Download Branching out in new directions: the control of root

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Branching out in new directions: the
control of root architecture by lateral
root formation
Author for correspondence:
J. C. Coates
Tel: +44 121 414 5478
Fax: +44 121 414 5925
Email: [email protected]
C. Nibau*, D. J. Gibbs* and J. C. Coates
School of Biosciences, University of Birmingham, Edgbaston, Birmingham B15 2TT, UK
Received: 21 December 2007
Accepted: 14 March 2008
Contents
Summary
595
I.
Background
595
II.
Formation of lateral roots
596
III.
Endogenous factors regulating the stages of lateral
root development
V.
Transcriptomic studies to identify potential new
regulators of lateral root development
VI. Conclusions and future challenges
IV. Plasticity: modification of lateral root development
by the environment
597
608
608
Acknowledgements
608
References
609
603
Summary
Key words: abiotic stress, biotic stress,
lateral root development, nutrients, plant
hormones, root system architecture,
transcriptomics.
Plant roots are required for the acquisition of water and nutrients, for responses to
abiotic and biotic signals in the soil, and to anchor the plant in the ground. Controlling
plant root architecture is a fundamental part of plant development and evolution,
enabling a plant to respond to changing environmental conditions and allowing plants
to survive in different ecological niches. Variations in the size, shape and surface area of
plant root systems are brought about largely by variations in root branching. Much
is known about how root branching is controlled both by intracellular signalling pathays and by environmental signals. Here, we will review this knowledge, with particular
emphasis on recent advances in the field that open new and exciting areas of research.
New Phytologist (2008) 179: 595–614
© The Authors (2008). Journal compilation © New Phytologist (2008)
doi: 10.1111/j.1469-8137.2008.02472.x
I. Background
A plant’s root system is the site of water and nutrient uptake
from the soil, a sensor of abiotic and biotic stresses, and a
*These authors contributed equally to this work.
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structural anchor to support the shoot. The root system
communicates with the shoot, and the shoot in turn sends
signals to the roots. A plant root system initially consists of a
primary root (PR) formed during embryogenesis that has
dividing cells in a meristem at its tip. As the seedling develops,
certain other cells within the PR acquire the capability to
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Fig. 1 Components of the root system. (a) A typical dicot
(e.g. Arabidopsis) seedling root system, consisting of a primary
root (PR) originating from the embryo, lateral roots (LR) branching
out from the PR during seedling development, and root hairs
(RH) that originate from PR epidermal (Epi) cells (shown at
higher magnification to the right (inset)). Ultimately, the LRs
will undergo higher-order branching to form secondary and
tertiary LRs. Adventitious roots (AR) form at the shoot–root junction.
(b) A typical cereal (e.g. rice, maize) seedling root system consisting
of a primary root (PR) originating from the embryo, seminal roots (SR)
that originate postembryonically close to the top of the primary root,
and crown roots (CR) that originate from the stem. PR, SR and CR all
form LR and undergo higher-order branching.
divide, eventually forming new roots, called lateral roots (LRs)
(Fig. 1a). These branch out from the PR, greatly increasing
the total surface area and mechanical strength of the root
system and allow the plant to explore the soil environment.
Ultimately, millions of higher-order root branches can form,
resulting in hundreds of miles of root system in a small area
of soil (Dittmer, 1937). New roots, called adventitious roots
(AR), can also be formed postembryonically at the shoot–root
junction, optimizing the exploration of the upper soil layers
(Fig. 1a). In cereals such as rice and maize, root structure
becomes more complex, with the formation of additional
shoot-borne and postembryonic roots, which in turn undergo
higher-order branching (Hochholdinger et al., 2004; Hochholdinger & Zimmermann, 2008; Fig. 1b).
The root system architecture (RSA) of plants varies hugely
between species and also shows extensive natural variation
within species, reflecting the plethora of environments in
which plants can grow (Cannon, 1949; Loudet et al., 2005;
Osmont et al., 2007). Root system architecture manipulation
is instrumental in the domestication and breeding of crop
plants, because using water and nutrients from the soil in the
most efficient manner affects a plant’s ability to survive in
stressful or poor soils. Changes in RSA can therefore have
huge impacts on the final yield of a crop (reviewed in de
Dorlodot et al., 2007). Of the factors that control total RSA,
LR formation and growth is one of the most important.
Many of the hormonal and environmental signals affecting
LR development also affect other components that have a
bearing on RSA and the overall root surface area, namely, root
New Phytologist (2008) 179: 595–614
hair development, primary root (PR) growth and AR formation.
However, an extensive analysis of how these structures are
controlled is outside the scope of the present review and the
reader is referred to several other excellent reviews (Dolan &
Costa, 2001; Carol & Dolan, 2002; Scheres et al., 2002; Casson
& Lindsey, 2003; Hochholdinger et al., 2004; Samaj et al., 2004;
Serna, 2005; Scheres, 2007). Moreover, colonization of certain
plant roots by symbiotic bacteria or fungi leads to the formation
of modified LRs (root nodules, mycorrhizas or proteoid roots)
that carry out specialized functions such as nutrient acquisition
(Oldroyd & Downie, 2004, 2006; Autran et al., 2006).
In addition to signals that regulate many components of
RSA (and sometimes also shoot development), there is
mounting evidence that some signalling networks are specific
for LR formation (Rogg et al., 2001; Hochholdinger et al.,
2004; Loudet et al., 2005; Coates et al., 2006), potentially
highlighting novel strategies for manipulating root branching
in crop plants.
Because of the major contribution they play in the control
of RSA, this review focuses on LRs: how they arise and develop.
It will pay particular attention to recent molecular and ‘omic’
developments that highlight the huge variety of genes,
proteins and mechanisms that interact together to coordinate
a process so central to plant development and survival.
II. Formation of lateral roots
In flowering plants and gymnosperms, LRs initiate from a
specialized cell layer in the PR called the pericycle. The
pericycle is the outermost cell layer of the vascular cylinder
and consists of two distinct cell types corresponding to the
underlying vasculature (Dubrovsky & Rost, 2005; Parizot
et al., 2008; Fig. 2a,b). In Arabidopsis and most other dicots,
LRs are formed only from pericycle cells overlying the
developing xylem tissue (the xylem pole pericycle) (Fig 2b). In
other species, particularly cereals such as maize, rice and wheat,
LRs arise specifically from the phloem pole pericycle, with
additional contributions from the endodermis (De Smet et al.,
2006a; Hochholdinger & Zimmermann, 2008; Fig. 2b).
Insights into the evolution of multicellular, branched root
systems come from ‘ancient’ plants. In a vascular nonseed
plant, the fern Ceratopteris, LRs arise from the endodermis
and may be regulated differently from those in flowering
plants (Hou et al., 2004). The bryophyte moss Physcomitrella
patens revealed a very ancient mechanism controlling the
development of tissues with a rooting function (Menand et al.,
2007). Physcomitrella possesses putative homologues of known
Arabidopsis LR regulators, many of which have no assigned
function (e.g. Axtell et al., 2007; Rensing et al., 2008).
Lateral root formation consists of four key stages: (i)
stimulation and dedifferentiation of pericycle founder cells;
(ii) cell cycle re-entry and asymmetric cell divisions to give rise
to a lateral root primordium (LRP); (iii) LRP emergence
through the outer layers of the PR via cell expansion; and (iv)
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Fig. 2 Root anatomy. (a) Longitudinal
section through an Arabidopsis primary
root tip, showing the different cell types.
LRC, lateral root cap (which is absent further
up the root); Epi, epidermis (which is the
outermost layer of the root above the
root tip); Co, cortex; En, endodermis; P,
pericycle; Vasc, vasculature (xylem and
phloem); QC, quiescent centre (maintains
the neighbouring stem cell population). (b)
Transverse section through an Arabidopsis
primary root. Epi, epidermis; RH, root hair;
Co, cortex; En, endodermis; P, pericycle;
XPP, xylem pole pericycle (the pericycle
cells adjacent to the xylem tissue, from
which lateral roots arise); Xy, xylem; Ph,
phloem. In monocots, lateral roots arise
from the phloem pole pericycle.
Fig. 3 Aspects of auxin signalling during lateral root (LR) development. (a) A pulse of auxin (light grey) in the basal meristem (BM) primes a
pericycle cell (dark grey) to become competent to form a lateral root initial cell. (b) Cells (white) leaving the basal meristem between cyclical
auxin maxima are not specified to become LR initials. (c) The first primed pericycle cell arrives at a point where it can initiate LR development;
meanwhile another pericycle cell (dark grey) is primed in the basal meristem by the subsequent auxin pulse. (d) Lateral root initiation begins
with auxin-induced IAA14 degradation. This allows activation of the ARF7 and ARF19 transcription factors, which activate expression of LBD/
ASL genes. LBD/ASL proteins in turn activate cell cycle genes and cell patterning genes, enabling formation of a new lateral root primordium
(LRP). Auxin also activates transcription of NAC1 to stimulate LR initiation, and at the same time induces expression of two ubiquitin ligases,
CEGUENDO and SINAT5, which feed back to attenuate the auxin response.
activation of the LR meristem that recapitulates PR growth
(Celenza et al., 1995; Cheng et al., 1995; Laskowski et al.,
1995; Malamy & Benfey, 1997).
III. Endogenous factors regulating the stages of
lateral root development
Underpinning each stage of LR development is the hormone
auxin (Casimiro et al., 2003; Woodward & Bartel, 2005;
Figs 3a–d and 4a–d). Comprehensive studies using a LR-
inducible system revealed that over 10% of the Arabidopsis
seedling root transcriptome was affected by treatment with
auxin (Himanen et al., 2002; Vanneste et al., 2005).
Auxin maxima appear at LR initiation sites and also later
during emergence and elongation (see section III.1). Auxin
‘hot spots’ within the root arise as a result of the regulated
positioning of auxin transporters within cells, in a process
conserved between lateral organ formation in the root and in
the shoot (Benkova et al., 2003). Interestingly, auxin signalling
regulates the differential positioning of auxin efflux carriers
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Fig. 4 Lateral root development in Arabidopsis shown in longitudinal section. P, pericycle; En, endodermis; Co, cortex; Epi, epidermis. (a) Early
initiation – a founder xylem pole pericycle cell (dark grey) undergoes initial anticlinal cell divisions (perpendicular to the surface of the root). (b)
Periclinal cell divisions (parallel to the surface of the root) begin and the lateral root primordium (LRP) begins to grow. (c) The LRP undergoes
further organized cell divisions and begins to emerge through the outer cell layers of the primary root, resulting in cell separation (asterisks). (d)
The new lateral root is fully emerged and its new meristem is activated (dark grey star). It will continue to grow and elongate. At each stage,
the effect of various key plant hormones is indicated. ABA, abscisic acid; BR, brassinosteroids.
and, consequently, the direction of auxin flow (Sauer et al.,
2006). This effect is mediated by the activity of VPS29, a
membrane-trafficking component that is involved in the
recycling of cargo molecules. Together with other proteins,
VPS29 mediates the dynamic arrangement of auxin efflux
carriers in response to auxin (Jaillais et al., 2007). The regulated
interplay between auxin transport and signalling is critical for
all stages of LR development, and many of the signals regulating
RSA impinge upon this pathway.
Many Arabidopsis and cereal mutants affecting auxin production, transport and metabolism have LR defects: their
involvement in LR formation has been described extensively
elsewhere (Casimiro et al., 2003; Woodward & Bartel, 2005;
Fukaki et al., 2007). Lack of detailed characterization of many
of these mutants prevents identification of the particular stage
of LR development at which they act (De Smet et al., 2006a).
Exhaustive description of all the proteins involved in
auxin-dependent lateral organ formation is beyond the scope
of this review and the reader is directed to recent reviews in
this area (De Smet et al., 2006a; Teale et al., 2006; De Smet
& Jurgens, 2007).
Other hormone pathways are also involved in the regulation
of LR formation, and recent research provides new insight
into these pathways. Below we will outline how plant hormones, with particular emphasis on auxin, interact with various
cellular processes to control each stage of LR development.
1. Lateral root initiation – stimulation of cell cycle
proliferation in the pericycle
In Arabidopsis, the xylem pole pericycle cells, from which LRs
arise, are smaller than other pericycle cells, indicating
New Phytologist (2008) 179: 595–614
differential cell cycle regulation between pericycle cell types.
Normally, not all xylem pole pericycle cells form LRP,
indicating that multiple levels of control occur in these cells
(Beeckman et al., 2001). However, exogenous application of
auxin can activate the whole pericycle to form LRPs, whereas
the application of auxin transport inhibitors blocks LR
formation without loss of pericycle identity (Casimiro et al.,
2001; Himanen et al., 2002). Therefore, all the cells within
the pericycle retain the ability to form LRs but only some of
them do so. It is thus suggested that the coordinated action of
auxin transport and signalling, cell cycle regulators and novel
root-specific proteins is necessary for LR initiation to occur.
Lateral root initiation requires auxin and regulated protein
degradation Auxin signalling during LR initiation is closely
coupled with regulated protein degradation (Fig. 3d). Proteins
are targeted to the cellular degradation machinery, the
proteasome, by the addition of a chain of ubiquitin monomers.
The process requires a ubiquitin-activating enzyme (E1), a
ubiquitin-conjugating enzyme (E2) and a ubiquitin-protein
ligase (E3), which transfers ubiquitin from the E2 to the
target (Petroski & Deshaies, 2005). Some E3 ubiquitin ligases
consist of multiprotein complexes, and SKP1-CULLIN1-F-box
(SCF) E3 ligases contain F-box protein subunits that confer
specificity, binding to particular target proteins.
Auxin receptors are a family of F-box-containing proteins
known as TIR1 and AFB1–3 (Dharmasiri et al., 2005a,b;
Kepinski & Leyser, 2005). It is thus not surprising that
mutants in components of the SCF complex and its associated
proteins have altered LR phenotypes (Gray et al., 1999; Hellmann et al., 2003; Bostick et al., 2004; Chuang et al., 2004;
Woodward et al., 2007).
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Auxin binding to TIR1/AFBs allows them to interact with
AUX/IAA proteins and target them for degradation. AUX/IAA
proteins are transcriptional repressors that dimerize with
auxin response factor (ARF) transcription factors, preventing
the latter from binding to promoter elements in auxinresponsive genes. Thus, auxin-induced degradation of AUX/
IAAs enables ARFs to activate auxin-responsive transcription
(Gray et al., 2001; Dharmasiri et al., 2005a,b; Kepinski &
Leyser, 2005). AUX/IAAs and ARFs exist as large, functionally
redundant protein families (Okushima et al., 2005; Overvoorde
et al., 2005).
One of the most important AUX/IAA proteins for LR
initiation is SLR1/IAA14. As a result of the stabilization
of IAA14, a gain-of-function slr1 mutant does not form LRs
(Fukaki et al., 2002). In wild-type plants, auxin triggers
the degradation of IAA14, enabling ARF7 and ARF19 to
activate transcription of LATERAL ORGAN BOUNDARIES
DOMAIN/ASYMMETRIC LEAVES LIKE (LBD/ASL) genes
(Fukaki et al., 2005; Okushima et al., 2007; and Fig. 3d).
LBD/ASL proteins, in turn, activate the transcription of cell
proliferation and patterning genes (Okushima et al., 2007).
In maize and rice, LBD genes regulate shoot-borne root
formation rather than LRs (Taramino et al., 2007; Hochholdinger & Zimmermann, 2008). ARF7 also interacts with
a MYB transcription factor that provides a link among auxin,
LR initiation and environmental responses (Shin et al., 2007;
and section IV.4).
Comparison of the auxin-induced transcriptomes of
wild-type and slr1 roots identified 913 specific ‘LR initiation’
genes that function downstream of the auxin/slr1 signalling
pathway. Many of these are cell cycle-associated genes or cell
division-associated genes, and genes involved in auxin signalling,
transport or metabolism. Other over-represented functional
categories include macromolecular biosynthesis, ribosome
biogenesis and DNA synthesis (Vanneste et al., 2005).
IAA28 is also important for LR initiation. The gain-offunction mutant iaa28 forms fewer LRs than the wild type:
IAA28 is degraded by auxin and represses auxin-induced
LR-formation genes. However, IAA28 mRNA levels are
repressed by auxin, indicating a complex regulation of IAA28
during auxin signalling (Rogg et al., 2001; Dreher et al.,
2006). The iaa28 mutant is also resistant to exogenous cytokinins and ethylene, suggesting an integration point for other
hormone pathways.
The VIER F-BOX PROTEINE (VFB) F-box proteins
are also important for LR formation in Arabidopsis. Mutants
deficient in VFB function have reduced LR formation. Microarray analysis demonstrated that loss of VFB function leads to
altered expression of both auxin-responsive genes and cell
wall-remodelling genes (Schwager et al., 2007). Despite this,
vfb mutant plants maintain full sensitivity to exogenously
applied auxin. VFBs may regulate auxin-induced gene expression, and consequently LR formation, by a pathway
independent of the auxin receptor TIR1 (Schwager et al.,
Review
2007). It will be important to determine whether IAA14/
SLR1 stability or cell cycle gene expression is affected in vfb
mutants.
The NAC1 transcription factor promotes LR initiation
(Xie et al., 2000) and may bind auxin-responsive promoters
to transmit the auxin signal (Fig. 3d). Interestingly, NAC1
overexpression can rescue the reduced LR phenotype of tir1
auxin receptor mutants. NAC1 is tightly regulated: NAC1
expression is induced within 30 min of auxin application,
suggesting that NAC1 may be an early auxin-responsive gene.
Auxin also induces the expression (albeit more slowly) of
SINAT5, a RING-finger ubiquitin E3 ligase (Fig. 3d). SINAT5
promotes NAC1 ubiquitination and subsequent degradation
(Xie et al., 2002). It will be interesting to determine if auxin
binds directly to SINAT5 in the SINAT5–NAC1 complex.
Yet another ubiquitin ligase involved in LR initiation is
XBAT32 (Nodzon et al., 2004). XBAT32 is a RING-finger
protein highly expressed in the vascular system close to sites of
LR initiation. Plants lacking XBAT32 develop fewer LRs than
wild-type plants and have reduced cell division in the pericycle.
XBAT32 may be involved in auxin transport: loss of XBAT32
may lead to suboptimal auxin levels for LR initiation (Nodzon
et al., 2004).
As only certain pericycle cells usually give rise to LRs, it is
crucial that auxin signals are tightly regulated. Interestingly,
auxin stimulates the transcription of ubiquitin ligases that
repress auxin signals, providing an elegant feedback mechanism
to maintain auxin sensitivity in the pericycle. The F-box
protein CEGENDUO (CEG) is a negative regulator of LR
formation whose transcription is induced by auxin (Dong
et al., 2006; and Fig. 3d). Further studies are needed to clarify
its role LR initiation.
It is thus clear that the action of auxin during LR initiation
depends heavily on the ubiquitin-proteasome pathway, both
to transduce signals by degrading repressors and also to reset
the system by destroying activators when they are no longer
needed. Protein degradation allows for rapid changes in
response to the ever-changing environment, as well as providing fine-tuning to sustained signals.
Which pericycle cells? Questions remain about where, when
and which pericycle cells are primed to become LR initiation
sites. In the last year, this problem has been addressed by new
molecular genetic and mathematical modelling studies.
De Smet et al. (2007) showed that the position of Arabidopsis
LR formation is determined in a region at the transition between
the meristem and the elongation zone, called the basal meristem
(Fig. 3a–c). Lateral roots occurred in a regularly spaced alternating
left–right pattern correlating with gravity-induced root waving.
Both responses are dependent on the auxin influx transporter,
AUX1. Furthermore, auxin responsiveness at the basal meristem
oscillates in a periodic manner, correlating with the timing of
LR formation. This, together with the observation of a lateral
gradient of auxin responsiveness with a maximum in protoxylem
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New Phytologist (2008) 179: 595–614
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cells, led the authors to suggest that auxin accumulation
alone is sufficient for the priming of founder cells (De Smet
et al., 2007). Consequently, one can suggest that all the other
factors involved in LR formation will act downstream of the
auxin signal. However, several factors have been suggested to
regulate LR formation independently of auxin (see section
III.1 ‘Hormone-independent signalling pathways that regulate
lateral root initiation’). Targeted manipulation of these genes
in pericycle cells in the basal meristem is necessary to clarify
this conundrum. In addition, it is well known that environmental factors can tap into the LR developmental program
and alter root architecture in regions outside the basal meristem
(see section IV).
Support for the co-regulation of LR formation and gravitropism came from a mathematical model suggesting that
gravistimulation concentrates auxin at a certain point in the
root, allowing the auxin threshold necessary for LR formation
to be reached. Lateral root initiation would, in turn, consume
the auxin pool in that area, preventing new LR initiation until
the pool had been refilled: this would be accelerated by a new
gravistimulation (Lucas et al., 2007). Two main ideas came
out of this study: first, there is an endogenous mechanism
regulating the periodicity of LR formation, revealed by the
existence of a minimum and maximum time between two
successive LR initiations. Second, this endogenous system is
sensitive to external cues such as gravity (Lucas et al., 2007).
This would provide increased plasticity for the root system to
adapt to new soil conditions.
Interestingly, the developmental window for LR initiation
in Arabidopsis displays natural variation between accessions
(Dubrovsky et al., 2006), which may indicate adaptation of
the system to different environmental niches. In addition,
LRP initiation and emergence are separable processes, again
providing greater plasticity to the root system (Dubrovsky
et al., 2006). The mathematical model used by Lucas et al.
(2007) suggests that LR formation in gravistimulated areas
may also optimize soil exploration. It will be interesting to
determine if biotic and abiotic factors that alter RSA also have
an effect on gravitropism-stimulated LR formation.
It is important to move this type of research beyond
Arabidopsis to agriculturally relevant plants, especially as the
mechanisms at work in crop plants may differ from those
in Arabidopsis. In many grasses, LRs initiate in phloem pole
pericycle cells and, because of varying root organization and
growth rates, the timing and spacing of LR initiation is also
different (Dubrovsky et al., 2006; Dembinsky et al., 2007).
Interplay between the cell cycle and auxin signalling It is
generally accepted that pericycle cells are arrested in the G1
phase of the cell cycle. Those pericycle cells that will give rise
to a LR proceed through S phase and arrest in G2. Lateral
root-inducing signals stimulate these cells to undergo
proliferative cell divisions (Beeckman et al., 2001). Cell cycle
re-entry requires changes in chromatin structure, increasing
New Phytologist (2008) 179: 595–614
the proportion of active chromatin in the genome (De
Veylder et al., 2007). Indeed, a chromatin remodelling factor
mutant has perturbed LR initiation (Fukaki et al., 2006). Cell
cycle progression from G1 to S requires the activity of the
retinoblastoma (RB)-E2F pathway (del Pozo et al., 2006; De
Veylder et al., 2007). Progression from G2 to M is regulated
by the opposing activity of B-type cyclin-dependent kinases
(CDKs) and CDK inhibitor proteins (KRPs) (Wang et al.,
1997; De Veylder et al., 2001; Verkest et al., 2005). Many cell
cycle components are transcriptionally regulated by auxin
(Himanen et al., 2002; Vanneste et al., 2005). Another level
of regulation involves cell cycle protein degradation (Verkest
et al., 2005; del Pozo et al., 2006).
In tomato, nitric oxide is required in the early stages of LR
formation to regulate the expression of cell cycle genes, downstream of the auxin signal (Correa-Aragunde et al., 2006).
Nitric oxide is also induced in Arabidopsis LRP by the auxin,
indole-3-butyric acid (Kolbert et al., 2007).
Despite the importance of the cell cycle in LR initiation,
increasing the mitotic index in roots or forcing excessive cell
divisions in the pericycle does not stimulate LR initiation or
morphogenesis (Vanneste et al., 2005; Wang et al., 2006).
Thus, pericycle cell divisions can be uncoupled from LRP
formation, and LR initiation seems to require the simultaneous
activation of cell cycle and cell fate genes triggered by auxininduced degradation of the SLR/IAA14 protein (Vanneste
et al., 2005). Conversely, although cell division and LR
morphogenesis are both controlled by auxin signalling, the
processes are regulated independently, as shown by tomato
diageotropica (dgt) mutants that have a number of auxin-related
phenotypes (including a lack of LRs) but have normal root
cell identities and patterning. dgt mutant pericycle cells maintain
their full proliferative capacity, but no LRs are formed, even
in the presence of exogenous auxin, which instead stimulates
further pericycle divisions to form multiple cell layers
(Ivanchenko et al., 2006). A similar mechanism operates in
Arabidopsis, as demonstrated by the wol mutant, which forms
very few LRs even in the presence of auxin (Parizot et al.,
2008). Close analysis of the pericycle cells in the wol mutant
showed that they express pericycle protoxylem markers and
are able to divide in response to auxin but no LRPs are formed
(Parizot et al., 2008). This again shows that cell cycle activation
is not sufficient for LR initiation to occur.
Heterotrimeric G-proteins may integrate auxin signalling and
cell cycle inputs during root branching as well as other developmental processes (Ullah et al., 2001, 2003; Chen et al., 2006b;
Trusov et al., 2007). Mutations in the β or γ G-protein
subunits show increased cell division and increased LR formation, and the normal function of Gβ and Gγ may be to attenuate
auxin signalling (Ullah et al., 2003; Trusov et al., 2007).
Other hormones affecting LR development In addition to
auxin, other hormone signals are important for LR initiation (Fig. 4a–d). Traditionally, cytokinin is thought to act
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antagonistically to auxin in many developmental processes:
indeed, cytokinin is a negative regulator of LR formation in
many plant species, including Arabidopsis, Medicago, tobacco
and rice (Werner et al., 2003; Rani Debi et al., 2005;
Gonzalez-Rizzo et al., 2006; Li et al., 2006b; Laplaze et al.,
2007). Plants with decreased cytokinin content have increased
LR numbers (Werner et al., 2001, 2003; To et al., 2004;
Mason et al., 2005; Gonzalez-Rizzo et al., 2006; Riefler et al.,
2006), whereas exogenous cytokinin inhibits LR initiation by
preventing pericycle cell cycle re-entry (Li et al., 2006b). An
elegant study by Laplaze et al. (2007) showed that exogenous
cytokinin disrupts both LR initiation and the organization of
cell divisions within developing LRPs: these defects cannot be
rescued by auxin. Targeted expression of cytokinin biosynthetic
and catabolic enzymes in specific cell types demonstrated that
cytokinin activity is required very early in the LR formation
process (Laplaze et al., 2007). Importantly, disrupting cytokinin
signalling in xylem pole pericycle cells leads to perturbation
of the auxin maximum in developing LRPs as a result of the
reduced expression of PIN auxin transporter genes and
mislocalization of PIN proteins (Laplaze et al., 2007).
Both ethylene and brassinosteroids affect LR formation via
an auxin-dependent pathway (Bao et al., 2004; Stepanova
et al., 2005). In rice, a casein kinase 1 gene, OsCKI, is upregulated by both brassinosteroid and abscisic acid (ABA) and
promotes lateral and AR formation as well as cell elongation.
OsCKI may affect LR development by regulating endogenous
auxin levels (Liu et al., 2003). Transcriptomic analysis of
OsCKI-deficient plants revealed alteration of several signalling,
developmental, transcriptional and metabolic genes (Liu
et al., 2003).
Unequivocal evidence for a role of ABA in LR initiation is
not available. However, the ABA-insensitive mutant abi3
shows decreased sensitivity to auxin-induced LR initiation
(Brady et al., 2003). Furthermore, 9-cis-epoxycarotenoid
dioxygenase genes (involved in ABA biosynthesis) are expressed
in pericycle cells surrounding LR initiation sites (Tan et al.,
2003). It is tempting to suggest that ABA may restrain cell
proliferation outside the LR initiation site, although ABA
being produced as a result of the stress caused by LR emergence
cannot be excluded (De Smet et al., 2006b). Indeed, ABA
upregulates the expression of KRP1, a cell cycle inhibitor
(Wang et al., 1998). Some auxin-induced LR-initiation genes
had previously been described as ABA-repressed (Vanneste
et al., 2005). Among these, AUXIN-INDUCED IN ROOT
CULTURES 12 (AIR12) and IAA19 function in LR formation
(Neuteboom et al., 1999; Tatematsu et al., 2004). Thus, ABA
and auxin could have an antagonistic effect on LR initiation
(Fig. 4a). Interestingly, the KNAT1 homeobox transcription
factor is expressed at the base of LR primordia (Truernit et al.,
2006) and is auxin-induced and ABA-repressed in LR primordia (Soucek et al., 2007), suggesting a possible point of
integration for the two signals. Further research is needed to
establish ABA as an inhibitor of LR initiation in Arabidopsis.
Review
Curiously, ABA seems to stimulate LR initiation in rice (Chen
et al., 2006a).
In addition to ‘classical’ plant hormones, several other signals
affect LR development both during initiation and at later
stages, in a variety of plant species. Salicylic acid promotes LR
initiation, emergence and growth, possibly via crosstalk
with cytokinin or auxin (Echevarria-Machado et al., 2007).
Melatonin promotes lateral and AR formation while decreasing root length, similarly to the effects of auxin (Arnao &
Hernandez-Ruiz, 2007). Alkamides are lipid-based secondary
metabolites that are novel regulators of plant growth and
development (Lopez-Bucio et al., 2006). They induce LR
initiation and growth in Arabidopsis (Ramirez-Chavez et al.,
2004), and regulate meristematic activity throughout the plant.
It is suggested that they regulate root pericycle cell activation,
possibly via cytokinin signalling (Lopez-Bucio et al., 2007).
Hormone-independent signalling pathways that regulate
lateral root initiation The best-characterized protein that
regulates LR initiation, independently of hormone signalling,
is ABERRANT LATERAL ROOT FORMATION 4 (ALF4).
The Arabidopsis alf4 mutant shows a complete absence of LRs,
even in the presence of auxin (Celenza et al., 1995). Cells
appear to be blocked at a premitotic stage of the cell cycle,
but the identity of the xylem pole pericycle itself is not
compromised. ALF4 is a nuclear-localized protein of unknown
function; a shorter protein generated by alternative splicing
localizes to the cytoplasm. Importantly, auxin has no effect on
ALF4 levels or intracellular localization (DiDonato et al.,
2004). In the current model, ALF4 maintains the pericycle in
a ‘competent’ state for cell division, allowing input from other
LR-inducing signals, including auxin (DiDonato et al., 2004).
The alf4 mutant maintains full responsiveness to auxin
inhibition of PR elongation, suggesting that LR formation is
not a simple recapitulation of the developmental program
producing PRs.
Other regulators of LR initiation, acting independently of
known pathways, are ARABIDILLO-1 and ARABIDILLO-2,
which act redundantly to promote LR initiation in Arabidopsis
(Coates et al., 2006). ARABIDILLOs are F-box proteins,
suggesting that they form ubiquitin E3 ligases. Importantly,
arabidillo mutants and ARABIDILLO-overexpressing plants
are able to respond to exogenous auxin similarly to wild-type
plants. In addition, auxin distribution in the root tip appears
to be normal in arabidillo mutants, and auxin does not affect
the nuclear localization of ARABIDILLO proteins (Coates
et al., 2006; C. Nibau, J. Coates, unpublished).
Transcriptomic comparison of wild-type, arabidillo mutant
and ARABIDILLO-overexpressing roots reveals changes in
some genes defined as pericycle-enriched and LR-enriched
(Birnbaum et al., 2003; Levesque et al., 2006; https://
www.genevestigator.ethz.ch/), but no strong overlaps with
other recent LR data sets defined by auxin induction, VFB
signalling, LR emergence or red light signalling (Vanneste
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New Phytologist (2008) 179: 595–614
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et al., 2005; Laskowski et al., 2006; Molas et al., 2006;
Schwager et al., 2007; J. Coates, unpublished). ARABIDILLOs might act very early in determining pericycle cell fate.
Given the recent suggestion that this determination occurs at
the basal meristem (section III.1 ‘Which pericycle cells?’) it
will be interesting to establish whether arabidillo mutants are
affected in this process.
2. Redifferentiation to form a new meristem that
recapitulates the root organization
Lateral root primordium formation and emergence A series
of well-characterized cell divisions gives rise to the LRP
(Malamy & Benfey, 1997). The coordinated pattern of cell
division is dependent on auxin signalling and on the activity
of the PUCHI gene. PUCHI is expressed in pericycle cells that
will form the LRP and in the LRP itself. PUCHI encodes an
APETALA2 (AP2) transcription factor that is upregulated by
auxin and acts downstream of auxin to restrict the area of cell
proliferation within the LRP. PUCHI is also necessary for
correct cell divisions within the LRP (Hirota et al., 2007). In
rice, the EL5 RING finger ubiquitin E3 ligase maintains
cell viability in the developing primordium. EL5 may act
downstream of auxin, cytokinin and JA to prevent meristematic
cell death (Koiwai et al., 2007). The identity of the target
proteins and the involvement of EL5 in LRP hormone
signalling pathways remain to be investigated.
Once an LRP has initiated, it must form a functional
meristem and emerge from within the parent root tissues. As
a result of rounds of cell division, the LRP increases in size,
forming a dome-shaped structure that penetrates the external
cell layers of the PR. This requires separation of cells in the
endodermis, cortex and epidermis for the passage of the LR to
the outside (Fig. 4c,d). This process must be tightly regulated,
as cell separation (particularly of the protective epidermal
layer) constitutes a risk to the plant, potentially allowing the
entry of pathogens from the soil into internal tissues.
Much less is known about how LRs emerge than how they
initiate. Changes in electrical potential occur around prospective
sites of LR emergence (Hamada et al., 1992) and auxin seems
to be required for LR emergence independently of its role
in LR initiation. Shoot-derived auxin is required for LR emergence in Arabidopsis until c. 10 d after germination (Bhalerao
et al., 2002), and auxin can induce root cell separation in
Arabidopsis (Boerjan et al., 1995; Laskowski et al., 1995).
Cell separation occurs via regulated activity of cell wallremodelling enzymes. Breakdown of pectin is particularly
important for cell separation, as the middle lamella between
adjacent cells is pectin-rich. Pectin is demethylated by pectin
methylesterases (PMEs) before its catabolism. Pectin breakdown
involves homogalacturonases called pectate lyases (PLAs).
Interestingly, during LR emergence, the pectin in the emerging
LR remains methylated, whereas the pectin in the overlying
parent root tissues becomes demethylated, possibly in prepa-
New Phytologist (2008) 179: 595–614
ration for its controlled breakdown as LRs emerge (Laskowski
et al., 2006). How this differential pectin methylation is fully
controlled remains an intriguing question.
Various Arabidopsis studies have shown that cell wallremodelling enzymes are induced by auxin in roots (Neuteboom
et al., 1999; Himanen et al., 2004; Vanneste et al., 2005;
Laskowski et al., 2006). Cell wall remodelling genes induced
by auxin include a PME, PLAs, and also an expansin and a
beta-xylosidase (Laskowski et al., 2006). AtPLA1 and AtPLA2
are both upregulated steadily for up to 24 h after only a 15-min
pulse of auxin: this response is blocked in the slr1/iaa14
mutant. In addition, expression of both AtPLAs is much
higher in LR initials than in the pericycle cells from which
they arise (Laskowski et al., 2006).
The polygalacturonase (PG) family of cell wall-degrading
enzymes may help to ‘prime’ the PR cells to separate ready for
LR emergence (Gonzalez-Carranza et al., 2007). An Arabidopsis
PG (PGAZAT) is expressed specifically in the cortical and
epidermal cells overlying the future site of LR emergence. A
pgazat insertion mutant has no obvious LR phenotype, but it
is likely that functional redundancy exists. Interestingly, root
PGAZAT expression is auxin-inducible (Gonzalez-Carranza
et al., 2007).
Activation of the lateral root meristem and lateral root
elongation Activation and maintenance of the LR meristem
requires polarized auxin transport to create an auxin maximum
at the tip of the LRP: this requires regulated activity of auxin
influx and efflux transporters. During LR development there
is an important change in the direction of auxin flow, brought
about by AUX/IAA-dependent repositioning of auxin efflux
carriers towards the tip of the newly formed LR. This results
in LR growth perpendicular to the PR (Benkova et al., 2003;
Sauer et al., 2006). The new LRP auxin maximum regulates
the activity of several transcription factors (Blilou et al., 2005).
It is proposed that regulated expression of known regulators
of PR meristem formation, such as the PLETHORA,
CLAVATA, SCARECROW and SHORT ROOT, is also
important for the maintenance of an active meristem in LRs
downstream of auxin (for a recent review see Scheres, 2007).
Because mutations in these genes severely impair PR growth,
their effect on LR development has not been investigated:
targeted overexpression and underexpression in LRs will
clarify this issue.
Abscisic acid can reversibly block meristem activation
postemergence by inhibiting the cell cycle gene expression
necessary for meristem activity, leading to LR growth arrest
(De Smet et al., 2003). This effect of ABA defines a new
auxin-independent checkpoint between LR emergence and
meristem activation, which may also be regulated by nitrate
levels (De Smet et al., 2003). In line with this observation, an
ABA receptor mutant is completely insensitive to ABA inhibition of LR development (Razem et al., 2006). No other
known ABA-insensitive mutants show insensitivity, suggesting
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Review
mutations in the auxin efflux transporter MDR1 cause nascent
LRs to arrest their growth (Wu et al., 2007). The ALF3 protein
elevates the levels of auxin at the LRP, probably by facilitating
auxin transport (Celenza et al., 1995). The auxin-induced
homeobox gene HAT2 may also modulate auxin distribution
within the primordium (Sawa et al., 2002).
Despite the fact that cytokinins inhibit LR initiation, they
have a positive effect on LR elongation in Arabidopsis and rice,
possibly via stimulation of cell cycle gene expression in an
auxin-independent process (Rani Debi et al., 2005; Li et al.,
2006b).
Fig. 5 Lateral root responses to nutrient deprivation. When nitrate
(N) levels are high, lateral root (LR) emergence and elongation is
represssed compared with normal conditions. Locally high levels of N
promote local LR proliferation. In low phosphate (P), primary root
growth ceases and LR density increases. In low sulphate (S), primary
root growth and lateral root density increase, with LRs originating
closer to the root tip. In low potassium (K), LR elongation is inhibited.
that ABA signalling during LR emergence involves specific
proteins (De Smet et al., 2003). ABI8, a plant-specific protein
of unknown function, is a possible novel signalling candidate.
abi8 mutant plants are less sensitive to ABA and, despite
being able to initiate LR, their LR meristem soon loses
competence to divide (Cheng et al., 2000; Brocard-Gifford
et al., 2004).
This checkpoint between LR emergence and meristem
activation provides an elegant way by which environmental,
nutritional and endogenous factors can modulate root architecture through ABA signalling (Signora et al., 2001; De Smet
et al., 2006b; and section IV). For example, stress-induced
oxylipin production affects LR development. Treatment of
Arabidopsis seedlings with 9-hydroxyoctadecatrienoic acid
(9-HOT), an oxylipin derivative, induces the accumulation of
arrested early stage LRPs, accompanied by the upregulation
of cell wall-associated genes (Vellosillo et al., 2007). The
not-responding to oxylipins2 (noxy2) mutant has more LRs
than the wild-type plant. 9-Hydroxyoctadecatrienoic acid and
related oxylipins are probably endogenous modulators of
LR emergence that may act via ABA signalling and they are
involved in RSA reprogramming in response to pathogen
infection (Vellosillo et al., 2007).
Interestingly, ABA appears to have the opposite effect on
LR emergence in legumes, stimulating LR formation in
Medicago (Liang & Harris, 2005). The Medicago latd mutant
has a reduced root surface area with short PRs, arrested LRPs
and disorganized meristems (Bright et al., 2005). The latd
phenotype can be at least partly rescued by the exogenous
application of ABA, and latd mutants seem to be impaired in
ABA perception or signalling (Liang et al., 2007).
Lateral root elongation occurs by cell division and elongation
from the meristem and is controlled by several factors. Auxin
transport and signalling are important in this process. Auxin
transport within the root is necessary for LR elongation, as
IV. Plasticity: modification of lateral root
development by the environment
Plants are sessile organisms that need to survive in a dynamic
environment. Consequently, their root systems need to
maintain plasticity to react to fluctuating abiotic and biotic
factors. Genetically identical plants can have very different
RSA when grown in varying environmental conditions. Plants
primarily respond to the abundance of macronutrients and
water to produce the best root network for optimum growth
and survival (Fig. 5). However, other exogenous factors, such
as plant–pathogen interactions, are also important for root
development. An overview of the current understanding
of how changing external conditions affect RSA is presented
here, with a particular focus on more recent advances.
1. Nitrogen availability and root system architecture:
local and global effects
Inorganic nitrogen Root adaptation to nitrogen levels is an
excellent example of a plants’ developmental plasticity. Nitrogen
is available in the soil as ammonia, nitrite, nitrate and organic
nitrogen. The abundance of these compounds is highly
variable and can have dramatic effects on LR development.
Species-specific differences in nitrogen responses are apparent:
LR length, number or both can be affected (Zhang & Forde,
1998; Linkohr et al., 2002; Boukcim et al., 2006).
Nitrate levels have strongly opposing effects on LR growth,
depending upon the context in which they occur. In low-nitrate
soils, patches of high nitrate have a localized stimulatory effect
on LR development in many species (Drew & Saker, 1975;
Zhang & Forde, 1998). However, where nitrate levels are
globally high (i.e. not growth limiting), LR growth is inhibited
(Zhang et al., 1999). Thus, there are two clear morphological
adaptations: a local stimulatory effect of exogenous nitrate
supply on LR elongation, and a systemic inhibitory effect of
high nitrate on LR meristem activation. This is caused by the
signalling effect of nitrate itself, rather than being a response
to downstream metabolites (Zhang & Forde, 1998). Some
species, including barley and cedar, but not Arabidopsis, are
also able to respond to a localized ammonium supply (Drew,
1975; Zhang et al., 1999; Boukcim et al., 2006).
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Key protein players specific to the ‘local nitrate’ response
include the Arabidopsis NITRATE REGULATED-1 (ANR1)
MADS-box transcription factor and the DUAL AFFINITY
NITRATE TRANSPORTER (AtNRT1.1). Downregulation
of ANR1 reduces LR stimulation in nitrogen-rich zones,
without compromising the overall nitrate-induced inhibition
of LRs (Zhang & Forde, 1998). Seven other MADS box genes
have a similar expression pattern to ANR1 under different
nitrate conditions (Gan et al., 2005). Three of these (AGL-16,
AGL-21 and SOC1) interact with ANR1, although whether
they represent nitrate signal transduction components is
unknown (de Folter et al., 2005).
AtNRT1.1 is induced by nitrate (Munos et al., 2004) and
Atnrt1.1 mutants exhibit a strongly decreased response to
local nitrate supply (Liu et al., 1999). Interestingly, this
reduced responsiveness is accompanied by a reduction of
ANR1 mRNA (Remans et al., 2006). Transporters have been
previously identified as nutrient sensors, but it is unclear
whether AtNRT1.1 is involved directly in nitrate sensing, or
in facilitating access of nitrate to another sensor.
High nitrogen inhibits LR development after emergence
but before meristem activation. This effect is reversible: transferring plants to nitrate-limiting media results in a release of
LR inhibition within 24 h, so plants can respond rapidly to
fluctuating environmental nitrate levels (Zhang & Forde,
1998). A high shoot nitrate status is important for the inhibitory response, and an Arabidopsis mutant lacking nitrate
reductase activity is hypersensitive to inhibition, suggesting
that systemic accumulation of nitrate causes LR inhibition
(Zhang et al., 1999).
How are nitrate responses regulated during LR development?
Various ABA-deficient Arabidopsis mutants have significantly
reduced levels of LR inhibition in abundant nitrate (Signora
et al., 2001). With both ABA and high nitrate, LRs are inhibited
immediately after meristem activation (Signora et al., 2001;
De Smet et al., 2003; and section III.1 ‘Other hormones
affecting LR development’). Arabidopsis LR ABA-insensitive
(LABI) mutants can still produce LRs in the presence of ABA:
they are also less sensitive to high nitrate, implying that the
inhibition of LRs by ABA and nitrate involves the same
mechanism (Zhang et al., 2007a). Interestingly, transferring
Arabidopsis and soybean from conditions of high nitrate to
low nitrate increases root auxin (IAA) levels, suggesting that
nitrate affects auxin synthesis or transport (Caba et al., 2000;
Walch-Liu et al., 2000).
Carbon : nitrogen (C : N) ratios affect RSA, further highlighting the complexity of the root response to nitrate. A high
sucrose : nitrate ratio suppresses LRs, and a mutation in the
high-affinity nitrate transporter AtNRT2.1 abolishes this
inhibition (Malamy & Ryan, 2001; Little et al., 2005). Like
AtNRT1.1, AtNRT2.1 may be a direct nitrate sensor (Little
et al., 2005). In addition, AtNRT2.1 may have different
functions depending on the degree and context of nitrate
deficiency (Remans et al., 2006).
New Phytologist (2008) 179: 595–614
Organic nitrogen Plants can use both organic nitrogen and
inorganic nitrogen as a nutrient source. Arabidopsis roots show
a specific set of responses to the amino acid l-glutamate,
which inhibits PR growth and causes concomitant increases in
LR density to varying degrees in different ecotypes (Walch-Liu
et al., 2006). This response is similar to roots grown in low
phosphate, forming a short and highly branched RSA (section
IV.2; Williamson et al., 2001; Walch-Liu et al., 2006).
Root responses to l-glutamate are accompanied by dramatic
cytological changes, including microtubule depolymerization
(Sivaguru et al., 2003).
The PR tip is the sensor for l-glutamate (as with phosphate;
section IV.2), which inhibits cell division in the meristem
(Walch-Liu et al., 2006). Interestingly, the auxin transport
mutant aux1 is somewhat insensitive to l-glutamate, whereas
the axr1 mutant (a modifier of auxin signalling and possibly
also other hormone pathways) is hypersensitive to l-glutamate,
and various other auxin-signalling mutants exhibit wildtype sensitivity to l-glutamate (Walch-Liu et al., 2006). lglutamate probably acts as a signalling molecule rather than
as a nutritional cue, because closely related amino acids do not
elicit changes in root architecture (Walch-Liu et al., 2006).
The molecular mechanism of l-glutamate perception at the
Arabidopsis root tip remains to be discovered. Interestingly,
rice with a mutant putative glutamate receptor (OsGLR3.1)
has short PRs and LRs, reduced cell division, and premature
differentiation and cell death in the root meristem (Li et al.,
2006a).
Carnitine, an organic nitrogenous cation, induces LR
formation. However, disruption of an Arabidopsis plasma
membrane-localized carnitine transporter, AtOCT1, led
to increased root branching (Lelandais-Briere et al., 2007).
AtOCT1 promoter activity is present in the root vasculature,
including at sites of LR initiation. This suggests a possible
modulatory role for carnitine movement or homeostasis in the
control of RSA. It seems that AtOCT1 negatively regulates LR
development, and the local concentration of carnitine in the
root may affect the C : N ratio and hence LR development
(Lelandais-Briere et al., 2007).
2. Phosphorous: modulating total RSA but sensed at
the root tip
Phosphorous is an essential nutrient, primarily taken up via
the roots as inorganic phosphate (Pi). Phosphate is one of the
most inaccessible macronutrients in the soil, as it forms
insoluble compounds with metals in acidic and alkaline soils
(Raghothama, 1999). Root system architecture modifications
in response to phosphate are critical for the fitness of the plant
and differ from those seen with nitrate, perhaps reflecting a
Pi-foraging strategy, in contrast to nitrate responses that
improve nitrogen uptake (Fitter et al., 2002).
The main adaptive trait for accessing phosphate is the ability
to explore different layers near the soil surface through
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changes in the RSA (Lopez-Bucio et al., 2000). Various
adaptations have evolved in different plants. In Arabidopsis, Pi
deficiency favours a redistribution of growth from the PR to
LRs. The PR stops growing and the density and elongation of
LRs increases, forming a shallow, highly branched root system
(Williamson et al., 2001; Lopez-Bucio et al., 2002). This
prevents further growth into less nutrient-rich deeper soils,
increasing the exploration of the more nutrient-rich upper
strata. In the bean Phaseolus vulgaris, a different strategy has
evolved for achieving a similar explorative result: the angle of
root growth is shifted to predominantly outwards instead of
downwards when Pi levels are low (Bonser et al., 1996). The
nitrogen-fixing white lupin forms proteoid (cluster) roots that
secrete organic acids and phosphatases into the surrounding
soil to solubilize phosphate and aid its uptake (Schulze et al.,
2006).
Unlike nitrate responses, the initial effect of low-Pi sensing
is the arrest of PR growth, with changes in LRs occurring later.
Loss of PR growth occurs via reduced cell elongation and a
progressive loss of meristematic activity (Williamson et al.,
2001; Sanchez-Calderon et al., 2005). The phosphate deficiency
response-2 (pdr2) mutant displays hypersensitive inhibition of
cell division in developing root meristems under Pi-limiting
conditions, suggesting that PDR2 is required for meristem
function where external Pi is low. It therefore represents a
Pi-sensitive checkpoint that monitors Pi status and allows the
root system to adjust accordingly (Ticconi et al., 2004). In
addition to soil Pi status, systemic Pi levels may also be
important for the induction of Pi-deficient RSA responses
(Williamson et al., 2001). Active photosynthesis, or the
presence of sugar, is also essential for RSA responses to limiting
phosphate (Karthikeyan et al., 2007).
Physical contact of the Arabidopsis PR tip with low-Pi
medium is necessary and sufficient to arrest primary growth
and reprogram root architecture (Svistoonoff et al., 2007).
Multicopper oxidase mutants LOW PHOSPHATE ROOT-1
and -2 (LPR-1/-2) form long PRs in low Pi and provide
evidence that the root cap has an important role in nutrient
sensing. Interestingly, LPR1 was previously identified as a
quantitative trait locus (QTL) important for phosphate
responses (Reymond et al., 2006). Despite highlighting a
novel role for multicopper oxidases in plant development, it is
unknown whether LPRs are directly involved in the stimulation
of LR growth in low Pi.
A variety of hormones may modify the Pi response.
Responses to low Pi correlate with increased auxin sensitivity
and changes in auxin transport (Lopez-Bucio et al., 2002; Jain
et al., 2007). Low phosphate resistant (lpr) mutants of BIG, a
protein required for wild type levels of auxin transport, have
reduced LRs in low Pi (Gil et al., 2001; Lopez-Bucio et al.,
2005). However, neither BIG nor auxin transport is required
for other RSA modifications seen in low Pi (Lopez-Bucio
et al., 2005). Interestingly, many root responses to phosphate
starvation are repressed by cytokinin signalling (Franco-Zorrilla
Review
et al., 2005). In addition, Pi starvation affects gibberellin
signalling in roots, whereas gibberellin can attenuate the
low-Pi response (Jiang et al., 2007).
A variety of protein regulators of the phosphate-deficiency
response have been uncovered, which affect transcription,
translation and post-translational modifications. PHOSPHATE STARVATION RESPONSE-1 (PHR1) is an Arabidopsis MYB-like transcription factor that regulates a number
of Pi-deficient responsive genes and is conserved in various
plant species (Rubio et al., 2001). Miura et al. (2005) reported
that PHR1 is a target of the small ubiquitin modifier (SUMO)
E3-ligase AtSIZ1 in vitro. Interestingly, Atsiz1 mutants exhibit
an exaggerated response to low Pi levels compared with wild
type, most notably an extensive increase in LR development
and a stronger PR inhibition (Miura et al., 2005). Although
no direct link between PHR1 and RSA modification has been
shown, two genes that belong to the PHR1 regulon (AtIPS1
and AtRNS1) are positively regulated by AtSIZ1 during the
initial stages of Pi limitation (Miura et al., 2005). However, it
is unknown whether the root phenotype of Atsiz1 mutants is
a result of PHR1 modification and subsequent downstream
gene expression, or whether the effect is pleiotropic, as AtSIZ1
also has roles in other developmental pathways (Jin et al.,
2008).
Other transcription factors identified, but not fully characterized as Pi-response components, include the basic leucine
zipper (bZIP) transcription factor, PHI-2, in tobacco, and
more recently OsPTF1, a bHLH transcription factor providing
tolerance to low-Pi conditions in rice (Sano & Nagata, 2002;
Yi et al., 2005). The WRKY75 transcription factor is strongly
induced during Pi deprivation (Devaiah et al., 2007). Several
genes are downregulated in plants with reduced levels of
WRKY75, including high-affinity Pi transporters, which
consequently leads to reduced phosphate uptake during Pi
starvation (Devaiah et al., 2007). WRKY75 may be a specific
modulator of LR development (rather than affecting the PR)
and may also act independently of the Pi status of the plant to
modify LR development (Devaiah et al., 2007).
3. Root responses to sulphur
Sulphur, in the form of sulphate, is required for the synthesis
of methionine and cysteine and is critical for cellular
metabolism, growth and development, and stress responses.
Sulphate deficiency is detrimental to a plant’s survival and
leads to the development of a prolific root system, usually at
the expense of shoot growth (Kutz et al., 2002). Sulphatedeficient roots elongate faster than those with sufficient
sulphate, with LRs developing earlier, closer to the root tip
and at a greater density (Kutz et al., 2002). This leads to an
increase in total root surface area and a greater exploration of
the soil.
Sulphur deprivation leads to transcriptional activation of
NITRILASE3 (NIT3), which converts indole-3-acetonitrile
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(IAN) to auxin (Kutz et al., 2002). Low sulphate-induced
LRPs exhibit high NIT3 promoter activity, thus generating
additional auxin close to the pericycle, allowing increased LR
initiation (Kutz et al., 2002). Sulphur deficiency also upregulates
the sulphate transporter genes SULTR1;1 and SULTR1;2 in
the epidermis and cortex of roots (Yoshimoto et al., 2002); both
transporters are reversibly downregulated in sulphate-replete
conditions (Maruyama-Nakashita et al., 2004). A sulphur
response regulatory element (SURE) is conserved in the
upstream region of a variety of sulphate-deficient response
genes, suggesting that RSA alterations in response to sulphur
levels are coordinately controlled in Arabidopsis roots
(Maruyama-Nakashita et al., 2004).
Interestingly, SURE regions contain ARF consensus
sequences, suggesting a role for auxin in the early sulphatestarvation response (Maruyama-Nakashita et al., 2004). A
number of AUX/IAA genes have been implicated in the sulphate response, and transcriptomic analysis suggests that both
auxin influx and IAA28 activity may modulate the response to
low sulphate, perhaps by acting in a negative regulatory manner
(Nikiforova et al., 2003, 2005). More recently, (Dan et al.,
2007) suggested that auxin is involved in a subset of sulphurdeficiency responses, with other hormones (such as cytokinin
and ABA) also playing a role. SULTR1 mRNA accumulation
can be reduced by exogenous cytokinin, further suggesting
points of regulation (Maruyama-Nakashita et al., 2004).
4. Potassium and lateral root development
Lateral roots of potassium-starved plants arrest their elongation
(Armengaud et al., 2004). Analysis of root transcriptomes
from potassium-starved seedlings which were then resupplied
with potassium revealed that certain genes were downregulated
by potassium resupply, including stress-induced genes,
transporters, calcium signalling components, sulphur
metabolism components, and cell wall-remodelling enzymes.
Conversely, upregulated genes were either transporters
(including three root-specific nitrate transporters) or cell wallremodelling enzymes. The transcriptomic profile of potassiumstarved plants overlaps with sulphur starvation, but not with
nitrate starvation or phosphate starvation, and also involves
changes in jasmonate/defence signalling (Armengaud et al.,
2004). Interestingly, the MYB77 transcription factor provides
a direct link between potassium starvation responses and
auxin signalling (Shin et al., 2007).
5. Water and salt stresses
Water stress Water availability has a profound effect on a
plant’s root system. Plant roots will grow towards wetter soil
and away from high osmolarity (Takahashi et al., 2003). As
water availability decreases (or osmotic stress increases), LR
emergence is repressed, although LR initiation is largely
unaffected (van der Weele et al., 2000; Deak & Malamy,
New Phytologist (2008) 179: 595–614
2005; Xiong et al., 2006). This is likely to be an adaptive
response encouraging increased water uptake from deeper soil
layers. The molecular mechanisms underpinning the response
are largely unknown, although ABA has an important role.
The ABA-deficient mutants aba2-1 and aba3-2 have increased
root system size compared with wild type under high osmotica
(Deak & Malamy, 2005). Plants mutant for the LATERAL
ROOT DEVELOPMENT 2 (LRD2) and Arabidopsis
CYTPOLASMIC INVERTASE (AtCYT-INV1) genes also
have a similar phenotype (Deak & Malamy, 2005; Qi et al.,
2007). Alongside ABA, LRD2 may be required to determine
the percentage of LRPs that become LRs under normal and
stress conditions (Deak & Malamy, 2005).
Abscisic acid and drought stress have similar and probably
synergistic effects on LR development. Several drought inhibition
of lateral root growth (dig) mutants have enhanced responses to
ABA and are also drought tolerant, whilst others have a
reduced LR-inhibition response to ABA and are drought
sensitive (Xiong et al., 2006). DIG3 is particularly important
for LR inhibition in response to ABA: dig3 mutants have
normal LR growth under stress and are susceptible to drought.
Interestingly, dig3 plants were smaller than wild-type plants
under well-watered conditions, suggesting that the ABA and
drought response involves factors required more generally for
growth (Xiong et al., 2006). Drought tolerance in crop species
is controlled by multiple QTLs (Nguyen et al., 2004): it will
be interesting to discover whether dig loci define droughttolerant QTLs that are important for responding to water-stress
in roots and globally.
Salt stress Salt stress, which is related to drought stress, also
reprograms RSA. Salt stress in Arabidopsis can induce root
swelling with shorter total root lengths, a seriously reduced
meristematic zone and a strong reduction in the number of
LRPs, accompanied by the downregulation of several cell cycle
genes (Burssens et al., 2000). However, salt stress may also
trigger an increase in LR number. In chickpea (Cicer arietinum),
the CAP2 (C. arietinum AP2) transcription factor is induced
upon dehydration and binds to dehydration-response elements
in many stress-inducible genes (Boominathan et al., 2004;
Shukla et al., 2006). Transgenic tobacco expressing CAP2 is
tolerant to salinity and osmotic stress, possibly because of a
large increase in LR number (Shukla et al., 2006). Many
auxin-response genes associated with LR development are
upregulated in these plants, indicating links between salt stress
responses and intrinsic auxin-associated development (Shukla
et al., 2006).
He et al. (2005) reported increased LR numbers and a
reduction of PR length in response to high levels of NaCl in
Arabidopsis. The NAC2 transcription factor is upregulated by
NaCl and its overexpression causes increased LR formation
specifically without a change in root length (He et al., 2005).
NAC2 is upregulated by ethylene, auxin and ABA, and its
induction by salt is compromised in auxin and ethylene
www.newphytologist.org © The Authors (2008). Journal compilation © New Phytologist (2008)
Tansley review
signalling mutants (He et al., 2005). These data highlight the
importance of phytohormone signalling in RSA responses to
salinity.
6. Effects of light on root architecture
Responding to light is key to plant survival. In addition to
having profound effects on the seed and shoot, light can affect
LR morphology. This can be direct (e.g. red light enhances LR
formation via the COL3 gene) (Datta et al., 2006) or indirect,
via effects in the shoot (Bhalerao et al., 2002).
The bZIP transcription factor LONG HYPOCOTYL 5
(HY5) is a key player in light-induced development in Arabidopsis (Koornneef et al., 1980). Initially noted for defective
light-induced hypocotyl elongation, hy5 mutants also have an
elevated number of LRs, which grow faster than wild-type
roots and are less responsive to gravity (Oyama et al., 1997).
The hy5 root phenotype occurs as a result of the underexpression of two negative regulators of the auxin signalling pathway:
AUXIN RESISTANT 2 (AXR2)/IAA7 and SOLITARY
ROOT (SLR)/IAA14 (see section III.1 ‘Lateral root initiation
requires auxin and regulated protein degradation’; Cluis et al.,
2004). This interaction between HY5 and auxin signalling
highlights the importance of both light-signalling networks
and hormone-signalling networks in the control of RSA. HY5
also interacts with SALT TOLERANCE HOMOLOGUE 2
(STH2) (Datta et al., 2007). The sth2 mutant phenocopies
the exaggerated root phenotype of the hy5 mutant, and
the authors suggest that light-dependent inhibition of LRs
by STH2 requires its binding to HY5, where it provides
transactivating potential to the transcription factor (Datta
et al., 2007).
HY5 HOMOLOGUE (HYH) is a functional equivalent of
HY5 with a similar expression pattern and responsiveness to
light (Sibout et al., 2006). hyh mutants show wild-type RSA.
However, hy5 hyh double mutants exhibit a suppression of
the hy5 phenotype, displaying less prolific root growth than
wild type (Sibout et al., 2006). It has been proposed that
these double mutants represent the morphological response
to a quantitative gradient in auxin signalling. This example
suggests that the inactivation of genes, both of which affect
the balance of a physiological process in the same manner, can
result in very different morphological changes (Sibout et al.,
2006).
Molas et al (2006) examined total gene expression in
dark-grown roots that were treated with red light for just 1 h.
Interestingly, genes affecting cell wall metabolism and remodelling were consistently downregulated. Genes involved in
hormone signalling (auxin, GA, ethylene) were also affected,
as were proteins involved in intercellular transport, various
transcription factors and several F-box proteins (Molas et al.,
2006). Thus, light-induced changes in RSA are likely to
happen rapidly and involve both signalling and remodelling
processes.
Review
7. Modulation of root architecture by biotic factors
Within the soil, plants must compete and interact with a
plethora of organisms, including microorganisms and other
plant root systems. Roots secrete chemicals into the soil that
affect other plant RSAs and also influence communication
with microorganisms (Bais et al., 2004). In turn, viruses,
bacteria and fungi can modify RSA. Many of these interacting
species are pathogens and result in plant defence responses,
while some can form symbiotic interactions leading to the
formation of root nodules or mycorrhizas/proteoid roots
(Autran et al., 2006; Oldroyd & Downie, 2006). In addition,
soil microorganisms can produce auxin and cytokinin that
dramatically affect RSA (see Section III) (Costacurta &
Vanderleyden, 1995). Some recent advances in the molecular
understanding of how pathogens modify RSA are presented
in the following sections.
Viral proteins Cucumber mosaic virus (CMV) infects a range
of dicots, inducing developmental and growth abnormalities.
The severity of disease symptoms is dependent on the
CMV-2b protein (Lewsey et al., 2007). CMV-2b bypasses
host defences both by inhibiting plant RNA silencing
mechanisms (thus promoting the undetected spread of viral
RNA) and by antagonizing salicylic acid signalling, which
normally inhibits viral replication and cell-to-cell spreading.
Arabidopsis infected with CMV or overexpressing CMV-2b
show perturbed RSA, specifically, shorter PRs, increased LR
density and increased LR length, leading to increased root
surface area (Lewsey et al., 2007). Interestingly, CMV-2b
stabilizes a number of endogenous Arabidopsis mRNAs that
are targets of degradation by microRNAs (miRNAs), including
the auxin signalling genes ARF17 and NAC1. Curiously,
stabilized ARF17 is proposed to inhibit LR formation (Mallory
et al., 2005), whereas NAC1 promotes LR development (Xie
et al., 2000), suggesting that targeting of NAC1 may be
particularly relevant during CMV infection. Cucumber
mosaic virus ultimately inhibits root growth, but it is possible
that transient increases in LR formation upon CMV infection
are advantageous during initial virus infection and spread,
because the presence of a higher number of emerging LRs and
an increase in root surface area could provide a greater number
of sites for virus entry.
Pathogenic bacteria and fungi Pathogenic bacteria and fungi
can directly influence LR development. Ralstonia solanacearum
inoculation leads to reduced formation and elongation of LRs
in petunia (Zolobowska & Van Gijsegem, 2006). Novel root
lateral structures develop, derived from the pericycle founder
cells that normally form LRs. These seem to act as colonization
sites, and this process probably requires secreted bacterial
proteins (Zolobowska & Van Gijsegem, 2006).
The bacterium Pseudomonas syringae stimulates LR
development and other auxin-related changes in Arabidopsis,
© The Authors (2008). Journal compilation © New Phytologist (2008) www.newphytologist.org
New Phytologist (2008) 179: 595–614
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whereas exogenous auxin promotes disease progression (Chen
et al., 2007). The authors suggest that auxin could promote
cell wall loosening. A Pseudomonas peptide was reported to
downregulate auxin signalling, enabling disease resistance;
however, the effect on LR development was not investigated
(Navarro et al., 2006). The rice transcription factor OsWRKY31
is induced by rice blast fungus and also by auxin. Overexpression
of OsWRKY31 confers resistance to rice blast fungus infection
and also inhibits LR formation (Zhang et al., 2007b) In addition,
OsWRKY31 upregulates auxin responsive genes, again linking
auxin signalling and/or transport with the defence response
(Zhang et al., 2007b).
Interestingly, the nitrate-inhibitory effect on LR development
is over-ridden when plants are inoculated with Phyllobacterium,
a growth-promoting rhizobacterium (Mantelin et al., 2006).
This effect was accompanied by altered expression of various
transport genes, including AtNRT1.1.
Continuing research in this new area will define the extent
to which plant–pathogen interactions affect RSA.
V. Transcriptomic studies to identify potential
new regulators of lateral root development
With the advent of high-throughput ‘omic’ experimental
techniques, it is possible to augment molecular genetic
studies of LR developmental mechanisms. This has included
the generation of data sets of genes that are upregulated
or downregulated specifically in different root cell types
or in response to specific signals (see various sections
above).
In terms of probing cell type-specific gene expression in
roots, studies from the Benfey laboratory have been influential.
An initial study identified several hundred genes enriched in
vascular tissues, including pericycle (Birnbaum et al., 2003).
More recently, a high-resolution gene-expression map of
all root cell types, including pericycle, xylem pole pericycle,
phloem pole pericycle and LRPs, has been created (Brady
et al., 2007). Analysis of this vast data set will provide new
insights into the gene regulation occurring during LR
development. For example, pericycle (in particular xylem pole
pericycle) is enriched in mRNAs encoding cell wall-modifying
enzymes, whereas genes encoding kinases and enzymes required
for cell wall loosening are enriched in LRPs. In addition, auxin
biosynthetic genes are enriched in pericycle and LRPs, and
ABA signalling components are enriched in pericycle (Brady
et al., 2007).
A meta-analysis identifying indirect targets of the SHORTROOT (SHR) transcription factor uncovered a number of
SHR-regulated genes that were enriched in or exclusive to
pericycle (Levesque et al., 2006). It is thus tempting to speculate that SHR signalling pathways may regulate LR formation
as well as PR development (Scheres et al., 2002), especially as
similar signalling may regulate AR formation in pine trees and
sweet chestnut trees (Sanchez et al., 2007).
New Phytologist (2008) 179: 595–614
Specific cell types have been isolated from maize roots by
laser capture microdissection (LCM) for transcriptomic
and proteomic analysis (Woll et al., 2005; Dembinsky et al.,
2007). Comparison of the pericycle transcriptome of
wild-type maize with that of a mutant that cannot initiate
LRs revealed that the majority of differentially expressed
genes are involved in transcription or metabolism, or have
unknown function. However, several genes involved in signal
transduction (especially protein kinases), cell cycle regulation,
cellular transport and defence were also identified (Woll et al.,
2005).
To identify genes involved in pericycle cell fate specification,
rather than LR formation per se, pericycle cells were dissected
from along the length of the root before the time that cell
divisions occur (Dembinsky et al., 2007). The pericycle
transcriptome and proteome was analysed, and further
pericycle-enriched genes were isolated from cDNA libraries
and expressed sequence tags (ESTs) (Dembinsky et al., 2007).
Around 40 ‘pericycle-specific’ genes were identified, of which
the largest two subsets were transcriptional regulators and
unknown genes. Compared with vascular cells, pericycle
appears enriched in genes involved in protein synthesis, but
low in genes regulating cell fate. Twenty abundant soluble
pericycle proteins were identified, of which 80% have a metabolic or energy function. There is only a small overlap between
the LR initiation data set (Woll et al., 2005) and the pericycle
data sets (Dembinsky et al., 2007), suggesting that specifying
pericycle cell identity is a distinct process from forming a
new LR.
VI. Conclusions and future challenges
A vast number of signals, both from within and outside the
plant, impinge on the root to regulate its final architecture and
branching pattern. Many challenges still exist for future ‘root
biologists’. We must understand at the molecular level how
these different signals work together to direct pericycle cell
behaviour and later LR developmental processes. We must
discover potentially novel signals that regulate LR developmental proteins which currently reside outside known
signalling networks. There are questions we can ask about the
evolution of RSA regulation across the plant kingdom. There
are huge transcriptomic data sets that will provide us with new
clues about the changes in gene expression necessary for LR
development to occur. Moving our knowledge gained in
Arabidopsis and genetically tractable crop plants into other
agronomically relevant species will provide an understanding
of how to engineer crop plants that can exist in a range of
potentially problematic environments.
Acknowledgements
The authors thank Laurent Laplaze and Jeremy Roberts for
useful comments.
www.newphytologist.org © The Authors (2008). Journal compilation © New Phytologist (2008)
Tansley review
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