenec, 1993a). These proteins
accumulate during autumn and early winter in taproot
tissues and rapidly decline in spring. Moreover, the depletion of specific amino acids and certain buffer-soluble
proteins from taproots during regrowth of defoliated
alfalfa suggests that these N-pools are utilized as a source
of N during foliar regrowth after defoliation (Hendershot
and Volenec, 1993*).
The aims of this work were (i) to quantify by 15N
labelling the contributions of N reserves and external N
to shoot regrowth of nodulated Trifoliwn repens L. supplied with NH4NO3 following defoliation, and (ii) to
identify proteins of stolons and roots which are involved
in N storage.
Materials and methods
Plant material and culture
White clover seeds (Trifoliwn repens L. cv. Huia) were inoculated
with wild Rhizobium trifolii strains, and germinated in sand.
Ten seedlings were then transplanted into a 91 culture pot when
the primary leaves had developed. Plants were grown hydroponically in a continuously aerated nutrient solution containing
1 mM NH4NO3 as described previously by Kim et al. (1991).
The natural light (January to May, daylength from about
9-14 h) was supplemented with fluorescent 'phytor' tubes
(Claude GTE, Puteaux, France, 150 /xmol m~2 s"1 at canopy
height) for 16 h d"1. The thermoperiod was 23°C (day) and
18°C(night).
Experimental design and
15
N labelling
Plants were grown for 3 months, and the leaves were then
submitted to a first defoliation (day 90). Plants were defoliated
by removing all petioles and leaves while the end of each stolon
was tagged with adhesive tape to distinguish old (i.e. already
present on the day of defoliation) and new stolons (i.e.
appearing during regrowth) during further regrowth. They were
then grown for 24 d (first regrowth) with 1 mM NH4NO3 and
labelled (day 104) with 1 mM I5NH4I5NO3 (15N excess of
1.02%) dunng the last 10 d. They were again defoliated (day
114) and allowed to regrow for 24 d in the presence of 1 mM
14
NH414NO3. Plants for 15N labelling and protein analysis were
harvested the day of defoliation (day 114) and after 6, 10, 14,
and 24 d of regrowth (days 120, 124, 128, and 138). Plants used
for electrophoresis analysis were harvested on the day of
defoliation and after 2, 6, 9, 15, and 30 d of regrowth (days
116, 120, 123, 129, and 144); control plants were taken 60 d
(day 144) after the first defoliation. Harvested plants were
separated into organs remaining after defoliation: roots (including nodules) and old stolons already present on the day of
defoliation and regrowing organs: new stolons, and regrowing
petioles and leaf laminae. Samples were immediately frozen in
liquid nitrogen, freeze-dried and then ground to a fine powder.
Nitrogen isotopic analysis and calculation of nitrogen flows
Determination of 15N and N contents in samples was performed
by a continuous flow isotope mass spectrometer (Twentytwenty, Europa Scientific Ltd, Crewe, UK) linked to a C/N
analyser (Roboprep CN, Europa Scientific Ltd, Crewe, UK).
The calculations performed in this study are dependent on a
number of assumptions, many of them supported by other
works. Previous studies (Ourry et al., 1990) have shown that
no significant efflux of 14N (as ammonium or nitrate) from
ryegrass roots to the labelled nutrient solution occurred in the
hydroponic conditions used in this experiment (solution renewed
every 5 d). Therefore, if it is assumed that no significant 15N
efflux occurred under these conditions (15N labelling followed
by a chase with unlabelled N), then (i) any increase in 15N in
regrowing tissues during 24 d of regrowth will be derived from
15
N previously stored in the organs remaining after defoliation
and (ii) any increase in 14N in the plant will be derived from N
uptake and N2 fixation which occur at values close to natural
abundance. Moreover, if it is assumed that 10 d of labelling are
sufficient to obtain uniform isotope distribution within the
different plant organs and within the different organ N pools,
then any mobilization of "N will be proportional to total N
mobilization (15N + 14N). This last assumption is supported by
the fact that estimations of N mobilization during ryegrass
regrowth are similar whatever the method of calculation or the
15
N labelling used, i.e. pulse-chase (Ourry et al., 1988) or
continuous labelling (Ourry et al., 19896) during regrowth.
Considering the previously stated assumptions, the apparent
change (dN/dt) in N content in a plant organ during the second
regrowth is the difference between nitrogen inflow and outflow
from this organ during the time dt:
dN/dt =
(1)
- ^outflow = N , + d l - Nt
are
where Nt and yVt+dt
the N contents at time t and t + dt,
respectively.
Therefore, nitrogen inflow derived from N uptake and N2
fixation during dt can be calculated from 15N dilution as:
"Inflow = " t + dt x ( 1 ~Ex+
from
ujx.ke
lnd
fiuUon
(2)
where Et and Et+<it are atom% "N excess in the plant part
measured at time t and t + dt, respectively. N outflow from a
plant organ, corresponding to endogenous nitrogen remobilization during dt, can therefore be calculated from Equation 1:
^outflow — ^ t ~ J% + dl + ^in/low
(3)
Substitution from Equation 2 gives:
NOutnow = (Nt x Et-Ni+dt x £, +dt )/£,
(4)
If there is the assumption that N outflow from all remaining
organs after defoliation is uniformly labelled and that no
isotopic discrimination occurs during further distribution to
regrowing laminae, petioles and new stolons, then the amount
of N mobilized to these organs is directly proportional to their
15
N contents.
Mobilization of N reserves during regrowth of white clover
Extraction of soluble proteins
Soluble proteins were extracted from 200 mg freeze-dried
samples at 4°C with 7 ml of 50 mM TRIS-HC1 buffer (pH 7.5)
containing 2 mM PMSF, 10 >iM leupeptin and 200 mM DTT.
After centrifugation (1200 g, 15 min), nucleic acids in the
supernatant were precipitated using protamine sulphate
(1 mg ml" 1 ) for 15 min at 4°C. The pellet was discarded after
centrifugation (18 000 g, 5 min) and proteins in the supernatant were precipitated with cold (4°C) acetone 80% (v/v).
They were then dissolved in 1.5 ml extraction buffer containing
150 /*g DNase I (Sigma) and 37.5 /ig RNase A (Sigma
Chemicals). After 15 min at 4°C, soluble proteins were
separated in five subsamples and again precipitated using cold
(4°C) acetone 80% (v/v). The first pellet was then resuspended
in a 50 mM HEPES-NaOH buffer (pH 7.5) (O'Farrell, 1975),
and protein concentration estimated using the Bradford assay.
The 15N content was analysed by mass spectrometry as
previously described on a subsample further precipitated
with trichloroacetic acid and sodium deoxycholate (Peterson,
1983).
SDS-PAGE electrophoresis of soluble proteins
One pellet of soluble proteins was resuspended in Laemmli
(1970) buffer, denatured for 5 min at 100 °C and centrifuged
(12 000 g, 5 min). The SDS-PAGE electrophoresis used a 15%
duracryl running gel with a stacking gel containing 5.5%
acrylamide. Samples of 40 ^g proteins were loaded in wells.
Two wells were used for loading known molecular weight
proteins. The gels were then run for 2.5 h at a constant 500 V.
2-D electrophoresis of soluble proteins
One pellet of soluble proteins resuspended in O'Farrell (1975)
buffer was used to run 2-D SDS-PAGE gels, loaded with
200 jxg proteins, according to a modified procedure from
O'Farrell (1975) on an Investigator System (Millipore Corp.).
First-dimension isoelectric focusing was in a 4.1% acrylamide
tube gel containing 9.5 M urea, 2% (v/v) Triton X-100,
5 mM (3-[(3-cholamidopropyl)dimethylammonio]-l-propanesulphonate), and 2% (v/v) Millipore 2D optimized carrier
ampholytes (pH 3-10). After 2 h of prefocusing to 1500 V with
current limiting to 110 /xA per tube, protein samples (20 fA)
were loaded on the basic end of the tube gel and focused with
200 V per tube for 17.5 h. The gels were removed and preequilibrated for 2 min in a 375 mM TRIS-HC1 (pH 8.6) buffer
containing 3% (w/v) sodium dodecyl sulphate, and 50 mM
dithiothreitol. Separation of the isoelectro-focused proteins in
the second dimension was as for the one-dimensional gels using
a 15% duracryl running gel, run 5 h at 500 V.
1113
30 d later and on controls on two independent samples, each
corresponding to the harvest of 14 plants.
Results
Dry matter accumulation and partitioning of previously
accumulated 15/V
Following defoliation, dry weight accumulations were
greater for regrowing laminae and petioles than the dry
weight production of new stolons, which occurred mainly
between days 14 and 24 (Fig. 1A). The dry weights of
roots and stolons remaining after defoliation were
unchanged during the first 10 d and 6d, respectively,
following leaf removal (Fig. IB). During the last 14 d of
regrowth, only the root dry matter increased significantly.
Partitioning of previously accumulated 15N during 24 d
of regrowth on a 14N medium is given in Fig. 2. The
decreases in 15N contents of all remaining tissues following defoliation (Fig. 2B) corresponded to the increases in
15
N measured in regrowing laminae, petioles and new
stolons (Fig. 2A). The total amount of 15N in the plant
remained at 572 ±34 ^g 15N per plant throughout
regrowth, suggesting that there was no significant loss of
15
N to the medium, which might have interfered with
estimation of endogenous N flows. The main decreases
in 15N contents were in stolons which remained after
defoliation (-70%, from 250 to 75 ^g 15N plant" 1 ) while
in roots, 15N content decreased by 55% during the first
10 d. Regrowing laminae were a stronger sink than
regrowing petioles and new stolons since they received
about 62% of the remobilized 15N (Fig. 2A). About 94%
1000
Silver staining and analysis of gels
Gels were subsequently silver stained as described by Lopez
el al. (1991). Bidimensional gels were analysed using the
Millipore Bioimage computerized image analysis system. Gels
were scanned, and the individual staining intensity of each
polypeptide was then expressed as a percentage of gel total
staining intensity. For each gel, silver staining intensities of
unknown proteins were estimated using silver staining intensities
of known molecular weight and isoelectric point proteins added
to each gel in constant amounts. Electrophoresis was performed
on samples from the day of defoliation and 2, 6, 9, 15, and
0
6
12
18
24
Days of regrowth
Fig. 1. Changes in dry weight (mg plant"1) of regrowing tissues (A):
stolons (V), petioles (D) and leaf laminae (O) and of organs remaining
after defoliation (B): roots ( • ) and stolons (T) during 24 d of
regrowth. Vertical bars, when larger than symbols, indicate ±se for
1114
Corre et al.
200-
0
6
12
18
24
Days of regrowth
Fig. 2. Changes in 15N contents (jig plant"1) of regrowing tissues (A):
stolons (V ), petioles (D) and leaf laminae (O) and of organs remaining
after defoliation (B): roots ( • ) and stolons ( • ) during 24 d of
regrowth. Vertical bars, when larger than symbols, indicate ±se for
n = 4.
of 15 N mobilization occurred during the first 10 d of
regrowth.
Mobilization of endogenous nitrogen and translocation of
newly assimilated nitrogen
The amounts of total N mobilized were calculated from
Equation (4) given in Materials and methods, and are
presented in Fig. 3. About 71% of the total endogenous
nitrogen mobilized to regrowing organs (Fig. 3A)
30-
occurred during the first 2 weeks of regrowth. At this
time, laminae, petioles and new stolons received 19.3, 5.9
and 2.2 mg N plant" 1 from roots and stolons which
remained after defoliation. Nitrogen endogenous to roots
and old stolons accounted equally for the total N reserves
mobilized for regrowth (Fig. 3B).
Nitrogen used by regrowing organs was derived from
N reserves and external N (Fig. 4). Translocation of
newly assimilated nitrogen from the medium to regrowing
organs (Fig. 4) followed a different pattern to that of N
reserves. During the first 6 d of regrowth there was no
significant translocation of exogenous nitrogen into any
organ, despite the fact that nodulated plants were also
supplied with ammonium nitrate. Translocation of exogenous N from the medium to regrowing organs increased
significantly from day 6 (Fig. 4). Regrowth can be divided
into two different periods, characterized in terms of N
source. During the first 6 d, nitrogen of regrowing organs
(Fig. 4) came mainly from remobilization of N reserves,
while thereafter it also came from assimilation of external
N. About 67, 72 and 67% of laminae, petiole and new
stolon N, respectively, were derived from reserves 6 d
after defoliation, and 53, 45 and 18% after 14 d of
regrowth (Fig. 4). However, the majority of total reserves
(about 90%) was used for regrowth of leaves
(laminae + petioles), which therefore constituted the
stronger sinks.
Overall flows of N from reserves or from the medium
(as N 2 , NO3~ or N H / ) , during 24 d of regrowth, are
•
Exogenous N
N from reserves
A
20-
1a 10Af
Of
© 30-
E
£
Z
AB
1
20-
OJ)
E
10-
V
0
6
12
18
24
Days of regrowth
Fig. 3. N remobilization (mg N plant"1) to regrowing tissues (A):
stolons (V), petioles (H) and leaf laminae (C) from reserve of organs
remaining after defoliation (B): roots ( • ) and stolons (T) during 24 d
of regrowth. Vertical bars, when larger than symbols, indicate ±se for
*=4
0
6
12
18
Day» of regrowth
Fig. 4. Total nitrogen increments (mg N plant"1) in regrowing tissues:
leaf laminae (A), petioles (B) and stolons (C) during 24 d of regrowth
after defoliation and its origin as N from reserve or derived from
exogenous N (NOf, NH< and N 2 ). Vertical bars, when larger than
symbols, indicate ±se for n = 4.
Mobilization of N reserves during regrowth of white clover
1115
Mobilization of N from reserves
26.5±4.9
Translocation of N from the medium (NOj, NH4, N2)
6
12
18
24
Days of regrowth
Fig. 6. Soluble protein contents (A,mg g" 1 dry weight) and soluble
proteins l5N contents (B, ^g "N in soluble proteins g"' dry weight) in
organs remaining after defoliation during 24 d of regrowth: roots ( • )
and stolons (T). Vertical bars, when larger than symbols, indicate ±se
forn = 4.
Fig. 5. Mobilization of N from reserves and translocation of N derived
from the medium (NO3~, NH4+ and N2) during 24 d of regrowth after
defoliation. Each value ±se for n=4 is given in mg N plant"1.
summarized in Fig. 5. Leaf removal induced a large
remobilization of root and stolon N reserves. Whilst
regrowing laminae and petioles were the stronger sinks
for endogenous nitrogen reserves in defoliated plants,
new stolons received only 10% of mobilized N reserves.
In a way, this is not surprising as new stolons will act in
the mid-term as new storage organs. Nitrogen from the
medium derived from N2 fixation and/or NOf and
NH^" uptake, was translocated mainly to regrowing laminae (32%), roots (29%), remaining (16%) and regrowing
stolons (14%), and the rest to regrowing petioles.
Furthermore, when N flow from reserves or from the
medium were compared, it was clear that N mobilization
in roots was fully compensated for by exogenous N
translocation, while this was not the case for remaining
stolons. Moreover, after 24 d, the total N increase in
regrowing organs was 102.2 mg N plant"1, 39% being
derived from reserves.
Soluble protein contents and electrophoretic pattern
Soluble protein contents represented the largest N pool
in roots (32 mg total N g" 1 DW) and in stolons (30 mg
total N g~' DW) on the day of defoliation. If it is
assumed that soluble proteins contained 6.25% N, and
using data from Fig. 6A, then they may account for
approximately 77% and 83% of total N in roots and
stolons, respectively. Soluble protein contents were exam-
ined in organs remaining after defoliation (i.e. roots and
remaining stolons) over 24 d of regrowth (Fig. 6A). They
decreased during the first 6 d in roots and during the first
10 d in stolons by 40% of initial values. After this first
period, soluble protein contents increased to about 90%
of the value found on the day of defoliation in roots, and
to a value equal to the original for stolons. When the
stolon and root proteins were submitted to isotopic
analysis, the decrease in their 15N contents was apparently
very large (Fig. 6B). It was reduced by 57% and 66% of
initial 15N soluble protein contents in roots and stolons,
respectively, after 6 d of regrowth, and remained at a
steady-state level thereafter. This result suggests that the
magnitude of proteolysis was partly hidden by de novo
protein synthesis from unlabelled amino acids. Moreover,
when compared to the decrease of total 15N contents in
these tissues at the same time (Fig. 2), i.e. —32% and
41% in stolons and roots, respectively, this strongly
reinforces the role of soluble proteins as a storage
component.
Analysis of root and remaining stolon proteins by
SDS-PAGE electrophoresis (Plate 1A, B) and their relative contribution to total staining intensity per gel, was
used to detect putative storage proteins. The results
indicate that specific proteins decreased during the first
6 d of regrowth. In roots (Plate 1A), a band with molecular mass of 15 kDa (Plate 1A), was hydrolysed between
6 d and 9 d of regrowth. In stolons, a protein of 17.3 kDa
was also largely hydrolysed within 6 d of defoliation
(Plate IB). Analysis of roots and remaining stolons by
1116 Cone et al.
r
A
kDa
66.2
- •
4.8
kDa
IEF
6.7
kDa
- 66.2
.43
. 45
45-
:
w
O
31
- 31
a.
•>
.30
a
21.5 -
21.5
17.3 1
^ 17.3
20.1
15 <
A
0
26
9
15
Days of regrowth
3 0 C
A
A
.14.4
•
B
kDa
kDa
66.2'
66.2
• 45
45
A : SOLUBLE PROTEINS IN ROOTS
- •
IEF
• 31
3121.5
.21.5
417.3
14.4
17.3 ^
14.4
2
6
9
IS
Days of regrowth
30
20.1
Plate 1. SDS-of soluble proteins of Trifolium repens L. roots (A) or
stolons (B). Each well was loaded with 40 /*g soluble proteins except
two wells loaded with a constant amount of known molecular weights.
Putative vegetative storage proteins are identified with an arrow. The
position of molecular weight markers is indicated on both sides of the
gel. (C) Control plants non-defoliated.
two-dimensional electrophoresis was used to confirm and
quantify the previous results.
In roots (Plate 2A), two proteins with molecular mass
of 15 kDa, but differing by their isoelectric points, i.e. 6.6
and 7.0 (Plate 2A), were largely hydrolysed during the
first 6 d. Their staining intensities decreased by 99% and
81% of initial values, respectively (Table 1). In stolons, a
protein of 17.3 kDa with an isoelectric point of 5.3
(Plate 2B) was also partially hydrolysed, which was confirmed by a 60% decrease of its staining intensity during
the first 6 d following defoliation (Table 1). As with roots,
when the changes of soluble protein contents (Fig. 6A)
are considered, an estimation that 80% of the polypeptide
of 17.3 kDa underwent proteolytic degradation during
the first 6 d of regrowth can be made. This protein seems
14.4
B : SOLUBLE PROTEINS IN STOLONS
Plate 2. Two-dimensional gel electrophoresis of soluble proteins of
Trifolium repens L roots (A) or stolons (B) harvested on the day of
defoliation. Each gel was loaded with 200 ^g soluble protein. Putative
vegetative storage proteins are identified by arrows.
to accumulate to a large extent in stolons as it represented
about 22% of the total staining intensity per gel, on the
day of defoliation. This protein was apparently detected
in roots (Plate 2A), but followed a less intense hydrolysis
(Table 1).
These three putative vegetative storage proteins
(Plate 2) were then accumulated during the last 9 d of
regrowth to an amount similar to those found on the day
of defoliation (Plate 1).
Mobilization of N reserves during regrowth of white clover
Table 1. Changes in relative staining intensities (as a percentage
of total gel staining intensity) of 15 and 17.3 kDa proteins
estimated by image analysis of 2-D gels loaded with 200 fj.g
soluble proteins from roots and stolons during 24 d of regrowth
Each value is the mean of two independent replicates and the numbers
between brackets give the difference between the mean and each
replicate; n.d.: not detected, MW: molecular weight, IP: isoelectric point.
Localization
Stolons
Roots
MW
17.3
15
IP
5.3
6.6
7.0
Days
0
6
10
14
24
7.7 (±1.4)
3.0 (±0.5)
4.9 (±0.1)
6.3 (±0.4)
12.0 (±0.4)
7.7 (±2.1)
0.1 (±0.1)
n.d.
0.3 (±0.1)
8.9 (±1.2)
12.5 (±1.4)
2.3 (±0.1)
4.3 (±0.9)
5.0 (±1.0)
21.4 (±1.5)
17.3
5.3
4.7 (±1.3)
4.2 (±1.3)
3.5 (±0.3)
5.0 (±1.9)
9.8 (±0.7)
Discussion
After a single defoliation, N2 fixation by nodulated white
clover declined by more than 80% within 3 h, by nearly
100% by 24 h, and then recovered to its initial value 15 d
after defoliation (Gordon et al., 1990). The fact that
activities of enzymes involved in sucrose, carboxylic acid
and amino acid metabolism in nodules (Gordon and
Kessler, 1990) were all depressed following defoliation
suggests that the effect is more general than specific.
Moreover, these authors considered that the loss of
protein was unlikely to be due solely to the observed
increase in endopeptidase activity, which occurred only
after significant declines in nodule protein content. The
declines in protein content may also be related to changes
in the rate of protein synthesis (Gordon et al., 1990).
After an initial lag of 24 h after defoliation, specific
activities of alfalfa nodule glutamine synthetase, NADHglutamate synthase and NAD-glutamate dehydrogenase
also decreased (Groat and Vance, 1981), but less rapidly
than nitrogenase (Cralle and Heichel, 1981; Groat and
Vance, 1981; Ta et al, 1990; Denison et al., 1992) while
protease activity reached a maximum 7 d after harvest
(Vance et al., 1979). In non-fixing plants like perennial
ryegrass (Ourry et al., 1988; Macduff and Jackson, 1992)
and non-nodulated alfalfa (Kim et al., 1993), shoot
harvest decreased nitrogen uptake from the medium as
nitrate or as ammonium during the first week of regrowth.
The explanation for these observations is probably that
shoot regrowth had not advanced sufficiently for renewed
phosynthesis to support nodule metabolism, and also the
energetic requirement for root N uptake. Therefore,
mobilization of N reserves seems to be a prerequisite for
shoot regrowth in many species like Loliwn perenne
(Ourry et al., 1988), Bromus mollis (Phillips et al., 1983),
Medicago sativa (Kim et al., 1991, 1993) and Trifolium
subterraneum (Culvenor and Simpson, 1991).
In this study, it has been shown that nitrogen was
1117
mobilized equally from the roots and stolons of Trifolium
repens (Figs 3, 5) to provide about 50% of the nitrogen
required for regrowing tissues during the first 10 d after
defoliation (Fig. 4). Even with nodulated white clover
supplied with ammonium nitrate, the mobilization of
nitrogen still supplied the demands of regrowth, showing
that this process was largely independent of the form of
N supplied. The decline in soluble protein of 32% from
roots and stolons during the first 6 d (Fig. 6A) was similar
to that reported from the stubble of perennial ryegrass
(33%) 4d after defoliation (Ourry et al., 1988). In this
last species, the decrease in soluble protein contents was
correlated with an increase in endoproteolytic activities.
It can be hypothesized that the same process occurred in
roots and stolons of clover during early regrowth and
this can be further supported by the fact that defoliation
increases nodule endoproteolytic activities in clover
(Gordon et al., 1990). When expressed as a percentage
of the initial values, the decline in 15N in soluble proteins
(Fig. 6B: -57% in roots and -66% in stolons) observed
during the first 6 d of regrowth, which were larger than
total 15N contents (Fig. 2B: -40% in roots and -32% in
stolons) and the decreases in soluble protein level
(Fig. 6A: -41% in roots and -33% in stolons) strongly
suggests that components of this N pool may act as
storage proteins. The changes in the pattern of SDS-and
2-D electrophoresis of root and stolon proteins after
defoliation, show that a specific protein of 17 kDa in
stolons, and 2 polypeptides of 15 kDa in roots, all largely
abundant, followed a typical cycle of hydrolysis/synthesis
during regrowth and, therefore, were probably involved
in N mobilization and storage.
According to a recent review (Staswick, 1994), several
criteria may be used to define a protein as a vegetative
storage protein. Amongst them, are their relative abundance which must exceed 5% of total soluble proteins,
their transient accumulation followed by an ample
degradation. The three polypeptides of 17.3 or 15 kDa
found in stolons and roots (Plates 1, 2; Table 1) met these
criteria. The data did not indicate that they lack any
metabolic activity or structural role or that they are
specifically localized in the vacuole, which are other
criteria used by Staswick (1994) to define VSPs.
Furthermore, it has been shown that the degradation of
these three putative VSPs was concomitant to a large
mobilization of N between tissues (Figs 3, 4), and,
moreover, that their accumulation followed a seasonal
pattern (data not shown) with a peak of accumulation at
the beginning of winter and an ample spring degradation
as in woody species VSPs (Staswick, 1994). These latter
data give further evidence that these three polypeptides
act as VSPs. It should be noted that three vegetative
storage proteins of 32, 19 and 15 kDa were identified in
alfalfa taproots (Hendershot and Volenec, 1993ft), but it
is not known yet if they are related to putative storage
1118 Cone et al.
vegetative proteins of clover. However, their difference in
molecular weight support the hypothesis that they are
species-specific, which may not facilitate the elucidation
of their functional nature, if any.
The indication from the results presented here, and
from increased knowledge of metabolic changes occurring
after defoliation, is that N mobilization can be considered
as a general process for forage species, and probably
involves specific storage proteins. It is not known, however, if their availability at the beginning of regrowth can
be responsible for higher regrowth yield under optimal
conditions.
Future work will, therefore, be directed towards the
production of polyclonal antibodies raised against these
three polypeptides in order to screen genotypes differing
by their regrowth ability, over different seasons. Secondly,
immunolocalization of these polypeptides and their partial micro-sequencing will be used as a first step to assess
their functional nature and the control of their synthesis/
degradation.
Acknowledgement
We are grateful to Miss Micheline Meyer for her excellent help
for electrophoresis.
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