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Rhizodeposition of Organic C by Plant: Mechanisms
and Controls
Christophe Nguyen
Abstract During their life, plant roots release organic
compounds in their surrounding environment. This
process, named rhizodeposition, is of ecological im-
portance because (1) it is a loss of reduced C for the
plant, (2) it is an input flux for the organic C pool of
the soil and (3) it fuels the soil microflora, which is
involved in the great majority of the biological activ-
ity of soils such as the nutrient and pollutant cycling
or the dynamics of soil borne pathogens, for example.
The present review first examines the mechanisms by
which major rhizodeposits are released into the soil:
production of root cap cells, secretion of mucilage,
passive and controlled diffusion of root exudates. In a
second part, results from tracer studies (43 articles) are
analysed and values of C flux from the plant root into
the soil are summarized. In average, 17% of the net
C fixed by photosynthesis is lost by roots and recov-
ered as rhizosphere respiration (12%) and soil residues
(5%), which corresponds to 50% of the C exported by
shoots to belowground. Finally, the paper reviews ma-
jor factors that modify the partitioning of photoassimi-
lates to the soil: microorganisms, nitrogen, soil texture
and atmospheric CO2concentration.
Keywords Carbon rRhizodeposition rRhizosphere r
Tracer studies
C. Nguyen ()
UMR 1220 TCEM, INRA, 71 avenue Edouard Bourlaux,
F33883, Villenave d’Ornon, France
e-mail: Christophe.Nguyen@bordeaux.inra.fr
Résumé La rhizodéposition de C organique par les
plantes: mécanismes et contrôles Au cours de leur
vie, les racines des plantes libèrent des composants
organiques dans leur environnement proche. Ce pro-
cessus, nommé rhizodéposition, est d’importance
écologique car (1) c’est une perte de C réduit pour la
plante, (2) c’est une flux d’intrant pour la réserve en C
organique du sol et (3) il alimente la microflore du sol,
qui est impliquée dans la grande majorité de l’activité
biologique des sols tels que par exemple le cycle des
éléments nutritionnels et des polluants ou encore les
dynamiques des éléments pathogènes apportés par le
sol. La présente revue examine en premier lieu les
mécanismes par lesquels les rhizodépôts majeurs sont
libérés dans le sol: la production de cellules de la coiffe
racinaire, la sécrétion de mucilage, la diffusion passive
et contrôlée d’exudats racinaires. En second lieu, les
résultats de traceurs (43 articles) sont analysées et les
valeurs de flux de C allant de la racine de la plante
au sol sont synthétisées. En moyenne, 17% du C net
fixé par la photosynthèse est perdu par les racines et il
est restitué dans la respiration de la rhizosphère (12%)
et dans les résidus de sol (5%), ce qui correspond à
50% du C exporté par les pousses vers le sous-sol. En-
fin, l’article répertorie les facteurs principaux qui mod-
ifient la répartition des photoassimilats vers le sol: mi-
croorganismes, azote, texture du sol et la concentration
en CO2de l’atmosphère du sol.
Mots clés Carbone rRhizodéposition rRhizosphère
rTraçage
E. Lichtfouse et al. (eds.), Sustainable Agriculture, DOI 10.1007/978-90-481-2666-8_9, 97
c
Springer Science+Business Media B.V. - EDP Sciences 2009. Reprinted with permission of EDP
Sciences from Nguyen, Agronomie 23 (2003) 375–396. DOI: 10.1051/agro:2003011
98 C. Nguyen
1 Introduction
During their life, plant roots release organic com-
pounds in their surrounding environment. This phe-
nomenon is now being studied for more than one cen-
tury. Indeed, the very complete book of Krasil’nikov
(1961), reports that root excretion was first evidenced
in 1894 by Dyer, who observed the excretion of acidic
compounds from the roots of plants. Then, numerous
workers identified sugars, organic and amino acids and
other compounds in the nutrient solution in which dif-
ferent plants were grown. Krasil’nikov (1961) reported
that as early as in 1927, Minima observed that root ex-
cretions of organic compounds by lupine, bean, corn,
barley, oat, and buckwheat cultivated in Knop’s nutri-
ent solution, were maximum during the fourth week
of growth. Afterwards, these excretions decreased and
stopped altogether with plant growth. In the beginning
of the twentieth century, it was already estimated that
root-released compounds yielded 0.6–27% of the plant
dry weight and studies also demonstrated that greater
amounts of substances could be obtained if the nutrient
solution was replaced (Krasil’nikov,1961).
The release of organic compounds by living plant
roots referred to as rhizo-deposition (Shamoot et al.,
1968) is a process of major importance that is still sub-
ject of investigations for several reasons. Firstly, rhi-
zodeposition is an input of organic C into the soil.
The soil is the second largest C compartment .1:5
1012 tC/after oceans .3:8 1013 tC/and before at-
mosphere .7:5 1011 tC/and plant biomass .5:6
1011 tC/(estimates from (Schlesinger and Andrews,
2000)). Each year, it is estimated that 7:5 1010 tofC
return to the atmosphere due to soil respiration. Con-
sidering that in average, shoots export to belowground
about half of the C fixed by photosynthesis (Lambers,
1987), it is of major importance to determine how
much of this flux enters the soil organic C pool. This
is particularly relevant if the soil is expected to seques-
trate C in response to elevation of atmospheric CO2.
Secondly, rhizodeposition represents a loss of en-
ergy for the plant. At first sight, the release of organic
C from roots into the soil might figure as a lost pool
of reduced C that does not contribute to dry matter
production. However, it is well established that rhi-
zodeposits stimulate the biological activity in the rhi-
zosphere, which have important positive feedbacks for
the plant such as enhancing of nutrient availability for
instance (Jones and Darrah,1996). However, we still
have no idea on the efficiency of rhizodeposition. In
other words, would a plant gain extra advantages in
terms of mineral nutrition for example, if it would de-
posit more C into the soil? The response to this ques-
tion is fundamental with respects to outlooks aimed at
engineering the rhizosphere.
In the past decades, many studies have focussed on
rhizodeposition. Authors have concentrated on deter-
mination of C flows from plant roots to soil and on
factors that affected them. Results have been reviewed
at regular interval (Grayston et al.,1996;Hale and
Moore,1979;Kuzyakov and Domanski,2000;Rovira,
1969;Whipps,1990). Briefly, these articles outlined
that: (1) plant roots are able to release a wide range
of organic compounds, (2) there is a great degree of
uncertainty about the amounts and the quality of or-
ganic C deposited in soil conditions; this comes from
the major difficulty to estimate root-derived C in the
presence of microorganisms that rapidly assimilate rhi-
zodeposits, (3) in soil conditions, many factors are as-
sumed to alter both the amount and the nature of the
C compounds released from roots but little is known
about how these factors operate.
Our knowledge of rhizodeposition is too much in-
complete. As a result, the effective outputs of re-
search on rhizodeposition are lacking despite virtual
outputs are potentially numerous such as manipulating
C flow to the rhizosphere to alter the microbial dynam-
ics and the related processes (nutrient cycling, organic
matter dynamics, pollutants bioavailability, soil-borne
pathogen and inoculants dynamics, etc.).
Consequently, two areas of investigations could be
suggested. On one hand, new methodologies have to
be developed to obtain more reliable estimates of rhi-
zodeposition under various environmental conditions.
On the other hand, if a major goal is to manipulate rhi-
zosphere processes through plant ecophysiology and
through the quantity and the quality of rhizodeposits,
it is necessary to obtain more information about the
different mechanisms by which C is lost by roots as
well as their regulations by plant genetic and by envi-
ronment. The present article concentrates on that latter
point. It first reviews literature related to the mecha-
nisms by which major rhizodeposits (in terms of quan-
tity) are deposited into the soil: sloughing-off of root
cap cells, secretion of mucilage, passive diffusion of
root solutes (exudation) and senescence of epidermal
and cortical cells. The article then examines tracer
studies .14C/to summarize the main factors that are
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 99
assumed to affect rhizodeposition. It is attempted to re-
late their effects to the aforementioned mechanisms of
C release from roots. Finally, some outlooks are pro-
posed for future investigations on rhizodeposition.
2 Mechanisms of Release of Organic
C from Living Roots
2.1 Sloughing off of Root Border Cells
Apical meristems of plant roots are covered by a group
of cells arranged in layers, the root cap, the surface of
which sloughs-off as the root tip wends its way through
the soil (Barlow,1975). In mature branched roots, the
entire cap itself can be lost as the results of pathogen
attacks or as part of a normal developmental process
as it was observed in field-grown maize (Varney and
McCully,1991). The cap initials generate cells that are
displaced from the inner zone towards the periphery
of the cap where they slough-off. During their transit
through the cap, the cells first differentiate into stato-
cytes, i.e. gravity-perceiving cells and then into cells
able to secrete mucilage (Sievers and Hensel,1991).
The separation of cells from the periphery of the cap
can easily be observed under a microscope for numer-
ous plant species. In field-grown maize, the detached
cells were found alive at some distance from the root
tip (Vermeer and McCully,1982), which indicates that
border cells are still viable several days after their sep-
aration from the root. Among plants belonging to ten
families, the viability of border cells after they sepa-
rate from the root was demonstrated to be of 90% or
higher in most cases except in the Compositae sun-
flower and Zinnia for which most of the border cells
are dead when they detached from the cap (Hawes,
1990). Furthermore, in pea, detached cap cells exhibit
different gene expression than that of attached cap cells
(Brigham et al.,1995). It is suggested that they play
a significant role in engineering the rhizosphere ecol-
ogy (Hawes et al.,1998) and therefore, the term border
cells was proposed instead of the original denomina-
tion “slough off cap cells” (Hawes,1990). The sug-
gested functions of root border cells are numerous:
decrease of frictional resistance experienced by root
tips (Bengough and McKenzie,1997), regulation of
microbial populations in the rhizosphere by attracting
pathogens and preventing them from damaging root
meristem and by promoting growth gene expression
in symbiotic microorganisms (Hawes,1990;Hawes
et al.,1998,2000;Zhao et al.,2000), protection against
heavy metal toxicity such as aluminium (Miyasaka and
Hawes,2001).
In maize seedlings, the number of cells in the cap
ranges from 3,900 to 20,900 (Clowes,1976). It de-
creases with root age due to the reduction in the width
of root apices (Clowes and Wadekar,1988). In labo-
ratory experiments, the cap removed artificially is re-
generated in 1–9 days (Barlow,1975;Clowes,1976;
Sievers and Hensel,1991). The maximum number of
cells released daily from the cap is very variable rang-
ing from a dozen in tobacco to more than 10,000 for
cotton and pine but it is conserved at the species level
(Hawes et al.,2000). In maize, the daily production of
cap cells increases from 356 cells day1at 15ıCtoa
maximum of 3,608 cells day1at 25ıC and declines to
851 cells day1at 35ıC(Clowes and Wadekar,1988).
The production of border cells by roots growing in soil
is poorly understood. In laboratory experiments, it has
been demonstrated that environmental conditions ex-
perienced by root tips strongly influence border cell
production. For example, atmosphere with high CO2
and low O2partial pressure inhibits border cell separa-
tion in pea during germination whereas later in devel-
opment, it increases the total number of border cells
that accumulate over time (Zhao et al.,2000). The
mechanical impedance experienced by maize roots
creates friction that is decreased by the sloughing
of root cap cell (Bengough and McKenzie,1997).
Consequently, in maize seedling grown in compacted
sand the number of shed cap cells increases exponen-
tially with the penetration resistance from 1,900 cells
day1(56 cells mm1root elongation) for loose sand
(resistance to penetration: 0.29 MPa) to 3,200 cells
day1(750 cells mm1root elongation) for compacted
sand (resistance to penetration: 5.2 MPa) (Iijima et al.,
2000) (Table 1). The authors estimated that this corre-
sponded to 1.5 and 2:6 gC per day, respectively.
There are evidences that the number of border
cells is also controlled at the genetic level. In pea,
the separation of cells from the cap has been shown
to be closely correlated to the expression of an in-
ducible gene coding for a pectinmethylesterase, which
is thought to solubilize cell wall polymer (Wen
and Hawes,1999). Furthermore, there are evidences
that cap cells synthesized a factor that accumulates
100 C. Nguyen
Table 1 Production of root cap cells and mucilage by roots of different plant species
References Plant Nature of C Amount Units Comment
Iijima et al. (2000)Zea mays Root cap cells 1,900 cells/day Seedling grown in sand:
resistance to
penetration D
0.3 MPa. For
calculations, the root
cap cell is considered
as a cylinder with a
length of 80 manda
diameter of 21 m, a
density of 1g=cm3
and a dry matter/fresh
matter ratio of 0.072
(Iijima et al.,2000)
1.52 g C/day/root
Root cap cells 3,200 cells/day Same conditions as above
except the resistance
to penetration D
5.2 MPa
2.56 g C/day/root
Newman (1985)Zea mays Root cap cells 7 gDM/mgDM
of root
growth
2.8 gC/mgDMof
root growth Calculated assuming a C
content of root cap
cells of 40%
Convulvus arvensis Root cap cells 4 gDM/mgDM
root growth
Hawes et al. (2000)Pinus gossypium Root cap cells 10,000 cells/day
McLeod (1976)Vicia faba Root cap cells 420–636 cells/day
Chaboud and
Rougier (1991)Zea mays Mucilage 34 gDM/mgDM
root growth
Vancura et al. (1977)Zea mays Mucilage 11–17 gDM/mgDM
root growth Growth in axenic nutrient
solution for 28 days
Triticum aestivum Mucilage 29–47 gDM/mgDM
root growth Growth in axenic nutrient
solution for 25 days
Bowen and Rovira
(1973)Triticum aestivum Root cap
cells C
mucilage
3.2–6.4 gDM/mgDM
root growth
Samsevitch (1965)Triticum aestivum Root cap
cells C
mucilage
700 m3=ha Calculated from the size
of the droplet at the
root tip
Zea mays Root cap
cells C
mucilage
1,250 m3=ha Calculated from the size
of the droplet at the
root tip
Griffin et al. (1976)Arachis hypogea Root cap
cells C
mucilage
0.13–0.27 mg MS/plant/day Growth in axenic nutrient
solution for 2 weeks
Root cap
cells C
mucilage
0.15 % of root C
extracellularly and inhibits the production of new cells
by the cap meristem without inhibiting cell mitosis in
the root apical meristem (Hawes et al.,1998). Hence,
the cap turnover is stopped unless the factor is diluted
or unless cells from the periphery are shed. Therefore,
in soil, it can be assumed that the production of cap
cells is favoured on one hand by rain, irrigation and the
soil microporosity, which all facilitate the diffusion of
the inhibitor away from the cap and on the other hand,
by frictional forces that shed the cells from the root tip.
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 101
2.2 Secretion of Mucilage by Roots
A mucilaginous layer has been frequently observed on
the root surface of many plants (Oades,1978) and more
particularly at the root tip where it can form a droplet
in the presence of water (Samsevitch,1965). There
is no clear evidence that the epidermis and the root
hairs secrete mucilage (Peterson and Farquhar,1996).
In Sorghum, Werker and Kislev (1978) reported small
drops of mucilage secreted by root hairs in addition to
a fibrillar mucilaginous layer secreted by the epider-
mal cells. However, the mucilaginous layer observed
on these parts of roots may derived from the mucilage
secreted by the root cap (Vermeer and McCully,1982),
from the degradation of epidermal cell walls (Foster,
1982) or may be synthesised by rhizosphere microor-
ganisms (Rovira et al.,1979). However, for most of the
plants examined, the mucilage is secreted by the outer
layers of the cap cells (Paull and Jones,1975;Rougier,
1981) and it can be seen at the root tip of several plants
(Miki et al.,1980).
The mucilage is composed of polymerised sug-
ars and of up to 6% proteins (Bacic et al.,1987;
Rougier,1981). The major sugars identified are arabi-
nose, galactose, fucose, glucose, xylose (Bacic et al.,
1987;Knee et al.,2001). In maize, the root cap
polysaccharide has a molecular weight greater than
2:106daltons, a density of 1:63 gcm
3(Paulletal.,
1975), a C content of 39% and a C/N ratio of 64 (Mary
et al.,1993).
The initiation of mucilage synthesis takes place in
the endoplasmic reticulum and completes in the Golgi
saccules. The slime is transported to the plasmalemma
by the Golgi vesicles. The mucilage is discharged
between the plasmalemma and the cell wall by exo-
cytosis (Morre et al.,1967;Rougier,1981). All these
processes are energy-dependant. The passage through
the cell wall is not systematic and the mucilage can ac-
cumulate at the inner wall surface. It is assumed that
if both the degree of hydratation of the mucilage and
the cell tugor are sufficient, the slime moves passively
through the cell wall and forms a droplet at the root tip
(Morre et al.,1967). The passage trough the cell wall
is probably due to an increase in the permeability of
the middle lamella (Lynch and Staehelin,1995). Un-
der controlled conditions, the formation of the droplet
follows a 3–4h cycle (Morre et al.,1967). However,
in these laboratory experiments, the saturated mois-
ture and the periodic complete removal of the mucilage
might probably have increased the droplet formation.
In soil, it can be assumed that conditions might not be
as favourable to the production of such an important
amount of polysaccharide.
The properties of the mucilage secreted by the root
cap have been extensively studied. The COOgroups
of the mucilage can bind to cations and in particu-
lar, those fixed to clay (Guckert et al.,1975;Jenny
and Grossenbacher,1963). Consequently, soil struc-
ture is affected and stability of aggregate is generally
increased (Czarnes et al.,2000;Traore et al.,2000).
Heavy metals also bind to root cap slime (Moreletal.,
1986) and this may play a significant role in the pro-
tection of the root tip against their toxicity (Miyasaka
and Hawes,2001).
The root cap mucilage is able to hydrate exten-
sively. Fully hydrated mucilage has a water content of
100,000% of dry weight but such a hydration is only
obtained in the presence of free water (McCully and
Boyer,1997). Indeed, in mucilage collected on nodal
roots of maize, the water content (% of dry weight) in-
creases only up to 450% when the water potential of
the mucilage increases from 11 MPa to 0:01 MPa
(McCully and Boyer,1997). Thus, unless the soil is
saturated with water, the root cap mucilage appears as
a dry coating over the apex and does not form a droplet
as it is often observed in vitro (McCully and Sealey,
1996;Sealey et al.,1995). Furthermore, the surfactant
and viscoelastic properties of the mucilage (Read and
Gregory,1997) might favour the adhesion of root cap
cells to the soil particles and hence, their separation
from the cap as the root tip moves trough the soil. This
process is consistent with the rhizosheath observed on
rootsofgrasses(Vermeer and McCully,1982;Watt
et al.,1994). The sheath consists in soil Cmucilage
and living border cells tightly adhering to the root. The
mucilage originates both in the root cap and in micro-
bial syntheses (Watt et al.,1993). The sheath is not
observed just behind the root tip because the epider-
mis of this area has a thick complex surface on which
mucilage does not adhere (Abeysekera and MacCully,
1993;McCully,1999). The rhizosheath may function
like a biofilm involved in plant nutrition and may have
an important role in resistance to drought (Watt et al.,
1993).
The formation of the rhizosheath from root cap mu-
cilage suggests that its mineralization by microorgan-
isms is reduced or very slow. In vitro, root mucilage
can readily be utilised by rhizosphere bacteria as a sole
102 C. Nguyen
source of carbon (Knee et al.,2001). Furthermore, in a
laboratory experiment, Mary et al. (Mary et al.,1993)
demonstrated that maize mucilage incubated in soil
was mineralised at 45% of the added C within 2 weeks.
However, in the rhizosphere, mucilage mineralization
may be delayed by the preferential use by microorgan-
isms of root exudates, which are more readily available
and by the protection of mucilage due to its adsorption
to the soil matrix (Sollins et al.,1996).
The amounts of mucilage synthesized in vitro
ranges from 11 to 47 gMS=mg MS root growth
(Table 1). However, these quantities were determined
from roots grown in water or in nutrient solution,
which increases the outward diffusion of the mucilage
from the periplasmic region and probably stimulates
the biosynthesis of the slime (Sealey et al.,1995). Con-
sistently with this, the estimation of the quantity of
mucilage produced in soil based on the size of the
droplet surrounding the root cap in vitro might be over-
estimated: 700 and 1;250 m3=ha for wheat and maize,
respectively (Samsevitch,1965) (Table 1). At present
time, the amount of mucilage produced in soil remains
unknown.
2.3 Root Exudation
Excretion of organic compounds from roots was first
reported as early as the end of the nineteenth century.
In 1894, Dyer demonstrated the release of acidic
substances from roots of barley, wheat and others
(Krasil’nikov,1961). The biochemical nature of com-
pounds excreted by roots demonstrates a wide variety:
simple and complex sugars, amino acids, organic acids,
phenolics, alcohols, polypeptides and proteins, hor-
mones, enzymes (Curl and Truelove,1986;Grayston
et al.,1996;Neumann and Römheld,2000). In the
literature, the meaning of the term “exudation” may
differ significantly. Sensu stricto, exudates were first
defined as low molecular weight compounds diffusing
passively from intact cells to the soil solution (Rovira
et al.,1979). However, “root exudates” is often used to
describe more generally the low molecular compounds
released from roots regardless of the process by which
they are deposited into the rhizosphere. The main low
molecular weight compounds released passively from
roots are sugars, amino acids and organic acids. They
diffuse passively from the cytoplasm that is commonly
three orders of magnitude more concentrated than the
soil solution (mM vs M, respectively) (Neumann
and Römheld,2000). For example, in maize roots,
average concentrations are 86 mM for sugars (Jones
and Darrah,1996), 9.5 mM for amino acids (Jones
and Darrah,1994) and 10–20 mM for organic acids
(Jones,1998). The lipid bilayer of the plasmalemma
is a barrier to free diffusion of solutes because its
permeability is reduced, especially for charged com-
pounds compared to neutral molecules. However, the
protons excreted by the HC-ATPase provide an elec-
trochemical gradient for the diffusion of anions (Jones,
1998). Transient defects in the plasmalemma can also
significantly increase its permeability as suggested for
amino acids (Chakrabarti and Deamer,1992).
Membranes of plant cells bear sugar and amino
acids proton-coupled ATPase transporters that mediate
assimilate imports into cells (Bush et al.,1996).
Hence, it is not surprising that in vitro, plant roots are
able to actively take up sugars and amino acids from
asolution(Jones and Darrah,1994,1996;Schobert
and Komor,1987;Soldal and Nissen,1978;Xia and
Saglio,1988). The consequence of this influx on net
exudation may be important in axenic nutrient solution
but in soil, the evidences are less obvious. Indeed,
microorganisms are also very efficient competitors for
the uptake of sugars and amino acids (Coody et al.,
1986;Jones,1999;Nguyen and Guckert,2001;Vino-
las et al.,2001). The injection of labelled compounds
into the rhizosphere indicated that plant capture was
of minor importance compared to microbial uptake
of glucose and of charged or uncharged amino acids
(Nguyen et al.,2002;Owen and Jones,2001;Schobert
et al.,1988). Therefore, it is not known if the plant can
tune the net exudation in non-sterile soil by altering
the influx of sugars and amino acids.
In maize, the spatial examination of exudation indi-
cates a greater efflux of solutes close to the root apex
(McCully and Canny,1985;McDougall and Rovira,
1970). This does not seem to relate to variability in
the plasmalemma permeability nor to the spatial repar-
tition of transporters, which is uniform along maize
roots (Jones and Darrah,1994,1996). The greater exu-
dation behind the root apices is consistent with the con-
centration gradients of sugars and amino acids inside
the root (Jones,1998) and with the diffusion through
the apoplast of sugars from the phloem to the apical
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 103
meristems (Jones,1999), diffusion that is supported
by experimental and theoretical evidences (Bret-Harte
and Silk,1994).
The amount of C exuded has been expressed in a
wide range of units. Table 2gives some estimates re-
viewed from the literature for plants cultivated in nu-
trient solution. Due to the re-sorption of exudates by
plant roots, these values have to be considered with
caution if extrapolations to soil conditions are aimed.
It can also be seen that the proportion between sugar,
amino acids and organic acids vary greatly, especially
between sugars and organic acids. The relative propor-
tions of sugars and amino acids exuded reflect quite
correctly the relative concentrations of root tissues for
these solutes.
Besides the passive diffusion of solutes, plants are
able to response to environmental conditions by al-
tering their excretion of organic compounds. For ex-
ample, in response to environmental nutrient stress
such as P or Fe deficiencies, anion channel proteins,
embedded in the plasmalemma, increase significantly
the passive efflux of carboxylates (malate, citrate, ox-
alate) whose complexing properties facilitate nutri-
ent mobilization by the plant (Jones,1998;Neumann
and Martinoia,2002;Neumann and Römheld,2000).
The chelating properties of organic acids are also a
central mechanism involved in rhizosphere detoxifi-
cation as demonstrated in aluminium tolerant plants
(Barcelo and Poschenrieder,2002;Gaume et al.,2001;
Ma et al.,2001). Apart from organic acids, many
other compounds are released by plant roots in re-
sponse to environment. The most studied are phos-
phatases excreted by roots in P-stressed plants (Gaume
et al.,2001;Miller et al.,2001), phytosiderophores re-
leased in Gramineous plants and which are involved in
micronutrient acquisition (Crowley,2000;VonWiren
et al.,1996) and some phenolics such as flavonoids,
which play an important role in symbiosis establish-
ment (Werner,2000). A comprehensive review cov-
ering these compounds is available in (Neumann and
Römheld,2000). There are numerous evidences that
both the amount and the nature of root exudates are
very variable according to the physiological status of
the plant and to the plant species (Fan et al.,2001;
Grayston et al.,1996;Neumann and Römheld,2000).
Therefore, it can be assumed that the controlled re-
lease of particular exudates in response to sensed en-
vironmental stimuli is probably a major mechanism
that allow the plant to face unfavourable rhizosphere
conditions such as nutrient deficiencies, toxicities or
proliferation of pathogenic microorganisms.
2.4 Senescence of Root Epidermis
Behind the root tip, epidermal cells differentiate either
into hair cells (trichoblast) or non-hair cells (atri-
choblat). Root hairs are involved in anchorage, in water
and nutrient uptake and in symbiosis (Hofer,1991;Pe-
terson and Farquhar,1996). In the past recent years,
extensive research detailed the genetic control of root
hair development, especially in Arabidopsis (reviewed
in (Gilroy and Jones,2000;Schiefelbein,2000)). From
a study carried out by Dittmer (Dittmer,1949)on37
species belonging to 20 angiosperm families, the size
of root hairs is quite constant within a given species
but is very variable between species. Root hairs are
typically 80–1;500 m long and have a diameter of
5–20 m. The root hair zone is in average 1–4cm
long (Hofer,1991). Literature gives evidences that
root hair density is also very variable between plants:
1–180 hairs mm1of root, 70–10,800 hairs cm2of
root (Table 3).
Furthermore, environment strongly influences root
hair development. For example, low levels of minerals,
especially P and nitrate (Jungk,2001;Ma et al.,2001),
mechanical constraint, low O2partial pressure or high
temperatures stimulate root hair formation. Similar ef-
fects can be observed when roots are exposed to ethy-
lene which suggests that ethylene could be involved
in the regulation of root hair development by environ-
mental factors (Michael,2001).
There is little information about the lifespan of root
hairs. Based on the loss of the nucleus, it was esti-
mated that the longevity of root hairs was 2–3 weeks
in wheat, barley and maize (Fusseder,1987;Holden,
1975). However, microscopical examinations indicate
some cytoplasm lyses in 4 days old hairs in maize
(Fusseder,1987). Thus despite the cell wall can per-
sist for several weeks or months (Hofer,1991), the life
span of root hairs is probably shorter i.e. 2–3 days. If
root hairs are considered as cylinders that have a dry
weight:fresh weight (DW:FW) ratio of 0.072, a den-
sity equal to 1gcm
3and a C content of 40% DW, the
calculation for a hair density of 50 hairs mm1root
indicates that small hairs (80 m of length, 5mofdi-
ameter) correspond 2:2 ngC mm1root whereas large
104 C. Nguyen
Table 2 Quantities of C in root exudates of different plant species
References Plant Amount Units Compounds Comments
Barber and Gunn
(1974)Hordeum vulgare 76–157 g/plant/day Exudates Depending on mechanical
constraint, 21 days of
growth
0.2–0.4 %root DM/day Exudates Calculated from original
data
5–9 %root DM
Haggquist et al.
(1984)Brassica napus 16–21 gC/plant/day Total C Sterile and non sterile
roots, calculated from
original data
In Hale and
Moore (1979)Acer saccharum 2.7–6.7 %root DM/day Exudates Defoliated-control,
calculated from Smith
(1971)
Agropyron smithii 0.01 %root DM/day Reducing
sugars Defoliated/control, effect
of temperature,
calculated from
Bokhari and Singh
(1974)
Kraffczyk et al.
(1984)Zea mays 0.03–0.06 %root DM/day Sugars Sterile and non sterile
roots, 23 days of
growth
0.03–0.04 %root DM/day Organic acids Idem
0.001 %root DM/day Amino acids Idem
0.02–0.03 %root DM/day Sugars Three levels of K tested,
nitrate Cammonium,
23 days of growth
0.01–0.07 %root DM/day Organic acids Idem
0.0005–0.0007 %root DM/day Amino acids Idem
0.001–0.002 %root DM/day Sugars Three levels of K tested,
nitrate, 25 days of
growth
0.016–0.019 %root DM/day Organic acids Idem
0.0004–0.001 %root DM/day Amino acids Idem
Prikryl and
Vancura
(1980)
Triticum aestivum 121–153 g C/cm root
growth Exudates Sterile, nutrient solution:
2 or 4 day replacement
196–226 gC/mgDMroot
growth Exudates Sterile non sterile nutrient
solution: 2 day
replacement
576–1,174 gC/mgCroot
growth Exudates Nutrient solution: 2 day
replacement,
sterile-inoculated with
Pseudomonas putida
Exudates
Jones and Darrah
(1993)Zea mays 0.1–1.2 % root DM/day Exudates Calculated from original
data, Sterile, no or
daily changes of
nutrient solution,
10 day culture
1.22 gC/root tip/h Exudates Standard values for model
simulation
0.24 gC=cm of root/h Exudates Idem
hairs (1;500 m of length, 20 m of diameter) are
equivalent to 680 ng C mm1root. Medium size hairs
(500 m of length, 10 m of diameter) correspond to
56 ngC mm1root. Theoretically, these amounts of C
should be deposited into the soil after the hair death.
However, to our knowledge, it is unknown if the cy-
toplasm material is released into the soil or recycled
within the root tissue.
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 105
Table 3 Root hair density in different plant species
Root hairs Root hairs Root hair
References Plant Root radius (cm) .mm1root) .cm2root) length (mm)
Gahoonia et al.
(1997)Triticum aestivum 8:50 10338 7,115 1.27
Triticum aestivum 8:00 10325 4,974 0.74
Triticum aestivum 7:50 10324 5,093 0.49
Hordeum vulgare 8:50 10330 5,617 1.1
Hordeum vulgare 7:50 10327 5,730 0.52
Hordeum vulgare 7:50 10331 6,578 1
Hordeum vulgare 8:00 10330 5,968 0.64
Föse (1971)in
Jungk (2001)Spinacia oleracea 1:07 10271 10,561 0.62
Brassica napus 7:30 10344 9,593 0.31
Lycopersicon esculentum 1:00 10258 9,231 0.17
Triticum aestivum 7:70 10346 9,508 0.33
Allium cepa 2:29 1021690.05
Lolium perenne 6:60 10345 10,851 0.34
Phaseolus vulgaris 1:45 10249 5,378 0.2
Masucci et al.
(1996)
Arabidospis thaliana 53–63
Bouma et al.
(2000)Elymus pycnathus (L.)
Main root 44 0.37
First-order branching 7 0.32
Second-order branching 3.5 0.32
Puccinellia maritima (L.)
Main root 20 0.51
First-order branching 11 0.47
Second-order branching 5 0.5
Spartina anglica (L.)
Main root 21 0.17
First-order branching 10 0.24
Second-order branching 5 0.25
Wulfsohn et al.
(1999)Agropyron cristanum
Main root 71 0.19
Branchings 181.6 0.153
Despite it is not a general rule, there are numerous
reports that cells from the root epidermis senesce (Curl
and Truelove,1986). For instance, in maize, the senes-
cence is extensive proximal to the region where the late
metaxylem matures (Wenzel and McCully,1991). The
senescence can even concern cortical cells. The nuclear
staining with acridine orange pointed out that senes-
cence of cortical cells concerns the old parts of the
roots but some works also indicated the absence of nu-
cleus in the cortex of young roots in cereals (Fusseder,
1987;Henry and Deacon,1981;Holden,1975). How-
ever, the impermeability of the cell wall to the stain
may cause an artefact that biases the evaluation of
the cell vitality (Wenzel and McCully,1991). Thus,
it would be necessary to gain more information about
(1) the life span of the root epidermis (including root
hairs) and of the root cortex in soil conditions, (2)
the fate of the intracellular content of the senescing
root cells.
2.5 Relative Proportion of Rhizodeposits
Due to the very different units used to express the
quantities of C from rhizodeposits, comparisons are
difficult. However, from Table 1, it is reasonable to es-
timate that border cells represents 1–3gCmg
1DM
of root growth, or 1:5–2:5g C day1root1.Inav-
erage, mucilage ranges between 2 and 20 gCmg
1
dry matter (DM) of root growth, assuming a C con-
tent of 39% DM. In comparison, Table 2indicates
mean exudation values of 150 gCmg
1DM of root
growth, 0.2–7% root DM/day. These calculations sug-
gest that exudation releases almost 10–100 times more
carbon than border cells and mucilage. As calculated
previously, the death of root hairs with a medium size
and density would deposit 56 ng C mm1root. If root
hair decay concerns 1 cm of root per day, which is
reasonable, the amounts deposited are 3 orders of mag-
nitude less than exudation.
106 C. Nguyen
3 Factors Affecting C Fluxes
to the Rhizosphere
Factors affecting the release of C from roots into
the soil are numerous and have been extensively re-
viewed e.g. (Curl and Truelove,1986;Grayston et al.,
1996;Hale and Moore,1979;Rovira,1969;Whipps,
1990). The literature points out that the total amounts
of organic C deposited in the rhizosphere can vary
greatly according to the plant ecophysiology. This
can be explained as follows. Both the environment
and the plant genetics and physiology can influence
(1) the flux of C from each root to the rhizosphere,
which is related to the root functioning and (2) the
size and the morphology of the overall root. There-
fore, any attempt to model rhizodeposition will have
to consider the plant ecophysiology. The aim of that
part of the paper is to examine tracer studies to anal-
yse the main factors that affect rhizodeposition. To
reach that goal, we have analysed the partitioning of
net fixed C between the plant-soil pools from 14C
tracer experiments. The main factors examined are
plant age, microorganisms, soil texture, soil nitro-
gen and atmospheric CO2concentration. There are
of course numerous other factors that alter rhizode-
position. They were not detailed in the present study
because no sufficient data from 14C-labelling experi-
ments were available. Among them are light intensity
(Hodge et al.,1997), photoperiod (Todorovic et al.,
1999), temperature (Martin and Kemp,1980), soil pH
(Meharg and Killham,1990a), anoxia (Meharg and
Killham,1990c), defoliation (Holland et al.,1996;
Paterson and Sim,2000).
Tracer experiments have been chosen because stud-
ies are numerous and because the expression of results
in terms of partitioning coefficients of net fixed C is a
common basis for the majority of articles. Indeed, in
non-tracer experiments, the comparison between stud-
ies is difficult or impossible because the classification
of rhizodeposits is not uniform among articles (soluble,
insoluble, sugars, total C, etc.) and because results are
expressed in a wide range of units (Toal et al.,2000).
Among tracer experiments, labelling of photoassimi-
lates with 14C is the most commonly used technique to
study C flow to the rhizosphere. Others isotopes have
also been used for labelling experiments such as 11C
and 13C and C flows to the soil can also be studied
using natural abundance of 13C. All these techniques
have been reviewed elsewhere e.g. (Kuzyakov and
Domanski,2000;Meharg,1994;Morgan and Whipps,
2000) and will not be detailed here.
The present study examines experiments in which
plants shoots were exposed to 14C except (Palta and
Gregory,1997) in which 13C was used as tracer. The
exposition of shoots to the tracer was either as a pulse
(few minutes to several hours) or as a permanent ex-
position from germination until sampling. These two
procedures refer to as pulse or continuous labellings.
Briefly, pulse labelling experiments are very useful to
obtain information on C fluxes in relation to the plant
ecophysiology but due to the brief exposition of the
plant to the tracer, this technique fails to provide reli-
able C budgets, which can be assessed by continuous
labelling. Moreover, on a technical point of view, con-
tinuous labellings differ from pulse-chase experiments
in that they are cumbersome, expensive and hardly ap-
plicable in field situations (Meharg,1994). However,
Warembourg and Estelrich (Warembourg and Estel-
rich,2000) compared 298 h and 78 day long labellings
in Bromus erectus. They concluded that reliable esti-
mations of C fluxes to the rhizosphere can be obtained
from an intermediate strategies consisting in repeated
short-term labellings of a few days each.
The tracer experiments reviewed for the present
study expressed results as 14C-partitioning coefficients,
i.e. percentages of the net fixed C allocated to C com-
partments. The compartments are shoot and root C,
CO2from rhizosphere respiration (root respiration C
the rhizomicrobial respiration i.e. microbial respiration
derived from rhizodeposition) and C in soil residues.
The respective partitioning coefficient are SHOOT,
ROOT, RR, RES. We have also investigated parti-
tioning of 14C between belowground compartments as
percentages of labelled C exported by shoots. These
partitioning coefficients are ROOTBG, RRBG and
RESBG. Articles for which 14C partitioning was not
complete were discarded. Data were analysed with
SAS V 8.02 for Windows (Microsoft), The SAS
Institute Inc., Cary NC, USA.
3.1 Data Overlook
There were 43 articles examined. A given article
presents as many sets of 14C-partitioning coefficients
as experiments/treatments. For example, an article that
examines the effect of nitrogen fertilization .CN;N/
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 107
and of elevated CO2(elevated, ambient) will provide
four sets of coefficients. Hence, the total number of
coefficient sets analysed was 237 (Table 4). There are
more data for pulse labellings than for continuous la-
bellings (137 sets of coefficients vs 100, respectively)
despite the number of articles analysed are compara-
ble, around 20. Whatever the labelling procedure, the
maximum contribution of an article to the total number
of coefficient sets is 12–26% (data not shown).
The data point out that tracer experiments focus on
a restricted number of plants (Table 4). There are 1.5
times more data for annual plants than for perennials.
Among annuals, Triticum aestivum and Hordeum vul-
gare and Zea mays represent 88% of the coefficient sets
and among perennials Lolium perenne represents more
than 35% of the coefficient sets. More than 65% of the
coefficient sets concern Lolium perenne and Bromus
erectus.DataonBromus erectus were drawn from a
single article. Examination of plant age indicates that
the great majority of the data concerns juvenile stage
of development (Table 5). In continuous labelling ex-
periments, the mean is 37 days and the median 28 days.
In pulse labelling studies, the mean is 146 days and the
median is 87 days but the coefficient of variation for
the mean is two times greater that that in continuous la-
bellings. The difficulty to maintain an atmosphere with
a constant 14CO2activity and constant CO2concentra-
tion (Warembourg and Kummerow,1991) can greatly
contribute to explain the fact that continuous labelling
focussed on younger plants compared to pulse-chase
studies. Indeed, in pulse labelling experiments, late de-
velopment stages such as flowering and grain filling
have been investigated (Keith et al.,1986;Meharg and
Killham,1989;Swinnen and Van Veen,1994).
Table 6outlines that the partitioning coefficients
from continuous labelling studies are normally dis-
tributed except the RES and RESBG coefficients (14C
in soil residues), the distributions of which are skewed
to low values (Data not shown). This means that the
majority of the data are low and few coefficients extend
to greater values. The root sampling procedure may
provide a possible explanation for the non-normality
of these coefficients. Indeed, it is very difficult to
separate, by hand picking/sieving, the fine roots from
the soil and a variable proportion of them may be left
in the soil, increasing artificially the values of RES and
RESB coefficients. Moreover, the washing of roots ex-
tracts some soluble 14C, which can also overestimate
labelled soil residues (Swinnen et al.,1994). In pulse
labelling experiments, none of the partitioning coef-
ficients are normally distributed (Table 6). The dis-
tribution of PA coefficients has a low Kurtosis (data
not shown). For the other coefficients, once again,
data are skewed to the low values. The non-normality
of the distribution of the partitioning coefficients in
pulse-chase experiments can be explained by the non-
standardization of the labelling procedures. Among the
studies, both the exposition of shoots to 14CO2and
the chase period are very variable in length (Table 5).
The length of the labelling ranges from 20 min to
720 h (data not shown) with a median of 6h and a
mean of 108 h. The length of the chase period, that is
the time elapsed between the labelling and the sam-
pling, is probably the key point that affects assimi-
lates partitioning. In the articles reviewed, the chase
period ranges from 30 min to 504 h (data not shown)
with a mean and a median equal to 145 h and 48 h,
respectively (Table 6). This indicates that in general,
the chase period is short, which could lead to an in-
complete partitioning of 14C and to an overestimation
of the 14C recovered in shoots and an underestima-
tion of C flows to belowground. This is supported by
the greater mean SHOOT coefficient in pulse-labelling
studies compared to continuous labelling studies (64 vs
57, Table 6). Conversely, a long chase period may in-
crease the labelled carbon retrieved in the rhizosphere
respiration and decrease the 14C in the soil residues.
The mean partitioning coefficients for SHOOT,
ROOT, RR and RES determined from pulse and con-
tinuous labelling experiments are 64%, 20%, 12% and
5% and 57%, 22%, 14%, 7%, respectively. This indi-
cates that shoots export almost half of the net fixed C to
belowground (Lambers,1987). In average, among the
net C allocated belowground, half is retained in root
tissues, a third is lost as root Crhizomicrobial respira-
tion and more than 15% is retrieved as soil residues. It
is interesting to note that in pulse-chase experiments,
SHOOT coefficients for annual plants are greater than
that of perennials, the contrary being observed for
ROOT and RR coefficients. There are also differences
between perennials and annuals for the belowground
budget. Further investigations are needed to explore
if these results are representative of a different strat-
egy of assimilates partitioning to the soil between an-
nual and perennial plants (Warembourg and Estelrich,
2001).
108 C. Nguyen
Table 4 Number of partitioning coefficient sets reviewed for different plant species. A partitioning coefficient set consists in the
percentages of the tracer allocated to shoots, roots, rhizosphere respiration and soil residues
Annual (A)/
perennial (P) Plante Labelling References
Partitioning coefficient sets
Number Total/species
% of total
relative to
labelling % of total
ATriticum aestivum CBarber and Martin (1976);
Billes et al. (1993);
Liljeroth et al. (1994);
Martin (1977); Martin
and Kemp (1980);
Merckx et al. (1985,
1986); Swinnen and Van
Veen (1994)
45 56:3
APGregory and Atwell (1991);
Keith et al. (1986); Palta
and Gregory (1997);
Paterson et al. (1996);
Swinnen et al. (1994);
Swinnen et al. (1995);
Whipps (1984); Whipps
and Lynch (1983)
30 75 49:2 53:2
AHordeum vulgare CBarber and Martin (1976);
Johansson (1992);
Whipps (1984); Whipps
and Lynch (1983); Zagal
(1994)
13 16:3
APGregory and Atwell (1991);
Jensen (1993); Swinnen
et al. (1995)
13 26 21:3 18:4
A Zea mays C Helal and Sauerbeck (1984,
1986); Liljeroth et al.
(1994); Martens (1990);
Merckx et al. (1987);
Whipps (1985)
15 18:8
APHolland et al. (1996);
Kisselle et al. (2001);
Todorovic et al. (2001)
8 23 13:1 16:3
ABromus madritensis PWarembourg and Estelrich
(2001)
8 8 13:1 5:7
ABrassica napus CZagal (1994)22:5
APShepherd and Davies (1993)13 1:6 2:1
ALycopersicon esculente CWhipps (1987)22 2:5 1:4
APisum sativum CWhipps (1987)22 2:5 1:4
AMedicago truncatula CCrawford et al. (2000)11:3
APCrawford et al. (2000)12 1:6 1:4
Total continuous labelling
experiments 80
Total pulse labelling
experiments 61
Total 141
PLolium perenne PDomanski et al. (2001);
Meharg and Killham
(1989,1990a–c); Paterson
et al. (1996,1999);
Rattray et al. (1995)
23 30:3
(continued)
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 109
Table 4 (continued)
Annual (A)/
perennial (P) Plante Labelling References
Partitioning coefficient sets
Number Total/species
% of total
relative to
labelling % of total
PCGorissen et al. (1996); Hodge
et al. (1997); Van Gink e l
et al. (1997); Zagal (1994)
12 35 60:0 36:5
PBromus erectus PWarembourg and Estelrich
(2001)
28 28 36:8 29:2
PCastanea sativa PRouhier et al. (1996)8 8 10:5 8:3
PTrifolium repens PTodorovic et al. (1999)55 6:6 5:2
PFestuca arundinacea CGorissen et al. (1996)4 4 20:0 4:2
PPinus taeda PReid et al. (1983)44 5:3 4:2
PPopulus tremuloides PMikan et al. (2000)44 5:3 4:2
PFestuca pratensis CJohansson (1992a,1993)3 3 15:0 3:1
PCynodon dactylon PPaterson et al. (1996)22 2:6 2:1
PLolium multiflorum P Henry et al. (in press) 22 2:6 2:1
PBouteloua gracilis CDormaar and Sauerbeck
(1983)
11 5:0 1:0
Total pulse labelling
experiments 76
Total continuous
experiments 20
Tot a l 9 6
CDcontinuous labelling, P Dpulse labelling
Table 5 Age of plants and labelling characteristics in the tracer experiments reviewed
Continuous labelling experiments Pulse labelling experiments
Annual (A)/
perennial (P) NaMean Median CV of
meanbNaMean Median CV of
meanb
Age (days) A 80 31:7 24:0 51:2 60 97:5 82:5 68:2
P20 56:0 59:0 31:1 66 190:5 93:0 97:5
Tot al 100 36:6 28:0 52:1 126 146:2 86:5 101:9
Chase (h) A 60 230:6 92:0 96:5
P72 73:6 48:0 78:1
Tot al 132 145:0 48:0 120:0
Length of labelling (h) A 60 60:1 1:8 237:8
P72 148:2 76:0 105:8
Tot al 132 108:2 6:0 144:6
aNumber of partitioning coefficient sets
bCoefficient of variation of the mean (%)
The SHOOT coefficient is significantly and nega-
tively correlated to all of the belowground coefficients
(ROOT, RR and RES) (Table 7). The correlations are
stronger in pulse-chase experiments, which can be re-
lated to the greater temporal resolution of pulse la-
belling compared to continuous labelling. Whatever
the kind of labelling, the ROOTBG, coefficient is
significantly correlated positively to RRBG and to
RESBG. This suggests a strong link between rhizode-
position and the metabolic activity of roots. This is
consistent with the mechanisms involved in the re-
lease of C from roots. Indeed, an important exporta-
tion of photoassimilates from the shoots to the roots
is expected to maintain the solute gradient between
the root tissue and the soil solution and so, to favour
the passive diffusion of root exudates into the soil.
Moreover, a rapid root growth should increase the
number of border cells and mucilage deposited into
the soil as the result of frictional forces experienced
by the foraging root apices.
110 C. Nguyen
Table 6 14C budget for tracer studies of C translocation into the soil. Results are expressed as percentages of the net 14Cfixed
Annual (A)/
Labellingaperennial (P) NbMean Maximum Minimum Median CV of meancPnd
C A 80 56.6 78.9 34.8 55.2 16.8
C P 20 58.2 86.4 22.0 60.1 26.9
SHOOT C All 100 57.0 86.4 22.0 55.6 19.2 0.302
P P 76 56.5 99.1 18.8 56.8 33.9
P A 61 72.8 97.4 25.6 73.9 25.8
PAll 137 63.7 99.1 18.8 65.8 32.3 0.003
C A 80 21.2 37.5 3.9 21.8 37.1
C P 20 27.1 40.4 9.1 28.9 33.8
ROOT C All 100 22.4 40.4 3.9 23.4 37.7 0.336
P P 76 24.0 55.0 0.1 21.8 60.5
P A 61 13.8 55.9 0.4 10.4 95.3
PAll 137 19.5 55.9 0.1 16.1 75.9 <0.0001
C A 80 14.5 26.1 0.1 15.3 44.5
C P 20 10.1 22.0 0.5 9.3 60.7
RR C All 100 13.6 26.1 0.1 13.9 48.5 0.050
P P 76 14.3 57.0 0.5 14.2 66.6
P A 61 8.9 35.7 0.9 6.2 81.2
PAll 137 11.9 57.0 0.5 10.9 75.3 <0.0001
RES C A 80 7.7 30.4 1.2 6.4 78.2
C P 20 4.5 23.0 1.9 3.6 100.1
CAll 100 7.1 30.4 1.2 5.3 83.1 <0.0001
P P 76 5.1 16.0 0.0 4.6 72.6
P A 61 4.5 20.7 0.1 3.5 95.2
PAll 137 4.9 20.7 0.0 4.2 82.2 <0.0001
C A 80 48.4 76.8 10.0 51.2 28.2
C P 20 66.0 84.8 40.3 67.3 16.9
ROOTBG C All 100 52.0 84.8 10.0 55.2 28.7 0.181
P P 76 53.9 90.8 1.6 57.9 40.6
P A 61 44.1 86.2 11.2 42.3 48.0
PAll 137 49.6 90.8 1.6 52.1 44.5 0.006
C A 80 34.0 79.3 0.2 36.2 41.9
C P 20 22.6 45.1 3.6 24.8 46.4
RRBG C All 100 31.7 79.3 0.2 33.4 45.0 0.024
P P 76 34.2 86.4 1.2 29.2 64.4
P A 61 35.6 63.8 7.0 37.6 41.0
PAll 137 34.9 86.4 1.2 33.1 54.7 0.015
C A 80 17.6 69.4 2.4 15.3 68.0
C P 20 10.9 29.5 4.3 8.8 57.6
RESBG C All 100 16.3 69.4 2.4 14.4 69.9 <0.0001
P P 76 11.8 35.6 0.1 11.9 61.1
P A 61 20.2 69.9 1.1 19.6 77.1
PAll 137 15.6 69.9 0.1 13.9 79.5 <0.0001
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated to belowground
aCDcontinuous labelling, P Dpulse labelling
bNumber of partitioning coefficient sets
cCoefficient of variation of the mean (%)
dTest for the normality of the distribution (Shapiro-Wilk test). Probability associated to the null hypothesis of normality of the
distribution
Data from continuous labelling experiments indi-
cate that the RRBG and RESBG coefficients are sig-
nificantly negatively correlated .rD0:32/. Hence,
this might reflect the fact that according to the studies,
a variable fraction of the rhizodeposits are mineralised
by the microorganisms, which consequently alters sym-
metrically the RESBG and RRBG coefficient. If this
hypothesis is valid, this means that the rhizomicrobial
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 111
Table 7 Pearson correlation coefficients between partitioning coefficients in tracer studies. The value in italic is the probability
associated with the null hypothesis Rho D0
LabellingaSHOOT ROOT RR RES
C SHOOT 1
CROOT0:675 1
<0.0001
CRR0:433 0:117 1
<0.0001 0.2459
CRES0:417 0:043 0:141 1
<0.0001 0.6731 0.1614
PPA1
PRAC0:857 1
<0.0001
PRR0:648 0.217 1
<0.0001 0.011
PRES0:515 0.257 0.341 1
<0.0001 0.0024 <0.0001
Belowground budget ROOTBG RRBG RESBG
CROOTBG1
C RRBG 0:696 1
<0:0001
CRESBG0:452 0:324 1
<0:0001 0.001
PROOTBG1
P RRBG 0:782 1
<0:0001
PRESBG0:535 0:008 1
<0:0001 0.9239
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated to belowground
aCDContinuous labelling, PDPulse labelling
respiration contributes significantly to the rhizosphere
respiration. In pulse labelling experiments, no such cor-
relation is observed probably because of the variability
of the length of the chase period.
3.2 Factors that Affect the Partitioning
of 14C-Assimilates to the Soil:
A Quantitative Approach
3.2.1 Methods for Calculations
The following methods were applied to appreciate the
effect of a factor on the partitioning of 14C between
plant-soil C pools. Be a given factor F tested at n levels,
the relative variation (RV) in a partitioning coefficient
‘PC’ was calculated as RV D.PCnPCn1/=PCn1.
The level ‘n’ of the factor F was always high relative
to the level n 1. For example, if an article reports
on the effect of N fertilization tested at N1 <N2
levels, the relative variation in partitioning of 14C
to shoots was calculated as follows: RVSHOOT D
.SHOOTN2 SHOOTN1/=SHOOTN1.Thesame
calculations were performed for the other partitioning
coefficients. Hence, a positive RV indicates that the
factor increases the partitioning of assimilates to the
compartment considered. If a second factor was stud-
ied such as the concentration of atmospheric CO2, ap-
plied at two levels L1 <L2, the effect of nitrogen was
calculated at the two levels of CO2W.SHOOTN2L1
SHOOTN1L1/=SHOOTN1L1 and .SHOOTN2L2
SHOOTN1L2/=SHOOTN1L2. If a factor was tested at
more than two levels L1 <L2 <L3 ::: < Ln, the
effects were calculated relative to two subsequent
levels: Ln vs Ln 1; Ln 1vs Ln 2:::L2 vs L1.
For each factor investigated, the relative variation
coefficients RVs were classified according to the la-
belling procedure (continuous or pulse). Then, the
maximum, the minimum, the median, the mean and its
112 C. Nguyen
coefficient of variation were calculated. The normality
of the RVs distributions were tested by the Shapiro-
wilk test. If normalilty was accepted (at alpha D5%),
the Student t test was used to test the null hypothesis:
mean D0, otherwise the non-parametric sign test was
applied to test the null hypothesis: median D0.
3.2.2 Plant Age
The data from pulse-chase experiments summarized in
Table 8clearly demonstrate that plant age significantly
influences C partitioning of photoassimilates between
plant-soil compartments. As the plant gets older, less
carbon is partitioned to belowground. Data being non
normally distributed, medians are examined. They are
43% to roots, 28% to rhizosphere respiration and
20% to soil residues (Table 8). The variability is very
important as illustrated by the coefficients of variation.
This is not surprising because the effect of age are more
marked for young plant than for older ones, which is
not taken into account in the calculations. No clear sig-
nificant effect of age on C partitioning between below-
ground compartments can be observed. The medians
of ROOTBG, RRBG and RESBG suggest that C al-
located to belowground is less retained in roots when
plant age increases. The partitioning coefficients from
continuous labelling experiments do not evidence this
pattern due to their low temporal resolution and due to
the fact that very young plants were considered in these
studies (37 day old in average, Table 5). Thus, the de-
cline in C inputs into the soil with plant age is related to
the decrease of assimilates partitioning to roots, which
is particularly marked for annual plant (Keith et al.,
1986;Swinnen and Van Veen,1994;Swinnen et al.,
1995).
3.2.3 Microorganisms
Only eight experiments are reported here for contin-
uous labelling experiments and one in case of pulse
labelling study (Table 9). This does not mean that the
effects of microorganisms have not been investigated,
but here, we only consider soil or sand culture exper-
iments. In the literature, due to the difficulty to ster-
ilized soil microcosms efficiently, the great majority
of works investigating the influence of microorgan-
isms on rhizodeposition have been performed in nutri-
ent solution e.g. (Lee and Gaskins,1982;Meharg and
Killham,1991).
The results indicate that microorganisms strongly
increased the 14C partitioned to the rhizosphere. In av-
erage, in non-sterile cultures, the RR and SOIL co-
efficients are significantly increased of C249% and
C37%, respectively (Table 9). The same effects are ob-
served for belowground budget. Belowground, less la-
belled C is partitioned to roots .10%/whereas 14Cin
rhizosphere respiration and in soil residues increased
of C199% and C24%, respectively, but the variation of
RESBG is not significant. Despite the small number of
articles considered here, there are strong evidences that
microorganisms increase greatly the partitioning of as-
similates to the rhizosphere. There are several possible
explanations. First, in non-sterile conditions, roots can
establish symbiosis with mycorrhizal fungi. Mycor-
rhizae represent a significant sink for plant assimilates
(Leake et al.,2001;Wu et al.,2002) since up to 30% of
the photoassimilates can be allocated to the symbiotic
fungus (Nehls and Hampp,2000). Consequently, fun-
gal respiration could explain the greater allocation of
labelled C to rhizosphere respiration whereas growth
of extraradicular hyphae and the hyphal C exudation
contribute to a large extend to the plant-derived carbon
retrieved as soil residues (Högberg and Högberg,2002;
Johnson et al.,2002;Sun et al.,1999). On the other
hand, non-symbiotic rhizosphere microorganisms take
up and assimilate soluble low molecular weight com-
pounds released passively from root and hence, they
maintain the C gradient between the internal root tis-
sues and the soil solution. Furthermore, rhizosphere
microflora can synthesize enzymes or metabolites that
can alter the integrity of root cells or the permeability
of their membrane. Finally, root morphology can be
modified directly by phytohormones produced by rhi-
zosphere microorganisms or indirectly by the changes
in nutrient availability resulting from microbial pro-
cesses. Consequently, any changes in root branching
pattern would be expected to have significant conse-
quences on root exudation, which can be more impor-
tant at the root apices, such as in maize for instance
(Jones and Darrah,1996). Besides quantitative aspects
of root exudation, both free and symbiotic microorgan-
isms change the quality of root exudates. For example,
Pinior et al. (Pinior et al.,1999) demonstrated that ex-
udates from non mycorrhizal roots of cucumber stimu-
late hyphal growth of the mycorrhizal fungi Gigaspora
rosea and Gigaspora intraradices whereas exudates
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 113
Table 8 Effect of plant age on labelled C partitioning between plant and belowground compartments. Effects are expressed as
relative variations, see the text for explanations about the calculation of the effects
Relative Variation(%)
Factor LabellingaReferences NbMean
CV of
mean Max Med Min PncPtdPme
Age (days)
CLiljeroth et al.
(1994); Merckx
et al. (1985,1986,
1987); Martens
(1990); Whipps
(1985,1987);
Zagal (1994)
SHOOT 21 4 503 59 4 33 0:376 0:373
Min D14 CROOT21 4 1;165 157 851 0:001 0:664
Max D76 CRR21 15 266 111 5 35 0:043 0:664
Mean D39 CRES21 11 381 81 19 68 0:033 0:064
Median D41 CROOTBG21 5 937 182 263 <0:001 1:000
CV D39 C RRBG 21 16 211 132 2 20 <0:001 0:007
CRESBG21 9464 95 21 65 0:075 0:335
PGregory and
Atwell (1991);
Jensen (1993);
Keith et al.
(1986); Meharg
and Killham
(1990b); Palta and
Gregory (1997);
Reid et al. (1983);
Rouhier et al.
(1996); Swinnen
and Van Veen
(1994); Swinnen
et al. (1995);
Warembourg and
Estelrich (2001)
SHOOT 45 17 196 103 8 57 0:013 0:000
Min D28 PROOT45 26 224 250 43 76 <0:001 0:000
Max D600 PRR45 110;764 311 28 85 <0:001 0:008
Mean D151 PRES45 18 596 374 20 93 <0:001 0:542
Median D106 PROOTBG45 6527 143 14 55 <0:001 0:096
CV D93 P RRBG 45 26 288 293 10 65 <0:001 0:291
PRESBG45 39 203 312 24 77 <0:001 0:096
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated to belowground
aCDcontinuous labelling, P Dpulse labelling
bNumber of partitioning coefficient sets
cP associated to Shapiro-Wilk test for normality
dP associated to Student test for location Mu D0for data normally distributed
eP associated to the non parametric sign test for location Median D0 for data non normally distributed
from roots colonized by Gigaspora rosea inhibited fur-
ther root colonization by Glomus mossae. Therefore,
the soil microflora strongly modifies root exudation,
which in turn alters both the size and the structure of
microbial populations in the rhizosphere (Brimecombe
et al.,2000;Grayston,2000).
3.2.4 Soil Texture
Here, we report on experiments that compared plants
grown on soils differing in their clay and loam con-
tents. It is important to note that the range of the
clay content reported here is low, from 2% to 15%
(Table 10) due to the difficulty to sample the roots in
soils with high clay contents. An increase in clay and
loam content of soil alters greatly the partitioning of
14C. Significantly more labelled C is retained above-
ground and less is allocated to roots (mean variation D
C15% and 25%, respectively, Table 10). Both the
global and the belowground 14C budget indicate that
partitioning of 14C to rhizosphere respiration and to
soil residues are also increased but these effects are
not significant due to the work of Whipps and Lynch
(Whipps and Lynch,1983) that indicated surprisingly
low values for RR in the soil with a light texture. The
increase in C loss from root in soil with increasing
clay and loam contents is not surprising because nu-
merous soil properties, which favour microbial activ-
ity and nutrient cycling, are related to the clay content:
water retention, organic matter stabilization, high
cation exchange capacity, for example. Thus, the sug-
gested stimulation of rhizodeposition in relation to clay
and loam contents of soil could be explained by some
114 C. Nguyen
Table 9 Effect of soil microorganisms on labelled C partitioning between plant and belowground compartments. Effects are ex-
pressed as relative variations, see the text for explanations about the calculation of the effects.
Factor LabellingaReferences Nb
Relative variation (%)
Mean CV of
mean Max Med Min PncPtdPme
Microorganisms
Non sterile vs sterile
C SHOOT 8 4316 21 524 0.841 0.400
CBarber and
Martin
(1976);
Martin
(1977);
Todorovic
et al. (2001)
ROOT 8 2 918 31 7 22 0.524 0.767
CRR8249 112 658 157 15 0.086 0.040
C RES 8 37 175 181 24 16 0.017 0.727
CROOTBG810 110 7 14 21 0.246 0.037
C RRBG 8 199 119 598 114 18 0.097 0.049
C RESBG 8 24 271 169 724 0.005 0.727
PWhipps and
Lynch (1983)
SHOOT 1 8888
PROOT126 26 26 26
PRR137 37 37 37
P RES 1 886 886 886 886
PROOTBG138 38 38 38
P RRBG 1 14 14 14 14
P RESBG 1 723 723 723 723
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated to belowground
aCDcontinuous labelling, P Dpulse labelling
bNumber of partitioning coefficient sets
cP associated to Shapiro-Wilk test for normality
dP associated to Student test for location Mu D0 for data normally distributed
eP associated to the non parametric sign test for location Median D0 for data non normally distributed
Table 10 Effect of soil texture on labelled C partitioning between plant and belowground compartments. Effects are expressed as
relative variations, see the text for explanations about the calculation of the effects
Factor LabellingaReferences
NbRelative variation (%)
Mean
CV of
mean Max Med Min PncPtdPme
Soil texture (% of clay/loam)
C SHOOT 11 15 130 42 11 16 0:455 0.029
Min D2/10 C Gorissen et al.
(1996);
Merckx et al.
(1985,1986);
Whipps and
Lynch (1983)
ROOT 11 25 118 44 22 66 0:195 0.018
Max D15/71 C RR 11 3;479 181 19;500 24 44 <0:001 1.000
Mean D9/29 C RES 11 67 185 233 6 72 0:025 1.000
Median D13/12 C ROOTBG 11 19 98 6 12 51 0:340 0.007
CV D71/83 C RRBG 11 4;789 184 27;158 45 38 <0:001 0.549
CRESBG11 77 169 320 24 61 0:136 0.079
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated belowground
aCDcontinuous labelling, P Dpulse labelling
bNumber of partitioning coefficient sets
cP associated to Shapiro-Wilk test for normality
dP associated to Student test for location Mu D0 for data normally distributed
eP associated to the non parametric sign test for location Median D0 for data non normally distributed
differences in fertility and in microbial activity. Be-
sides, the effect of the soil texture on C fluxes to
the rhizosphere can also be explained by the physi-
cal properties of the soil. Indeed, soil texture is in-
terrelated with bulk density and porosity and the re-
sulting mechanical impedance has been reported to
increase rhizodeposition (Boeuf-Tremblay et al.,1995;
Groleau-Renaud et al.,1998). On a theoretical point
of view, the mechanical impedance in soils with a fine
texture should promote the sloughing-off of root cap
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 115
cells. Root exudation may also be favoured by the
small size of soil pores, which increases the surface
of the root that is covered by soil aggregates and which
consequently facilitate mass flow diffusion of solutes.
Hence, both experimental data and theoretical consid-
erations support the relevancy of considering soil tex-
ture when investigating of C fluxes to the rhizosphere.
3.2.5 Soil Nitrogen
Soil nitrogen is a major factor that can severely limit
plant growth and therefore, the effect of N fertilization
on C fluxes to the rhizosphere is a highly relevant
question. We summarized 9 data sets for continuous
labelling experiments and 19 for pulse-chase studies,
the latter being mainly related to an experiment on
Bromus erectus (Warembourg and Estelrich,2001)
(Table 11). All data sets indicate that when plant are
N fertilized, there is a highly significant decrease of
labelled C partitioning to roots (14% and 35%
for continuous and pulse labellings, respectively) and
conversely, an increase of 14 C retrieved in shoots
(C11% and 36% for continuous and pulse labellings,
respectively) (Table 11). The coefficients of variation
are not excessive. The effect is more marked in
pulse-chase experiments. This is consistent with the
low root:shoot ratio of N-fertilized plants, which is
commonly observed experimentally and which is well
described by the functional equilibrium theory (Farrar
and Jones,2000). The global 14C budget does not point
out a significant effect of nitrogen on C allocation to
rhizosphere respiration and to soil residues whatever
the labelling procedure. However, the belowground
budget is altered by N fertilization, in pulse-chase
experiments. Indeed, the percentages of 14Cinrhi-
zosphere respiration and in the soil residues are both
significantly increased by nitrogen fertilization (mean
of increase is C25% and C82%, respectively). This
suggests that relative to C exported by shoots to be-
lowground, N fertilisation increases rhizodeposition.
This hypothesis has some theoretical basis. Indeed, if
rhizosphere microorganisms are in competition with
plant roots for mineral N (Kaye and Hart,1997), a
supply of nitrogen would be expected to stimulate
microbial growth and consequently to increase the flux
of passive exudation. Moreover, nitrogen deficiencies
were reported to affect root morphology by reducing
the branching (Baligar et al.,1998), which may
have significant consequences on the production of
mucilage and the release of root cap cells and on exu-
dation. Thus, despite there are clues indicating that N
fertilization increases the percentage of belowground
C that is released from roots, the overall effect of N
fertilization on rhizodeposition is difficult to predict
because in parallel, nitrogen stimulates the total plant
growth and photosynthesis and reduces the percentage
of photoassimilates that are allocated belowground.
3.2.6 Atmospheric CO2Concentration
The elevation in atmospheric CO2concentration con-
secutive to the use of fossil C has raised the question
as to C fluxes to the rhizosphere would be modified.
This is of particular importance for understanding nu-
trient cycling and C sequestration in soil under ele-
vated atmospheric CO2. We report here on 24 data sets
related to 14C distribution within the plant and to the
soil under elevated CO2(Table 12). Pulse and contin-
uous labellings are equally represented. Studies con-
cerned both herbaceous plants (ryegrass, wheat, maize)
and trees (aspen and chestnut, data not shown). There
is no clear effect of elevated CO2on the partitioning
of assimilates to shoots and roots. However, in con-
tinuous labelling experiments, the 14C retrieved in the
rhizosphere respiration is significantly increased un-
der elevated CO2.C36%/. This is consistent with the
data reviewed by Zak et al. (2000), which evidence an
increase in soil and microbe respiration under elevated
CO2. Apart from that, the data do not indicate a clear
effect of elevated CO2on C partitioning to the rhizo-
sphere. This is not surprising because atmospheric CO2
is not a factor directly connected to the rhizosphere.
Any effect of atmospheric CO2enrichment on rhizode-
position is through plant growth in contrary to factors
such as the soil texture or the presence of microorgan-
isms that act more directly on the release of C from
roots. Soil nitrogen can be considered as intermedi-
ate because it stimulates both the growth of plant and
the growth of microorganisms. Hence, elevated CO2
can alter the partitioning of assimilates to the rhizo-
sphere through several mechanisms such as a change
in plant structure, itself depending on the plant species
(Pritchard et al.,1999), a modification of the root to
shoot ratio (Rogers et al.,1994), an alteration in root
morphology, a nutrient stress due to the stimulation of
plant growth, etc.
116 C. Nguyen
Table 11 Effect of soil nitrogen on labelled C partitioning between plant and belowground compartments. Effects are expressed as
relative variations, see the text for explanations about the calculation of the effects
Factor LabellingaReferences
NbRelative variation (%)
Mean CV of
mean Max Med Min PncPtdPme
Soil nitrogen (mg/kg)
Soil content or applied as fertilization
CBilles et al.
(1993);
Johansson
(1992a);
Liljeroth
et al. (1994);
Merckx et al.
(1987); Van
Ginkel et al.
(1997)
SHOOT 9 11 78 27 9 0 0:470 0:005
Min D0C ROOT914 79 0 13 32 0:712 0:005
Max D505 C RR 96186 12 821 0:795 0:145
Mean D152 C RES 915 253 78 28 39 0:001 0:070
Median D73 C ROOTBG 92380 10 124 0:059 0:453
CV D118 C RRBG 9 7 127 18 9 7 0:429 0:045
CRESBG931;330 99 20 30 0:001 0:508
P Henry et al.
(in press);
Mikan et al.
(2000);
Warembourg
and Estelrich
(2001)
SHOOT 19 36 96 109 23 7 0:032 0:001
Min D38.5 P ROOT 19 35 84 38 39 84 0:006 0:001
Max D970 P RR 19 41;028 96 164 0:210 0:677
Mean D694 P RES 19 38 209 236 31 69 0:292 0:052
Median D750 P ROOTBG 19 16 167 67 25 42 <0:001 0:001
CV D32 P RRBG 19 25 156 90 29 49 0:717 0:012
PRESBG19 82 111 260 50 51 0:262 0:001
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated to belowground
aCDcontinuous labelling, P Dpulse labelling
bNumber of partitioning coefficient sets
cP associated to Shapiro-Wilk test for normality
dP associated to student test for location Mu D0 for data normally distributed
eP associated to the non parametric sign test for location Median D0 for data non normally distributed
4 Outlooks
Tracer experiments are very useful tools for investi-
gating C fluxes from plant roots to the soil because
they allow separating root-derived C from the C of the
native soil organic matter. With such techniques, in-
vestigations on rhizodeposition can be performed on
plants growing in soil including the microflora, which
is more realistic than experiments in nutrient solution.
In counterpart, in soil, it is difficult to estimate the frac-
tion of rhizodeposits that is mineralised by microor-
ganisms and consequently, the amount of C released
from root cannot be determined in a reliable manner.
The partitioning of rhizosphere respiration between
root and microbial contributions is of particular impor-
tance if rhizodeposition is investigated to understand
processes that are mediated by microbes. In that case,
it is essential to evaluate how much energy is avail-
able to microorganisms to predict microbial growth
in the vicinity of roots. Several attempts have been
made to evaluate the rhizomicrobial contribution to rhi-
zosphere CO2(Cheng et al.,1993;Helal and Sauer-
beck,1989;Johansson,1992a;Kuzyakov et al.,1999;
Todorovic et al.,2001;VonWiren et al.,1996) but at
present time, none of them is fully satisfactory be-
cause all these studies rely on strong assumptions that
are difficult to test. As an alternative, metabolic activ-
ity (growth, maintenance) of rhizosphere microbes can
be determined or compared between different treat-
ments to investigate its relationships with root activ-
ity (Nguyen et al.,2002;Soderberg and Baath,1998).
However, quantification of root-derived C fluxes in non
sterile soil is undoubtedly a key point that needs fur-
ther investigations and methodological developments
for aiming at engineering the rhizosphere to manage
nutrient and pollutant cycling or to control soil borne
pathogens.
Rhizodeposits cover a wide range of compounds
that have very different characteristics in terms of in-
teractions with the soil matrix, availability to microbial
assimilation, chemical properties, etc. Moreover, the
release of root C into the root environment originates
Rhizodeposition of Organic C by Plant: Mechanisms and Controls 117
Table 12 Effect of atmospheric CO2on labelled C partitioning between plant and belowground compartments. Effects are ex-
pressed as relative variations, see the text for explanations about the calculation of the effects
Factor LabellingaReferences Nb
Relative variation (%)
Mean CV of
mean Max Med Min PncPtdPme
Atmospheric CO2(ppm)
CBilles et al.
(1993); Gorissen
et al. (1996); Van
Ginkel et al.
(1997); Whipps
(1985)
SHOOT 12 6158 13 10 18 0:460 0:051
Min D350 C ROOT 12 21 201 113 5 23 0:032 0:774
Max D800 C RR 12 36 132 107 40 61 0:734 0:024
Mean D633 C RES 12 22;459 126 11 50 0:005 0:388
Median D700 C ROOTBG 12 5 461 54 222 0:022 0:146
CV D24 C RRBG 12 24 201 113 16 69 0:996 0:112
CRESBG12 13 280 90 20 45 0:003 0:039
PPaterson et al.
(1996,1999);
Rattray et al.
(1995); Rouhier
et al. (1996);
Mikan et al.
(2000)
SHOOT 12 3505 25 428 0:490 0:507
Min D350 P ROOT 12 10 315 87 7 28 0:041 0:774
Max D720 P RR 12 2 2;656 115 587 0:304 0:899
Mean D477 P RES 12 43 169 215 19 23 0:024 0:388
Median D450 P ROOTBG 12 3307 15 322 0:811 0:284
CV D32 P RRBG 12 41;269 129 986 0:053 0:790
PRESBG12 21 166 99 8 17 0:087 0:061
RR and RES are %14C allocated to rhizosphere respiration and soil residues, respectively
The suffix BG is used when partitioning coefficients are expressed as percentages of 14C allocated to belowground
aCDcontinuous labelling, P Dpulse labelling
bNumber of partitioning coefficient sets
cP associated to Shapiro-Wilk test for normality
dP associated to student test for location Mu D0 for data normally distributed
eP associated to the non parametric sign test for location MedianD0 for data non normally distributed
in various mechanisms (i.e. passive diffusion of solutes
to the soil solution, active secretion of molecules,
senescence of root tissues), the distribution and
intensity of which are not homogenously distributed
along the root. This complexity is well illustrated by
the great difficulty to propose a nomenclature for the
rhizodeposits. Consequently, the composition of C
released from roots is virtually extremely variable.
Indeed, the composition of rhizodeposits depends on
the relative proportion of each category (exudates,
secretion, senescing tissues) as well as of the intrinsic
composition of each of these categories. For example,
nutrient or toxicity stress is know to significantly
increase the concentration of organic acids in root
exudates (Jones,1998). It is thus crucial to investigate
in more detail the mechanisms by which root C is
released into the soil. For example, the production
of root cap cells and mucilage has been extensively
studied in vitro, under experimental conditions that
probably increase the phenomena. Very little is known
about environmental control of rhizodeposition by root
apices in soil conditions. If the mucilage sticks to the
root cap even at soil water potentials close to 0.01 MPa,
the continuous production of slime and the release of
root cap cells might not be as important as suggested
by laboratory investigations. There is also great debate
as to determine whether the plant does have a control
on the amount of C that passively diffuse to the soil
solution. The ATPase transporters, which can actively
reabsorb solutes in vitro, provide a mechanism by
which the root can virtually modulate exudation.
However, does the plant regulate the flux of exudates
by controlling the number of these transporters and
their activity? Research aimed at understanding the
regulation of these proteins is particularly relevant.
Indeed, these transporters would offer the opportunity
to manipulate the flow of exudation both in term
of quantity by over expressing or inhibiting the
transporters synthesis and in term of quality by acting
specifically on target transporters and thus on changing
the exudation flux of a particular compound.
The spatial heterogeneity of rhizodeposition along
a root segment outlines the need to link investigations
on rhizodeposition to the root. From a theoretical point
of view, the branching pattern, which determines the
number of apices, would be expected to have signifi-
cant effect on the number of slough-off root cap cells
as well as on mucilage production and on the release
118 C. Nguyen
of exudates if their diffusion is more important at the
root tips as it was observed in maize. Moreover, the ex-
udation, the release of border cells and the senescence
of epidermis is proportional to the root radius. Hence,
it is necessary to determine if in soil root morphology
has indeed significant effects on rhizodeposition.
The “rhizosphere effect” observed experimentally
for numerous soil processes mediated by the mi-
croflora is frequently related to the greater microbial
abundance at the soil-root interface compared to bulk
soil. Therefore, a major goal for investigations on rhi-
zodeposition is to predict microbial growth in the root
environment (Blagodatsky and Richter,1998;Darrah,
1991,1991;Newman and Watson,1977;Toal et al.,
2000). In the last decade, the development of tech-
niques to establish microbial fingerprints evidenced
that the structure of rhizosphere communities was both
physiologically and genetically different from that
of bulk soil and different between plant species. The
relationships between size of the microflora, structure
of microbial communities and functions performed by
them is far from being elucidated and there is a rele-
vant need to investigate the factors that determine the
structure of microbial communities in the rhizosphere.
Among them, rhizodeposits have been demonstrated to
be relevant (Benizri et al.,2002;Griffiths et al.,1999).
Since growth of soil microbes is generally limited by
availability of C, it can reasonably be assumed that
the dynamics of microbial community structure might
derive from the competitive ability of rhizosphere
microorganisms with respects to the amount of avail-
able C. On the other hand, plant roots are “chemical
factories” that synthesize a wide variety of secondary
metabolites (Bais et al.,2001), which are biologically
active and which might orient the dynamics of mi-
crobial communities. Root-microbe interactions might
not only be governed by trophic competition between
microorganisms for rhizodeposits and by sophisticated
signalling involved in symbiosis process. Allelopathy,
which can play a significant role in the dynamics
of plant community structure, might also contribute
to determine the structure of rhizosphere microbial
communities. The chemical diversity of secondary
metabolites released into the rhizosphere is probably
large and rhizodeposition of such compounds offers an
exciting area of investigations and additional outlook
to the use of the plant for engineering the rhizosphere.
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