Introduction
Tree carbon metabolism associated with photosynthesis, C allocation and
remobilization of C storage is well documented (Barbaroux et al.,
2003; Dickson, 1989), but tree nitrogen metabolism is less known.
Nevertheless, seasonal N cycling is a determinant of plant fitness in
perennials, particularly long-lived perennials such as forest trees
(Cooke and Weih, 2005). In early spring, trees' nitrogen
demand for growth can be satisfied either by uptake of external sources such
as ammonium, nitrate and organic N available from the soil
(Gessler et al., 1998a) or by remobilization of internal
stores (Bazot et al., 2013; Coleman and Chen, 1993; Cooke and Weih, 2005; El
Zein et al., 2011b; Gilson et al., 2014; Millard, 1996; Taylor, 1967). In many
species, N remobilization for growth in spring occurs before utilization of
N taken up by roots, typically during the 20–30 days before the roots
actively take up N. These species include deciduous species, such as
Quercus petraea (El Zein et al., 2011a), Malus domestica (Guak et al., 2003; Neilsen et al.,
2001), Populus trichocharpa (Millard et al., 2006), Prunus avium (Grassi et al., 2003),
Pyrus communis (Tagliavini et al., 1997) and Sorbus aucuparia (Millard et
al., 2001); marcescent/evergreen species, such as Nothofagus fusca (Stephens et al.,
2001) and coniferous evergreens, such as Picea sitchensis (Millard and Proe,
1993). In a few species (e.g. S. aucuparia), remobilization has completely finished
before any root uptake of N occurs, even if trees are supplied with an
adequate supply of mineral N in the soil. In contrast, other species have
been shown to begin taking up soil N through their roots concomitantly with
N remobilization. These include deciduous Juglans nigra × regia (Frak et al.,
2002), Pyrus communis (Tagliavini et al., 1997), Betula pendula
and evergreen Pinus sylvestris (Millard
et al., 2001). All of these studies were conducted on young trees and/or
under controlled conditions. Few studies have applied 15N-labelled
mineral fertilizer to larger, undisturbed trees growing in the field
(El Zein et al., 2011a), and even those only evaluated the
contribution of spring N uptake to leaf and twig growth, while the
contribution of stored N was indirectly estimated. However, in autumn, the
process of N storage (N translocation from leaves to sink compartments),
which starts concomitantly with leaf yellowing (Bazot et al., 2013), is
associated with a stimulation of soil nitrogen uptake (Gessler et al.,
1998b; Jordan et al., 2012; Kim et al., 2009). In the present study we
proposed to investigate the contribution of N storage and that of N taken up
from soil during autumn and spring to the development of new leaves of 20-year-old sessile oaks in the field, after budburst during the following
spring. Does soil N or foliar N contribute most to the storage of N
compounds in autumn? Does soil N or stored N contribute most to the
synthesis of new leaves in spring? Soil 15N labelling is a suitable
tool to quantify autumn and spring uptake of N by roots. Labelling of
foliage allows quantification of N remobilized from leaves to reserve
compartments. During three distinct labelling campaigns, 3 × 2 distinct
20-year-old sessile oaks received 15NH415NO3 applied to
their foliage (May) or on adjacent soil (September and March of the
following year). 15N partitioning in all tree–soil compartments, i.e.
leaves, twigs, trunk, roots, rhizospheric soil and microbial biomass, was
analysed regularly. The contribution of assimilated 15N to storage and
remobilization was investigated.
Materials and methods
Site description
The experiment was conducted in an area of 20-year-old naturally regenerated
oak in the Barbeau forest (48∘29′ N, 02∘47′ E), 60 km
southeast of Paris, France, at an elevation of 90 m on a gleyic luvisol. The
average air temperature is 10.5 ∘C and the annual rainfall in
this temperate location is 690 mm. Six 20-year-old sessile oaks (Quercus petraea L.) were
selected, their height ranged between 8 to 10 m and their average diameter
at breast height was 10 cm. In order to limit possible interference of root
cutting with nitrogen allocation, at least 5 months before labelling a
0.5–0.6 m deep trench was dug around each tree, then the trench was lined
with a polyethylene film and backfilled. All roots and root exudates inside
this perimeter therefore originated from the isolated tree, and were
contained in this trench volume. The area delimited by the trench was about
5 m2. The distance between each tree was at least 20 m.
15N pulse labelling
Three labelling campaigns were carried out: the first (L1) on the
foliage at the end of May (27 May 2009), the second (L2) on the soil at
the beginning of September (9 September 2009) and the third (L3) on the
soil the following March (20 March 2010). All labelling campaigns were
conducted on sunny days. Two oaks were labelled during each campaign: trees 1
and 2 during L1, trees 3 and 4 during L2 and trees 5 and 6 during
L3. Of buds showing leaf unfolding (Vitasse et al., 2009),
50 % occurred in those sessile oaks on 20 April 2010; this date was defined as
budburst. The L1 campaign consisted of homogenous spraying on all
foliage of 5 g 15NH415NO3 (98 at. %), i.e. 1.82 g of
15N, dissolved in 2.5 L distilled water. Prior to L1, the soil of the
surrounding trenches was protected with a plastic tarpaulin covering the
whole area of the trenched plot to avoid soil pollution with 15N. The
tarpaulin was sealed to the trunk at 50 cm height with Terostat-VII
(Teroson, Henkel, Germany). It remained on the soil for 2 weeks after
labelling. Before removing the plastic tarpaulin, crowns were sprayed with
distilled water in order to avoid any soil contamination after the
removing of the tarpaulin.
This first campaign aimed to label foliage and subsequently
the N reserves developed from the remobilization of leaf N the following autumn.
The L2 campaign consisted of homogenous spraying of 5g
15NH415NO3 (98 at. %), i.e. 1.82 g of 15N,
dissolved in 20 L distilled water on the soil of the trench plot of two
other selected oak trees (3 and 4). With this procedure, N reserves
developed from autumnal soil N uptake were expected to be labelled. The
third and last labelling campaign, L3, consisted of homogenous spraying
of 5 g 15NH415NO3 (98 at. %), i.e. 1.82 g of 15N,
dissolved in 20 L distilled water on the soil of the trench plot of trees 5
and 6, thus labelling their spring N uptake.
Sampling and analytical methods
Leaves, twigs, trunk phloem and xylem and soil monoliths (15 cm depth, very
few fine roots were present below 15 cm depth) of each labelled trees (1, 2,
3, 4, 5, 6) were sampled after labelling until the end of 2010 (Table 1). At
each sampling date 20 leaves and 20 twigs were collected randomly throughout
the crown. Sampling was always performed between 10:00 and 12:00 UTC. The
leaves were rinsed with distilled water to remove any excess 15N. At
each sampling date, two small disks of bark (14 mm diameter, 10 mm depth)
were collected at 1.3 m height using a corer. Thereafter, phloem and xylem
tissues were separated by hand with a cutter blade. The leaf mass per area
(LMA) was measured at each sampling date. Fine roots were hand picked from
the soil monoliths and washed with a 0.5 M CaCl2 isotonic solution.
Soil adhering to roots was removed with a brush and sieved at 2 mm. All
plant tissues and soil samples were brought to the laboratory in a cooler.
Plant tissues were lyophilized and ground to a fine powder with a ball mill
before analyses. At each sampling date, one aliquot of each plant powder (1 mg)
was transferred into tin capsules (Elemental Microanalysis, UK, 6 × 4 mm,
ref. D1006, BN/139877). On some dates (day after labelling (DAL) 1, 126,
337, 460 for leaves and twigs of L1; DAL 126, 337, 460 for roots of
L1; DAL 227 and 350 for leaves and twigs of L2; DAL 49 and 350 for
roots of L2 and DAL 40 and 166 for leaves, twigs and roots of L3),
four aliquots of powder were transferred into tin caps in order to test the
repeatability of the analysis. The total N concentration of plant and soil
samples was analysed by dry combustion using an N auto-analyser (Flash EA
1112 series, Thermofinnigan). 15N abundance was quantified in the same
plant and soil fine powder aliquots with a mass spectrometer (PDZ Europa,
University of Davis, Isotopes Facility, California).
Labelling characteristics and recovery of 15N administered in
each labelling campaign from the sampled compartments of each tree, on each
sampling occasion (DAL: days after labelling, JD: Julian day number).
Tree
1
2
3
4
5
6
Labelling date
2009/05/27
2009/05/27
2009/09/09
2009/09/09
2010/03/20
2010/03/20
DAL/JD
% of recovered 15N
DAL/JD
% of recovered 15N
DAL/JD
% of recovered 15N
Year 1
1/148
39
25
3/255
68
72
3/150
31
25
6/258
68
50
6/153
30
24
9/261
68
70
9/156
22
19
16/268
33
38
16/163
19
16
28/280
31
22
30/177
17
15
49/301
29
15
57/205
17
14
84/336
29
14
126/273
15
14
189/336
14
13
Year 2
318/98
8
7
208/98
24
14
20/98
65
28
337/118
11
13
227/118
12
10
40/118
63
40
358/139
10
13
247/139
16
20
61/139
16
14
370/152
14
14
260/152
22
21
74/152
20
25
397/180
11
10
287/180
38
18
102/180
20
25
460/244
13
11
350/244
13
12
166/244
18
21
509/293
7
5
399/293
10
8
215/293
11
21
Microbial N contents of fresh soil samples were determined using the
chloroform fumigation–extraction method (Vance et al., 1987).
Two fresh soil subsamples of 10 g were prepared. One subsample was fumigated
for 24 h with chloroform vapour, while the other was not fumigated. Nitrogen
extraction was performed using 50 mL of 0.5 M K2SO4 for 30 min
under vigorous shaking. The extracts (fumigated and not fumigated) were
filtered, then analysed for N content using an N analyser (TNM-1, Shimadzu,
Champs-sur-Marne, France). The microbial 15N abundance was estimated
using the same procedure, except that the extraction solution was 0.03 M of
K2SO4 in order to avoid any alteration of the mass spectrometer
with the K2SO4 salt during 15N analysis.
Calculations
All 15N enrichments were corrected for the background natural abundance
of this isotope, using control values determined in plants and soils just
before labelling. The seasonal variations of the natural 15N abundance
of each compartments were also followed throughout the season; those
variations were very weak, consequently we chose to use the
15N natural abundance of the labelled trees just before labelling. The
total weight of each analysed compartment (i.e. leaves, twigs, trunk phloem
and xylem and fine roots) was extrapolated from those of six equivalent
trees (same size and same diameter) grown on the same site and under the same
conditions. Those trees were felled as follows: two in October of the first
labelling year (2009), two in the following May (2010) and two the
following February (2011). Total leaf biomass was corrected according to the
LMA. All data were expressed as proportion of recovered 15nitrogen
(PRN) in a specific compartment using the following calculation Eq. (1):
PRN%=Q15NcompartmentMaxQ15N×100,
where Q15N was the quantity of 15N recovered from a compartment on
a specific date, and Max Q15N was the maximum quantity of 15N
recovered from all the sampled compartments during the experiment.
The % contribution of each 15N source (L1: leaves, L2:
autumn soil N, L3: spring soil N) to the 15N recovered in the
roots in autumn or in the leaves of the second year was determined according
to the following calculation Eq. (2):
%contribution15NL1,L2,L3=(Q15Ncompartment/MaxQ15N)L1,L2,L3Σ(Q15Ncompartment/MaxQ15N)L1,L2,L3×100.
Results
For each labelling, the two analysed trees displayed similar patterns of
total recovered 15N in each compartment (data not shown) and 15N
partitioning throughout the experiment. Moreover, the test of repeatability
of the analysis revealed very little variability in the 15N partitioning
at a specific date or in a specific compartment (Figs. 1, 2, 3). Consequently,
results were expressed as the mean of both trees (L1: 1 + 2; L2:
3 + 4, L3: 5 + 6).
Partitioning of recovered 15N (PRN%) from the sampled
compartments following the first labelling campaign, i.e. from 27 May 2009 to
20 October 2010. (a) Leaves
and
twigs ×, (b) phloem Δ, (c) fine roots
,
(d) rhizospheric soil
and microbial
biomass + (for those compartments the Y axis was adjusted to 1). DAL:
days after labelling. The two lines, continuous and dotted, correspond to
tree 1 and tree 2. Vertical bars indicate standard errors.
15N partitioning within the plant–soil system during the first
leafy season
After the foliar labelling in spring (L1, 27 May 2009)
The total balance for the administered 15N demonstrated maximum
recoveries of 15N within the plant–soil system of 32 %, 1 day after
leaf labelling. It decreased to 13.5 % of the administered 15N
recovered in the sampled compartments at the end of September (126 days
after labelling) (Table 1).
The PRN was at maximum in leaves (96 %, Fig. 1a) 1 day after L1, then
decreased continuously during the four following months (from 27 May to
30 September 2009, i.e. until the 126th day after labelling) with a
mean decrease of 80 % between these two dates (Fig. 1a). The same pattern
was observed in twigs, where the PRN decreased from 3 % on day 1 to
0.4 % on day 126 (Fig. 1a).
In the trunk phloem tissue and the fine roots, the PRN stayed relatively
stable or slightly increased until day 57 (24 July 2009). They then
increased until day 126 (30 September 2009), when they reached 4.75 % in
the phloem and 16 % in the roots (Fig. 1b, c). The PRN from the
rhizospheric soil and microbial biomass was less than 1 % (Fig. 1d).
During winter (2 December 2009; day 189) the PRN reached 18.5 % in fine
roots (Fig. 1c).
After the first soil labelling (L2, 9 September 2009)
The total balance for the administered 15N demonstrated maximum
recoveries within the plant–soil systems 3 days after L2 of 70 %.
By the end of October (49 days after labelling), recoveries from the sampled
compartments decreased to 22 % of the administered 15N (Table 1).
Three days after labelling, 3 % of the recovered 15N was present in
the fine roots (Fig. 2c). Nine days after labelling (18 September 2009),
the PRN showed that the majority of the 15N was recovered from
the soil, with 61 % of the 15N recovered from the rhizospheric soil
and 32.5 % from the microbial biomass (Fig. 2d). During the following 40
days (until 28 October 2009), the PRN from the soil decreased to 8.5 % in
the rhizospheric soil and 9.5 % in the microbial biomass (Fig. 2d). On the
same date, 6 % of the 15N was recovered from the fine roots (Fig. 2c).
Less than 1 % of the 15N was recovered from the phloem, xylem
and twigs (Fig. 2a, b). In December (day 84) the PRN from the soil was
similar to that of the previous date and 4 % of the 15N was recovered
from the fine roots (Fig. 2c, d).
Partitioning of recovered 15N (PRN%) from the sampled
compartments following the second labelling campaign, i.e. from 9 September
2009 to 20 October 2010. (a) Leaves
and twigs
×, (b) phloem Δ, (c) fine roots
,
(d) rhizospheric soil
and microbial
biomass +. DAL: days after labelling. The two lines for each category
(continuous and dotted) correspond to tree 3 and tree 4. Vertical bars
indicate standard errors.
15N partitioning within plant–soil system before and after
budburst
Almost one year after the first labelling (L1) and before budburst
(8 April 2010, 318 days after labelling), 7.5 % of the 15N were
recovered in the sampled compartments. Thereafter, recovery remained stable
at around 12 % until September (460 days after labelling, Table 1).
On 8 April 2010, i.e. 318 days after L1, 11.5 % of the recovered
15N was found in fine roots (Fig. 1c). Twigs contained 4.5 % of
recovered 15N (Fig. 1a), while phloem contained 4 % (Fig. 1b). Less
than 0.5 % of 15N was recovered from the rhizospheric soil and
microbial biomass (Fig. 1d).
Eight days after budburst (28 April, i.e. 337 days after L1), 25 % of
the recovered 15N was observed in new leaves. By 19 May this had
decreased to 17 % (Fig. 1a). On 28 April twigs contained 3.5 % of the
recovered 15N (Fig. 1a), phloem 4 % (Fig. 1b) and fine roots 10 %
(Fig. 1c). From then until September (i.e. 460 days after labelling), the
PRN from leaves remained relatively stable (22 %), whereas it
largely decreased in fine roots (0.35 %) (Fig. 1a, b, c). Less than
0.2 % of the total 15N recovered over the season was from the
rhizospheric soil and microbial biomass (Fig. 1d).
Just before budburst following the second labelling (L2, 8 April 2010,
208 days after labelling) 19 % of the administered 15N were recovered
from all the analysed compartments (Table 1). Most of it was from the
rhizospheric soil (14.5 %, Fig. 2d). The microbial biomass contained
9.5 % of the recovered 15N and the fine roots 2 % (Fig. 2d, c).
The rest of the 15N (less than 5 %) was distributed between the
twigs, trunk phloem and xylem (Fig. 2a, b). The same pattern was observed
8 days after budburst (227 days after labelling): most of 15N was
recovered from soil microbial biomass and rhizospheric soil (12 %,
Fig. 2d), 2.25 % was recovered from fine roots, 3.5 % of 15N was
recovered from phloem and xylem and only 0.5 % was recovered from new leaves
(Fig. 2a).
From 8 April (208 days after labelling) to 19 May (247 days after labelling and 30 days after budburst), the PRN decreased in soil microbial biomass and
rhizospheric soil (7 %), but increased in fine roots (9.5 %) (Fig. 2d,
c). A noticeable increase of the PRN from leaves was also observed at this
date (4.5 %, Fig. 2a). Thereafter, the PRN from soil microbial biomass and
fine roots decreased slightly from May 19 to June 28 (i.e. 247 to 287 days
after labelling), then remained stable until the end of August (Fig. 2d, c).
The PRN from leaves increased to 7 % in June (Fig. 2a).
For trees with soils that were labelled in spring (L3, 20 March 2010), the
maximum recovery of the administered 15N occurred 40 days later:
51.5 % from the sampled compartments. Recovery decreased thereafter and
stabilized at 19.5 % until autumn 2010 (Table 1).
Twenty days after labelling and before budburst, the soil microbial biomass
contained 44.5 % of the recovered 15N and the rhizospheric soil
39 % (Fig. 3d). The remaining 15N was mainly located in the roots
(2 % of recovered 15N, Fig. 3c). Eight days after budburst, the PRN was
quite similar: 61 % in microbial biomass and 32 % in rhizospheric soil
(Fig. 3d). 15N recovered from fine roots followed a pattern similar to
that observed on the previous sampling occasion (Fig. 3c). However, between
8 and 30 days after budburst (from 28 April to 19 May 2010, i.e. from 40 to
61 days after labelling), the PRN in microbial biomass and in rhizospheric
soil decreased sharply to 3.2 % (Fig. 3d). On that date, 17 % of the
15N was recovered from the fine roots (Fig. 3c) and 21.2 % from the
leaves (Fig. 3a). The PRN from leaves remained stable until the beginning of
June (74 days after labelling) (Fig. 3a). From that date until September the
PRN from leaves and fine roots declined slightly (Fig. 3a, c). The PRN from
microbial biomass decreased continuously throughout the season and reached
2.5 % in September (day 166 after labelling) (Fig. 3d).
Partitioning of recovered 15N (PRN%) from the sampled
compartments following the third labelling campaign, i.e. from 8 April 2010
to 20 October 2010. a Leaves and
twigs ×, (b) phloem Δ, (c) fine roots ,
(d) rhizospheric soil and microbial biomass +. DAL: days after labelling. The two lines for each
category (continuous and dotted) correspond to tree 5 and tree 6. Vertical
bars indicate standard errors.
Discussion
Efficiency of labelling
Isotope labelling experiments are technically challenging and as a
consequence are very scarce on trees growing in natural conditions. In this
paper, field labelling campaigns were conducted on 20-year-old naturally
regenerated oaks. For each campaign (only) two trees were labelled.
Nevertheless the similarity of the results between them suggests that the
observed 15N partitioning in soil and tree is a representative view of
the functioning of such systems
During the first labelling procedure (L1), a significant fraction of
the added 15NH415NO3 was incorporated into the leaves of
the sessile oaks. A significant proportion of the 15N was allocated to
the leaves: more than 90 % of the 15N was recovered from this
compartment. The total balance for the administered 15N demonstrated
maximum recoveries within the plant–soil systems of 32 %, 1 day after
leaf labelling. The remaining 15N was probably lost by leaf leaching.
However, soil protection with plastic tarpaulins avoided all contamination
of soil and roots as indicated by the 15N recovered in the belowground
compartments (Fig. 1d). Thereafter, the recovery of administered 15N
from the sampled compartments decreased to 14.5 %, probably due to
allocation of 15N to non-harvested compartments, such as old branches,
coarse roots or the inner part of the trunk. Indeed, data currently
available on woody plants show that nitrogen is re-translocated from leaves
to storage sites such as old branches, trunk or coarse roots
(Valenzuela Nunez et al., 2011; Bazot et al., 2013). The soil
15NH415NO3 labelling (L2) conducted in September
was also effective. Indeed, the total balance for the 15N applied to
the soil demonstrated maximum recoveries within the plant–soil systems of
70 %, 3 days after soil labelling. The rest of the 15N was most
probably lost by soil leaching (30 % of the 15N provide). Thereafter
the recovery of administered 15N from the harvested compartments
decreased to 22 %. As with the leaf-labelling experiment (L1), this
decrease was presumably due to allocation of 15N to non-harvested
compartments. Finally, the soil 15NH415NO3 labelling
carried out the following March (L3) was also effective, with maximum
recoveries within the plant–soil systems of 51.5 %, 40 days after soil
15N labelling. This recovery decreased to a mean of 19 % during the
rest of the season.
N dynamics in soil–tree systems during the first leafy season
Following the first labelling procedure, the 15N was quickly
incorporated into leaves; more than 90 % of the 15N applied was
accounted for in leaves 1 day after labelling. Thereafter this portion
decreased continuously throughout the season. The unaccounted-for fraction of the
15N had presumably been transferred to other compartments, including
those which were not sampled, i.e. branches and coarse roots.
This important foliar N remobilization was observed to continue in
leaf-labelled trees until yellowing, i.e. the end of September. Data
currently available on woody plants show that nitrogen is mainly
retranslocated from leaves to storage sites during the autumn (Coleman
and Chen, 1993; Cooke and Weih, 2005; Dong et al., 2002; Taylor, 1967), due to
the predominant role of leaf senescence in the constitution of N stores.
Leaf senescence leads to the breakdown of leaf proteins, the transfer of
their nitrogen to the perennial plant parts and the formation of N storage
compounds (vegetative storage proteins and amino acids) (Dong et al.,
2000; Tromp, 1983). In this study, a noticeable increase in the percentage of
recovered 15N in fine roots was observed on 30 September (16 %). This
compartment could be defined as a storage compartment in young sessile oaks.
Such an observation has been already reported for oaks of the same pole
stand (Gilson et al., 2014), and similar findings were reported
for field-grown adult peach trees by Tagliavini et al. (1997), being typical
of other young deciduous trees (Millard and Proe, 1991; Salaün et al.,
2005; Tromp and Ovaa, 1979; Wendler and Millard, 1996). On this date (end of
September), branches and coarse roots could also have contributed
significantly to N storage, as previously described (Bazot et al.,
2013).
Conceptual scheme representing percentage contributions of
15N (Eq. 2) from each labelling campaign (L1: white, L2:
light grey, L3: dark grey) in roots in the autumn and in new leaves in
the season following the first labelling campaign.
At the same time, root uptake can also contribute directly to storage, as
proposed by Millard (1996). Indeed, 49 days after labelled 15N had been
applied to surrounding soil (L2) in September, 5.75 % was recovered
from the trees' fine roots. It can be underlined that at the end of
September, foliage 15N made up 73 % of the 15N recovered in
roots, whereas soil 15N uptake contributed to 27 % of the 15N
recovered in roots (Eq. 2, Fig. 4). The soil N uptake in this period was
mainly recovered in the root system; there was little labelled N in the rest
of the trees. This is consistent with the results of Tagliavini et al. (1997)
and Jordan et al. (2012), who found a significant fraction of labelled N in
fine root samples of peach trees with 15N applied on soil
before fruit harvest in September.
Concomitantly with root N uptake for storage, notably in fine roots, a
strong immobilization of N in microbial biomass was observed. Indeed, on
October 7 (i.e. 28 days after labelling), when yellowing was well advanced,
12.5 % of the applied 15N was recovered in microbial biomass and
21.5 % in rhizospheric soil: there was a competition for soil N between
microbial N immobilization and reserve synthesis by root N uptake at that
time. This is consistent with the idea that soil microorganisms are strong
short-term competitors for soil N due to their high surface area to volume
ratio, wide spatial distribution in the soil and rapid growth rates
compared with plants roots (Hodge et al., 2000). Thereafter, root
N uptake was still efficient during late yellowing (between 7 and
28 October), since 15N recovered from the fine roots slightly increased
from 3.5 to 5.5 %, whereas the 15N that was recovered from microbial biomass
decreased from 12.5 to 10 %. This could be explained by microbial
mortality and turnover, which releases N to the soil, combined with the
capacity of plants to sequester N for longer (Barnard et al., 2006; Bloor
et al., 2009; Hodge et al., 2000).
After leaf fall, trees may have a significant capacity for nitrate uptake in
the fine roots in midwinter (i.e. in the absence of leaves), as already
shown in Japan oak (Ueda et al., 2010). However, in our case, N soil
uptake was limited by low soil temperature, which affected the
mineralization rate and root activity, since the 15N recovered from
roots slightly decreased between October 28 and December 2 (5.5 to 4 %)
and then declined to 1.75 % between 2 December and 8 April.
N dynamic in soil tree system the following spring
In April (before budburst), for trees with leaves labelled in the previous
year (L1), the most part of 15N was recovered in their roots
(11.5 %). On the other hand, at the same date, most of the labelled N
applied to soil in September (L2) was recovered from the rhizospheric
soil (14.5 %). When soil (and hence spring N uptake) was labelled
(L3) at the beginning of March, a month later most of the 15N was
recovered from microbial biomass and rhizospheric soil (81 %), but a small
proportion of 15N was recovered from the fine roots (1.5 %). The
latter demonstrated a small N uptake before budburst, as has previously been
observed in Japan oak (Ueda et al., 2010). This early N uptake from
the soil could be related to sessile oak's hydraulic properties. As a
ring-porous species, sessile oak achieves 30 % of its annual radial stem
growth before leaf expansion in spring (Breda and
Granier, 1996). Water flow pathways are then restored each spring before the
onset of transpiration (Breda and Granier, 1996). This
enables early root N uptake from soil as soon as a threshold soil
temperature is reached.
Eight days after budburst, most of the 15N applied to leaves (L1)
was recovered from new leaves (25.2 %) and new twigs (mean of 3.5 %).
This clearly underlined that a significant proportion of 15N used to
synthesize new leaves came from 15N stored during the previous autumn,
as shown for Ligustrum (Salaün et al., 2005). Moreover, this N
came from foliar N of the previous year, not from soil N uptake during the
previous autumn. Indeed, trees labelled the previous autumn on soil (L2)
showed a similar partitioning of 15N in leaves and twigs before budburst
(208 days after labelling) and 8 days after budburst (227 days after
labelling), there was no mobilization of 15N for the new leaves and
twigs synthesis for those trees. Less than 1 % of 15N taken up from
soil before budburst was recovered in leaves and twigs 8 days after budburst.
A distinction might be made between stored N sourced from leaves and that
sourced from soil, stored mainly in roots. N from leaves could be stored as
amino acids in branches, trunk and coarse roots, whereas N taken up from soil
could be stored in roots as NO3-. This N was not converted into amino
acids by glutamine synthetase/glutamate synthase enzymes during winter, most
probably due to low enzymatic activity in roots during winter (Bazot et al.,
2013). As a consequence, in the following spring trees first remobilized
easily, circulating forms of N and N to be stored nearer to demands. Indeed
in trees, NO3- is hardly transported to their leaves but rather
turned into amino acids in their roots (Morot-Gaudry, 1997). Indeed roots
were the main site of NO3- reduction (Gojon et al., 1991).
Consequently, soil 15N was not the main contributor to the synthesis of
new twigs and new leaves during the first 8 days after budburst.
Eight days after budburst, 95 % of new
leaves 15N came from 15N-labelled reserves, 2 % from soil labelled
the previous autumn and only 3 % from soil labelled in the current spring
(Eq. 2, Fig. 4). Previous studies have also found that N reserves contribute
significantly to leaf expansion in young trees: in white birch (Wendler and
Millard, 1996), sycamore maple
(Millard and Proe, 1991), Japan oak (Ueda et al.,
2009), pedunculate oak (Vizoso et al., 2008) and sessile oak
(El Zein et al., 2011a).
Considering trees with soil that had been labelled in autumn (L2), 8
days after budburst the proportion of recovered 15N in microbial
biomass decreased slightly, whereas it slightly increased in fine roots
compared to the previous sampling date. One can suppose that the increased
soil temperature and the first flux of C from plant to soil
(rhizodeposition)-stimulated microbial biomass turnover, making 15N
available for root uptake. Very little 15N was recovered from the other
compartments of the trees.
Soil N uptake became really effective between 8 and 30 days after budburst.
Indeed, whatever the date of the soil labelling (autumn or the current
spring), 30 days after budburst, a sharp decrease in 15N in the
microbial biomass was observed, depending on an increase of 15N in fine
roots and in young leaves. In 28 June (at leaf maturity), 40 % of the
15N recovered from leaves came from stored 15N, 10 % came from
15N applied to soil the previous autumn and 40 % came from 15N
applied on soil in March, 1 month before budburst (Eq. 2, Fig. 4).
This pattern of contribution was maintained throughout the season.
Similar findings have been reported for other species. For example,
20–30 % of shoot-leaf N was supplied by spring-applied fertilizer for
mature pear trees (Sanchez et al., 1990) and mature almond trees
(Weinbaum, 1984), while only 13 % of a solution of nitrate-N and
ammonium-N that was applied to soil contributed to total leaf N of apple trees
(Neilsen et al., 1997). Sorbus aucuparia had remobilized half the N from
storage before any was taken up by the roots (Millard et
al., 2001). Finally, there is a concomitant/concurrent remobilization and
uptake of N from the soil by some other species, as shown for Scots pine
(Millard et al., 2001) and walnut (Frak et
al., 2002).