Download Auxin and self-organization at the shoot apical

Journal of Experimental Botany, Vol. 64, No. 9, pp. 2579–2592, 2013
doi:10.1093/jxb/ert101 Advance Access publication 12 April, 2013
REVIEW PAPER
Auxin and self-organization at the shoot apical meristem
Massimiliano Sassi and Teva Vernoux*
Laboratoire de Reproduction et Développement des Plantes, CNRS, INRA, ENS Lyon, UCBL, Université de Lyon, 46 Allée d’Italie,
69364 Lyon Cedex 07, France.
* To whom correspondence should be addressed: Email: [email protected]
Received 8 February 2013; Revised 7 March 2013; Accepted 13 March 2013
Plants continuously generate new tissues and organs throughout their life cycle, due to the activity of populations
of specialized tissues containing stem cells called meristems. The shoot apical meristem (SAM) generates all the
aboveground organs of the plant, including leaves and flowers, and plays a key role in plant survival and reproduction.
Organ production at the SAM occurs following precise spatio-temporal patterns known as phyllotaxis. Because of the
regularity of these patterns, phyllotaxis has been the subject of investigations from biologists, physicists, and mathematicians for several centuries. Both experimental and theoretical works have led to the idea that phyllotaxis results
from a self-organizing process in the meristem via long-distance interactions between organs. In recent years, the
phytohormone auxin has emerged not only as the central regulator of organogenesis at the SAM, but also as a major
determinant of the self-organizing properties of phyllotaxis. Here, we discuss both the experimental and theoretical
evidence for the implication of auxin in the control of organogenesis and self-organization of the SAM. We highlight
how several layers of control acting at different scales contribute together to the function of the auxin signal in SAM
dynamics. We also indicate a role for mechanical forces in the development of the SAM, supported by recent interdisciplinary studies.
Key words: auxin, auxin signalling, mechanical forces, phyllotaxis, polar auxin transport, shoot apical meristem.
Introduction
One of the most striking differences between plants and
animals lies in the regulation of their organogenetic capacity over time. Unlike most animals, in which organogenesis
occurs during embryonic development, plants can form new
organs throughout their life cycle. Post-embryonic development occurs thanks to specialized tissues containing stem-cell
niches known as meristems. Plants possess different types of
meristems that control both primary and secondary growth.
Among the meristems controlling primary growth, the shoot
apical meristem (SAM) is located at the apex of the shoot
axis and is responsible for the generation of all the aerial
organs, in particular leaves and flowers. The SAM is thus not
only important for building up the plant architecture but also
for plant photosynthetic uptake and reproductive fitness. The
SAM is established during embryogenesis and remains active
throughout the plant life, which can last for up to thousands
of years, as in the case of the giant sequoia (Sequoiadendron
giganteum). A high level of coordination between the reiterative process of organogenesis and the self-renewal activity of
the stem cells is thus expected to maintain the SAM function
over long periods of time.
Coordination of cellular behaviours and cell fate is also
evident when considering the spatio-temporal patterns of
organogenesis at the SAM or phyllotaxis. Organs are indeed
initiated sequentially and at relatively precise spatial positions
in the SAM. A majority of plants, including the model species
Arabidopsis thaliana, have a spiral phyllotaxis where organs
are initiated one by one and at a 137° relative angle from the
previous organ. The regularity of these patterns has intrigued
not only biologists but also physicists and mathematicians
for several centuries. Both experimental and theoretical
works have led to the idea that phyllotaxis could arise from
Abbreviations: CZ, central zone; IAA, indole-3-acetic acid; OC, organizing centre; PZ, peripheral zone; SAM, shoot apical meristem; TF, transcription factor.
© The Author [2013]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For permissions, please email: [email protected]
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Abstract
2580 | Sassi and Vernoux
long-distance interactions between organs in a self-organization process. In this context, the phytohormone auxin is
a central regulator of the SAM organogenetic capacities but
also as an orchestrator of the spatio-temporal dynamics of
the SAM patterning. After presenting some of the key molecular players involved in regulating SAM function, we will discuss here both the experimental and theoretical evidence for
the implication of auxin in the control of organogenesis and
self-organization of the SAM. We will highlight how several
layers of control contribute together to the function of the
auxin signal in SAM dynamics. We will also indicate a role
for mechanical forces in SAM morphogenesis supported by
recent interdisciplinary studies.
Cellular organization and genetic
regulation of the SAM
Phyllotactic patterning as a self-organizing
developmental process
Phyllotaxis represents one of the most striking examples of
geometrical regularities that can result from a developmental patterning process. In higher plants, different phyllotactic patterns can be observed, being defined by the number
of organs formed at any given time and their relative spatial
distribution. The geometry corresponding to each phyllotaxis
is defined by precise mathematical rules. In the case of spiral
phyllotaxis, the most common phyllotactic pattern, the divergence angle (i.e. the angle between two consecutive organs)
approximates to 137.5°. This value corresponds to the golden
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
The structure of the SAM is relatively well conserved among
angiosperms, on which this review will focus. In general, the
SAM can be divided into an external layer called the tunica
and an inner region called the corpus. These two regions are
very well defined at a cellular level: the cells of the corpus
divide without a preferential cell division plane, whereas the
cells of the tunica mostly divide anticlinally (i.e. perpendicular to the SAM surface), generating a typical layered structure, as daughter cells usually remain in the same layer as
their parents. The tunica can be composed of several layers of
cells (two in the case of Arabidopsis, named L1 and L2), and
the layered organization is maintained only in the epidermal
layer once the cells are recruited in organ primordia.
Superimposed on the tunica–corpus organization, different domains of the meristem can also be defined based on
the rates of cell growth and division. In the central zone (CZ)
of the SAM, cells divide infrequently and grow at a low rate.
By contrast, in the peripheral zone (PZ) surrounding the CZ,
cells grow and divide at much higher rates. The PZ corresponds to the organogenetic domain of the SAM. As soon
as the organ primordia starts to bulge out, as a result of the
increase in cell proliferation and growth of a small number of
founder cells, another group of cells at the proximal end of
the organ will slow down their growth rates. These cells will
form the boundary between the organ and the SAM, providing a physical separation between these two structures (Traas
and Doonan, 2001).
Specific gene expression patterns also define functional
regions in the SAM that partly overlap with the zonation of
the SAM based on the structural organization and cellular
behaviours. A large number of these genes have been identified from mutants affected in SAM function. We will present
here only briefly the most important regulators, and we refer
the reader to recent reviews for a more exhaustive view (Aida
and Tasaka, 2006; Sablowski, 2007; Rast and Simon, 2008;
Dodsworth, 2009; Perales and Reddy, 2012).
The CZ is characterized by expression of the CLAVATA3
(CLV3) gene. CLV3 encodes a signalling peptide implicated
in a well-defined self-regulatory feedback loop, composed
of the CLV3 secreted ligand, a receptor signalling cascade
implicating notably CLV1 and the WUSCHEL (WUS)
homeodomain transcription factor (TF). The expression of
CLV3 is specific to the stem cells found in the CZ. WUS is
expressed in a small domain in the corpus of the SAM, below
the stem cells. The WUS domain defines the organizing centre
(OC), and WUS is involved in promoting the expression of
CLV3. The CLV3 peptide directly binds the receptor kinase
CLV1 and activates a signalling cascade that restricts the size
of the OC underneath. This non-cell-autonomous regulatory network implicating antagonistic factors determines and
maintains the integrity of the SAM stem-cell niche. Indeed,
wus mutants are unable to maintain an active SAM, whereas
the size of the SAM is dramatically increased in clv3 mutants
(Laux et al., 1996; Mayer et al., 1998; Schoof et al., 2000;
Brand et al., 2000; Ogawa et al., 2008; Yadav et al., 2011).
To give rise to a new organ in the PZ, a group of cells loses
its meristematic identity and acquire a new organ identity. The
homeodomain TF SHOOT-MERISTEMLESS (STM), which
plays a central role in meristem maintenance, is expressed
throughout the SAM but is excluded from organ primordia
(Endrizzi et al., 1996), and the loss of meristematic identity is
in part achieved by repression of STM in the organ founder
cells (Byrne et al., 2000; Heisler et al., 2005). In parallel, a
number of transcriptional regulators are induced in the organ
primordium and will determine organ fate and tissues identities. The expression of the TFs of the YABBY, KANADI, and
HD-Zip III families regulates the patterning of the organ by
establishing its dorso/ventral polarity (Bowman et al., 2002;
Byrne, 2006), whereas the expression of AINTEGUMENTA
(ANT) promotes organ growth at least partially by regulating
cell proliferation (Mizukami and Fischer, 2000). The expression of the CUP-SHAPED COTYLEDON (CUC) TFs specifies the boundaries as soon as the primordium starts to bulge
out and allows for the separation of the organ from the meristem (Furutani et al., 2004; Heisler et al., 2005). Moreover,
the absolute expression level of LEAFY (LFY) determines
whether the organ acquires a leaf (low LFY level) or a flower
(high LFY level) identity (reviewed by Moyroud et al., 2010).
This transcriptional programme is reiteratively started in the
PZ at each time a new organ is formed. This implies a tight
control of transcription in space and time in order to coordinate the continuous and regular production of the new organs
in the SAM and to establish phyllotactic patterns.
Auxin at the shoot apical meristem | 2581
Polar auxin transport and phyllotaxis at
the SAM
A possible mechanism explaining the inhibitory fields would
be the diffusion or transport of a molecule that inhibits
organogenesis (see, for example, Mitchison, 1977). However,
so far, no such inhibitory molecule has been identified. By
contrast, a number of experimental and theoretical evidence
have shown that auxin is both necessary and sufficient for
organ formation at the SAM and that polar auxin transport
depletes auxin in the region surrounding an organ, thus creating inhibitory fields. After presenting the mechanisms that
control polar auxin transport, we will discuss the data supporting this vision.
The regulation of polar auxin transport
The active polar transport of auxin in plant tissues has been
the subject of intense investigation for several decades. The
classical chemiosmotic hypothesis (Rubery and Sheldrake,
1974; Raven, 1975) proposed that the chemical properties
of the natural occurring auxin indole-3-acetic acid (IAA)
are at the basis of auxin transport in plants. As a weak acid,
IAA charge is pH dependent: in acidic (pH 5.5) environments such as in the apoplast, IAA is protonated (IAAH)
and can easily move and enter the cell via diffusion. At neutral intracellular pH, IAAH loses its proton and becomes
ionized (IAA–). Due to its charge, IAA– is trapped inside the
cells. The chemiosmotic hypothesis then predicts that auxin
efflux from cells requires an active carrier-dependent transport and that the polarity of the efflux carrier determines
the polarity of the auxin flux (Leyser, 2005; Vieten et al.,
2007; Friml, 2010).
Actually, several transmembrane carriers accounting for
both cellular influx and efflux of auxin have been identified in
Arabidopsis over the years. The cellular influx of auxin is mediated by the AUXIN RESISTANT1/LIKE AUX1 (AUX1/
LAX) family of amino acid permease-like proteins, whereas
the efflux from the cells is controlled mainly by members of
the PIN-FORMED (PIN) family of transmembrane proteins
(Leyser, 2005; Vieten et al., 2007; Friml, 2010). Members of the
subfamily B of ATP-binding cassette/P-glycoprotein (ABCB/
PGP) transporters have also been demonstrated to mediate
cellular auxin efflux and to interact with PIN transporters at
different levels (Blakeslee et al., 2007; Mravec et al., 2008). PIN
proteins are transmembrane carriers that perform a rate-limiting step in auxin efflux (Petrásek et al., 2006). As implied by the
chemiosmotic hypothesis, PIN carriers are localized polarly on
the plasma membrane of the cells and their polar localization
controls the direction of auxin transport (Wisniewska et al.,
2006). Thus, by observing the patterns of PIN polar localization in a plant tissue, it is possible to predict the direction
of the auxin movement. The regulation of polar PIN localization is still not fully understood, although possible mechanisms have been identified. PIN proteins undergo constitutive
cycles of clathrin-mediated endocytocis and recycling back
to the plasma membrane (Geldner et al., 2001; Dhonukshe
et al., 2007). Endocytic recycling has been shown to allow for
the polar localization of PINs after an initial apolar delivery
of newly synthesized PIN proteins (Dhonukshe et al., 2008).
Rapid changes in PIN polarity have been shown to be determined by a transcytosis-like mechanism in which, after an initial step of endocytosis, PIN proteins are recycled to a different
polar targeting domain on the plasma membrane (Kleine-Vehn
et al., 2008a, 2010). Moreover, reversible phosphorylation of
PINs by the PINOID (PID) serine/threonine protein kinase
and the regulatory subunits A of the protein phosphatase 2A
(PP2AAs) has also been demonstrated to play a central role in
the decision of targeting PINs to the apical versus the basal
membrane of cells (Friml et al., 2004; Michniewicz et al.,
2007). In particular, it was recently proposed that phosphorylation via PID and related AGC3 kinases WAG1 and WAG2
recruits PINs into an apical recycling pathway as opposed to
the default basal recycling that occurs in the absence of phosphorylation (Dhonukshe et al., 2010).
While PIN proteins regulate auxin distribution within a tissue, auxin can in turn regulate PIN activity in several different
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
angle defined from the Fibonacci series. Each term of the
Fibonacci series is given by the sum of the two previous ones
(mathematically defined by in=in–1 + in–2; i0=i1=1). The golden
angle is then the smaller angle resulting from the division of
the circumference of a circle according to the value of the
limit of the series obtained by dividing two consecutive terms
of the Fibonacci series, this value being known as the golden
ratio. Spiral phyllotaxis also generates a geometric distribution of organs at the apex where higher-order clockwise and
counterclockwise spirals, known as contact parastichies, can
be visualized by connecting adjacent organs. The number
of contact parastichies generated by a SAM generally corresponds to two consecutive terms of the Fibonacci series
(Reinhardt, 2005a,b; Kuhlemeier, 2007).
The seminal experiments from Snow and Snow, more
recently re-explored using modern laser-ablation approaches,
have suggested that existing organs inhibit the formation
of new primordia in their vicinity (Snow and Snow, 1931;
Reinhardt et al., 2005). These results, together with the observation that a new primordium forms in the largest available
space in the SAM (Höfmeister 1868), have fuelled the formulation of a number of theoretical models, collectively known as
inhibitory field models (reviewed by Adler et al., 1997). These
models postulate that organs generate inhibitory fields that
prevent new organ initiation occurring in their vicinity. Among
these, abstract dynamic models are sometimes considered as
a paradigm for phyllotaxis (Shipman and Newell, 2005) and
have been instrumental in exploring the putative role of inhibitory fields in the dynamics of phyllotaxis (Douady and Couder,
1996a,b,c). Indeed, computer simulations using dynamic models of inhibitory fields suggest that both the interval of time
between organ initiations (the plastochrone) and the relative
spatial position of the organs emerges from the local repulsive
interactions of organs and the growth in the SAM. In other
words, this theoretical work suggests that the formation of regular phyllotactic patterns is a self-organizing process that only
requires inhibitory fields produced by organs.
2582 | Sassi and Vernoux
PIN1-mediated polar auxin transport is essential for
organ initiation and phyllotaxis
Although the Arabidopsis genome encodes at least five PIN
efflux carriers active at the plasma membrane, only PIN1
plays a non-redundant role in auxin distribution at the SAM.
While pin1 mutants are able to form leaves during the vegetative stage (Guenot et al., 2012), they hardly produce any
lateral organs after the transition to the reproductive stage,
resulting in pin-like inflorescences (Okada et al., 1991;
Gälweiler et al., 1998; Vernoux et al., 2000; Reinhardt et al.,
2000). This indicates a limiting role both for PIN1 in auxin
transport and for auxin transport in organ initiation at the
inflorescence SAM, while other mechanisms might be at work
in the vegetative SAM. Moreover, local application of exogenous auxin at the periphery of the inflorescence SAM induces
de novo primordium formation in pin1 mutants (Reinhardt
et al., 2000, 2003). This suggests not only that auxin is necessary and sufficient to trigger organ initiation but also that
polar auxin transport leads to auxin accumulation at the site
of the incipient primordium, thus providing instructions for
organogenesis.
PIN1 has been shown to be expressed predominantly in the
L1 layer and in the provascular cells in the SAM (Vernoux
et al., 2000; Reinhardt et al., 2003; Benková et al., 2003;
Heisler et al., 2005; Barbier de Reuille et al., 2006). Patterns
of PIN1 polarity and expression on the SAM surface are
highly dynamic but are also stereotypical at the location of the
incipient primordia and early developing organ: PIN1 pumps
converge towards the forming organ, delimiting a small cluster of cells that display higher PIN1 expression levels without a clear polarity (Heisler et al., 2005; Barbier de Reuille
et al., 2006). These cells will be connected with the inner
PIN1-expressing provascular tissue at a slightly later stage
(Benková et al., 2003; Reinhardt et al., 2003; Heisler et al.,
2005). The PIN1 patterns in the L1 around organs suggest
that polar auxin transport concentrates auxin at the site of
the primordium formation, thus providing a mechanism for
auxin accumulation. This scenario is supported by the upregulation of the synthetic auxin-inducible DR5 marker during
organ initiation, indicating an activation of auxin signalling
(Benková et al., 2003; Heisler et al., 2005; Barbier de Reuille
et al., 2006; Smith et al., 2006), and by the results of computer simulations predicting auxin distribution in the SAM
based on the connectivity of the PIN1 network (Barbier de
Reuille et al., 2006). More recently, the DII-VENUS synthetic
auxin sensor, which was obtained by fusing the auxin-binding
domain of an Aux/IAA (see below) to a nuclear-targeted fastmaturating YFP (VENUS), allowed clear demonstration of
an accumulation of auxin starting from the earlier steps of
organ initiation (Vernoux et al., 2011; Brunoud et al., 2012).
If the primordium acts as a sink, attracting auxin from the
neighbouring cells through PIN1-mediated auxin transport,
this suggests that an auxin-depleted region surrounding the
primordium will be generated (Reinhardt et al., 2003; Heisler
et al., 2005; Barbier de Reuille et al., 2006). As a result, organ
formation will be inhibited in this auxin-depleted region.
Observation of the PIN1 polarity patterns thus leads to the
proposal that polar auxin transport could control not only
accumulation of auxin at the site of organ initiation but also
the creation of inhibitory fields around organs (Reinhardt
et al., 2003; Reinhardt, 2005a,b). In this scenario, polar
auxin transport alone would drive phyllotaxis through a tight
spatio-temporal control of auxin distribution in the SAM.
Analysing the DII-VENUS signal indeed confirmed this
depletion of auxin around the organ, starting from organ initiation onwards (Vernoux et al., 2011; Brunoud et al., 2012),
thus providing strong experimental support to this hypothesis. It is interesting to note that the depletion of auxin around
organs closely correlates with the expression of boundary
genes such as the CUC genes (Vernoux et al., 2000; Heisler
et al., 2005) and the specification of the organ boundary
identity might participate in inhibiting organogenesis close to
existing organs. Finally, further supporting the role of polar
auxin transport in the control of phyllotaxis, the AUX1/
LAX-dependent auxin influx has been shown to play a role in
promoting organ separation and stabilizing phyllotactic patterning against environmental fluctuations (Reinhardt et al.,
2003; Bainbridge et al., 2008). Although AUX1/LAX influx
carriers do not display clear cellular polarity, they are thought
to facilitate PIN1-mediated polar transport, notably by conveying auxin from the inner tissues to the L1 layer (Reinhardt
et al., 2003; Bainbridge et al., 2008).
Cell-based models suggest that phyllotaxis emerges
from the dynamics of PIN1 polarities
The dynamics of PIN1 pumps could thus explain the phyllotactic patterning, but there are two key questions. How is
this process coordinated throughout the SAM? And how can
PIN1 polarity precisely reorient to determine the next site of
primordium formation? Despite the substantial knowledge
on the molecular mechanisms of PIN polarization at a cellular level, an overall mechanistic view of how this process
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
ways. Auxin has been shown to inhibit PIN endocytosis, thus
enhancing its own efflux from the cell (Paciorek et al., 2005).
The AUXIN BINDING PROTEIN 1 (ABP1) protein acts
as an extranuclear auxin receptor and has been implicated
in this regulation. In particular, ABP1 has been shown to
inhibit clathrin-mediated endocytosis at the plasma membrane upon auxin binding (Robert et al., 2010). On the other
hand, nuclear auxin signalling through the TRANSPORT
INHIBITOR RESISTANT 1/AUXIN SIGNALLING
F-BOX (TIR1/AFB) receptors (see below) has been shown
to regulate the abundance of PINs by controlling both the
transcriptional activity of the genes and the turnover of
proteins (Vieten et al., 2005; Abas et al., 2006; Kleine-Vehn
et al., 2008b; Baster et al., 2012). In some tissues, auxin has
also been shown to promote shifts in PIN polarity in a TIR1/
AFB-dependent fashion (Sauer et al., 2006). Such feedback
regulations between auxin and PINs are thought to be crucial for the establishment and maintenance of the instructive
auxin gradients that regulate several developmental processes
including organogenesis at the SAM (Benjamins and Scheres,
2008; Vanneste and Friml, 2009)
Auxin at the shoot apical meristem | 2583
and can favour one mechanism depending on the intracellular auxin content. As a result, cells with low auxin concentration will promote a concentration-based transport mode,
whereas cells with high auxin concentration will activate a
flux-based mechanism. This combined model was also able
to reproduce realistically PIN polarization behaviour in different transport-dependent developmental contexts (Bayer
et al., 2009).
To date, all the theoretical models that have been developed assume the existence of hypothetical molecular components that can sense extracellular auxin or the intensity of
the auxin flux through the membrane and direct PIN polarization. These models clearly show that a cellular feedback
loop between auxin and the polarity of PIN1 is a plausible
mechanism to explain the emergence of a phyllotactic pattern
and self-organization in the SAM. However, they do not permit conclusions on the actual mechanisms involved. A recent
computational study might provide some first answers and
support canalization as a plausible scenario explaining
polarization of PINs in a multicellular context. Wabnik et al.
(2010) tested whether the modulation of PIN trafficking by
an extracellular auxin receptor could lead to PIN polarization as postulated in the canalization hypothesis. The authors
hypothesized that auxin binding to its receptor in the apoplast (ABP1 being an obvious candidate for such a receptor)
immobilizes the receptor and triggers inhibition of PIN internalization in the nearest cell, increasing the efflux locally. As
two neighbouring cells compete for auxin receptors in their
common walls, this model can lead to an asymmetric PIN
internalization in the two facing membranes. The authors also
included in the model the regulation of PIN transcription by
auxin through nuclear signalling and auxin transport through
both the influx and efflux carriers. The authors demonstrated
that their model can reproduce PIN-driven vascular formation in leaves and during vascular tissue regeneration in stem,
two biological contexts that are easily explained using the
canalization hypothesis (Wabnik et al., 2010). A key characteristic of the model used in this work is that PIN polarization
is driven by concentration sensors and not flux sensors, and
still the model appears to be able to simulate canalization. It
remains to be established whether the mechanism proposed
by Wabnik et al. (2010) formally behaves like a canalization
model and whether it also functions for phyllotaxis. This
point is notably important, as analysis of the role of mechanics in the SAM (which we will discuss later) rather supports
the concentration-based models of phyllotaxis. However, this
work suggests that building mechanistic models in parallel
with the exploration of the cellular mechanisms controlling
PIN polarity holds the key to the understanding of the selforganizing properties of the PIN1 network.
In this context, it should be stressed that the different
PIN1-based models of phyllotaxis are prone to instabilities
(Jönsson et al., 2006; Smith et al., 2006; Stoma et al., 2008)
and that this might be in part due to our partial knowledge
of the PIN1 polarization mechanisms. However, beyond
that, it could also be because other mechanisms regulating auxin homeostasis have been left out. For example,
none of the models have considered a positive-feedback
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
is developmentally regulated at the tissue level is still missing. Among the known regulators of PIN polarity, only PID
has been shown to play a role in organogenesis at the SAM
(Christensen et al., 2000; Friml et al., 2004). PID is expressed
specifically in organ boundaries (Christensen et al., 2000;
Furutani et al., 2004) and is unlikely to account for the complex dynamics of PIN polarization that underlie phyllotaxis
(Vernoux et al., 2010).
To overcome the limited biological information, and
inspired by the self-organization properties predicted by the
dynamical models of phyllotaxis, a number of cell-based
theoretical models have been developed to test whether the
PIN1 network self-organizes, and to explore plausible scenarios that could explain self-regulatory properties of the
auxin transport system in the SAM (reviewed by Wabnik
et al., 2011). Such models have been developed on the basis
of very simple hypotheses. A first set of models, inspired by
the seminal work of Tsvi Sachs (1969) on vascular development, is based on the assumption that cells can perceive the
flux of auxin across the plasma membrane and enhance it
by polarizing the efflux carriers in the direction of the preexisting flow (flux-based models). Such a feedback mechanism (canalization hypothesis) has the potential to canalize
auxin transport from sites with high hormone concentration
toward sites with low hormone concentration (Mitchison
et al., 1981). Although an auxin flux sensing mechanism
remains to be demonstrated (see below), flux-based models
can accurately predict PIN expression domains in the leaf
vasculature similar to those that have been observed experimentally (Rolland-Lagan and Prusinkiewicz, 2005; Scarpella
et al., 2006). For phyllotactic patterning, Stoma et al. (2008)
had to postulate two different canalization regimes (i.e. two
different feedback functions linking auxin flux to PIN protein insertion in membranes) in order to explain polarization
of PIN1 towards sites of high auxin concentration during
organogenesis. The weak regime (linear function) is active on
the SAM surface, while the strong regime (quadratic function) is active in the provasculature underneath. With the
assumption that, once established, sites of high auxin concentration act as sink by pumping auxin downwards by triggering the establishment of provascular cells, the model can
predict PIN1 polarization towards the auxin gradient and
the formation of inhibitory fields around the organ, as well
as PIN1 polarization in the provasculature (Stoma et al.,
2008). A second type of model was based on the hypothesis that, in a cell, PIN1 polarizes towards the cell neighbour
that possesses the highest concentration of auxin (concentration-based models). Simulations using concentrationbased models could reproduce realistic PIN1 polarization
and phyllotactic patterns in the SAM (Jönsson et al., 2006;
Smith et al., 2006). As such, concentration-based models can
also predict phyllotactic patterning, implying that flux-based
mechanisms must co-exist in the plant to regulate other
auxin transport-dependent patterning processes such provasculature development or leaf venation. Therefore, a third
model, integrating both the concentration-based and fluxbased hypotheses was developed. This model assumes that
cells are able to sense both auxin fluxes and concentrations
2584 | Sassi and Vernoux
Spatio-temporal regulation of auxin
signalling capacities also contributes to
the SAM patterning
The action of auxin in a patterning process depends not
only on the distribution of auxin in a tissue but also on the
cellular capacities to sense auxin and trigger specific transcriptional responses. Protein–protein interactions between
the Aux/IAA and auxin response factors (ARF) transcriptional regulators are central to auxin signalling. There are
29 Aux/IAA genes that encode mostly short-lived repressors of auxin-induced transcription. The 23 ARF proteins
can be either activators (the five Q-rich ARFs) or repressors of transcription. Aux/IAA and ARF proteins are able
to form homo- and heterodimers both within and between
the families. The instability of the Aux/IAAs is intrinsic to
the so-called domain II, which interacts directly with SCFlike ubiquitin protein ligases harbouring TIR1 or one of
the three related AFB F-box proteins (Dharmasiri et al.,
2005a,b; Kepinski and Leyser, 2005; Tan et al., 2007). TIR1
and the AFBs act as auxin co-receptors together with the
Aux/IAAs (Calderón-Villalobos et al., 2012), and their
activation leads to auxin-dependent degradation of Aux/
IAAs. A model for auxin transduction is that Aux/IAAs
dimerize with the ARF activators. These complexes bind
to auxin-inducible genes, thus preventing transcription. By
promoting the degradation of the Aux/IAAs, auxin would
allow the ARFs to activate transcription. Most Aux/IAAs
are themselves targets of the ARFs, thus establishing a
negative-feedback loop.
Treatment of pin1 meristems with exogenous auxin demonstrated that all the cells at the periphery of the meristem are competent to initiate organs in response to auxin
(Reinhardt et al., 2000). However, auxin cannot trigger
organogenesis at the centre of the meristem. Mutation in
MONOPTEROS/ARF5 (MP/ARF5) induces a pin-like phenotype and also blocks the ability of the PZ cells to respond
to exogenous auxin (Hardtke and Berleth, 1998; Reinhardt
et al., 2003). MP/ARF5 has been shown to exert its function in organogenesis, in part, by directly inducing, in an
auxin–dependent manner, the expression of LFY, ANT, and
ANT-like 6 (AIL6) TFs (Yamaguchi et al., 2013). As MP/
ARF5 is only expressed at the meristem periphery (Hardtke
and Berleth, 1998; Yamaguchi et al., 2013), the competence
for organ initiation at the periphery of the meristem thus
depends, at least in part, on a spatial modulation of auxin
signal transduction. Using mostly in situ hybridization, a
recent study identified TIR1, AFB1, and AFB5 and over 20
Aux/IAAs and ARFs as the effectors of auxin perception and
signalling in the SAM. This analysis also demonstrated that
most of these regulators are expressed differentially in the
SAM, with a low expression at the centre of the SAM and
high expression in the PZ. Notably, both ARF repressor and
ARF activator expression followed this general trend. A full
Aux/IAA–ARF interactome was obtained using a yeast twohybrid approach, and knowledge on the topology of the Aux/
IAA–ARF auxin signalling pathway was used to develop a
mathematical model of the control of gene transcription by
auxin in the SAM. This model predicted a role for the Aux/
IAA–ARF pathway in creating a differential sensitivity to
auxin between the centre and the periphery of the SAM,
together with a capacity to buffer fluctuations in the auxin
signal to stabilize the transcriptional response. These two
predictions could be validated by analysing the differences
between the spatio-temporal patterns of fluorescence of DIIVENUS and DR5 (Vernoux et al., 2011).
This work shows that the spatial distribution of ARF
genes in the SAM is essential to restrict high transcriptional
responses to auxin (including Aux/IAA expression) at the
periphery of the meristem and is essential to the definition
of the two main functional domains of the SAM, the CZ and
the PZ. In addition, the role of the signalling pathway in stabilizing transcriptional responses to auxin probably contributes to the robustness of patterning at the SAM and thus of
phyllotaxis (Vernoux et al., 2011). The dynamics of morphogenesis at the SAM thus appear to result from a dynamic integration of both spatial distribution of the auxin signal and
local signalling capacities in the spatial control of morphogenesis (Fig. 1), similarly to what has been observed recently
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
mechanism between auxin transport and its biosynthesis
that is supported by experimental evidence (Ljung et al.,
2001; Cheng et al., 2007). Recent breakthrough experiments have shown that auxin biosynthesis is regulated
mainly through a two-step pathway implicating the production of indole-3-pyruvic acid from tryptophan by members
of the TRYPTOPHAN AMINOTRANSFERASE OF
ARABIDOPSIS (TAA) family, followed by the conversion
of indole-3-pyruvic acid to IAA by YUCCA (YUC) flavin monooxygenase-like proteins (Mashiguchi et al., 2011;
Won et al., 2011). Several yuc mutant combinations display leaf and inflorescence developmental defects (Cheng
et al., 2006). Pin-like structures are also observed in the
yuc1,2,4,6 quadruple mutant and the expression patterns
of the YUC genes indicate that the YUC pathway regulates
lateral organ initiation and development of spatial and
temporal control of auxin biosynthesis. Cheng et al. (2007)
further observed that combining the pin1 mutation with
yuc1 and yuc4 mutations enhanced the pin1 organogenesis
phenotype, suggesting a coordinated action between auxin
transport and biosynthesis in the SAM. Feedback between
auxin transport and biosynthesis could provide robustness
to the auxin distribution dynamics. Indeed, the expression
YUC1 and YUC4 in the SAM is under the control of the
AP2 transcription factors of the PLETHORA (PLT) family (Pinon et al., 2013). It was proposed that phyllotactic
defects observed in the plt3plt5plt7 mutants arise from a
perturbation in auxin biosynthesis in the SAM, which, in
turn, alters PIN1 accumulation at the sites of incipient
primordia (Pinon et al., 2013). Further work on the spatial control of auxin biosynthesis and incorporating auxin
biosynthesis into the computational models of phyllotaxis
are likely to be crucial in refining our understanding of the
auxin-dependent self-organizing properties of patterning in
the SAM.
Auxin at the shoot apical meristem | 2585
for morphogen-driven developmental patterning in animal
systems (Kicheva et al., 2012).
Feedback between auxin-mediated
patterning and morphogenesis: a role for
biomechanics
Cell-wall properties and morphogenesis at the SAM
As discussed above, the coordinated regulation of auxin
transport and signalling provides instructions for patterned
organogenesis at the SAM. Such instructions will determine
local changes in cell growth that underlie primordium formation, but how is this achieved? How are the initial instructions
translated into a final shape? Plant cells are linked together
by the cell wall, which prevents cell sliding and migration,
and thus plant morphogenesis is the result of the coordination between localized cell division and selective cell growth
(Somerville et al., 2004; Cosgrove, 2005). At the cellular level,
growth is determined by the concerted action of the turgor
pressure of the protoplasm and the mechanical properties
of the cell wall. A plant cell can be abstracted as an inflated
balloon inside a rigid but deformable box: the pressure inside
the balloon will drive the growth of the whole structure if it
can overcome the physical resistance of the box. As such, the
water uptake into the plant cell will increase the turgor pressure, which, in turn, will trigger the extension of the pre-existing cell wall. At the same time, new polymers are synthesized
by the cell and delivered to the wall to avoid an excessive thinning and weakening of the structure (Cosgrove, 2005; Murray
et al., 2012). At the biochemical level, the cell wall is composed of stiff polysaccharides made of cellulose microfibrils
that are embedded and cross-linked in a matrix of hemicelluloses, pectins, and structural proteins: the chemical interactions between these components are commonly thought
to determine the mechanical properties of the cell wall (for
reviews on cell-wall biochemistry, see Somerville et al., 2004;
Cosgrove, 2005).
The irreversible extension of the cell wall during growth
(loosening) has so far been associated with acidic conditions
and the activity of some wall-loosening proteins such as
expansins that cleave the cross-links between polysaccharides
complexes, causing wall relaxation (Cosgrove, 2005; Murray
et al., 2012). Auxin-mediated cell growth has classically been
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Fig. 1. Multiple feedback loops acting across different scales drive auxin-driven organ formation at the SAM. Auxin accumulation in
cells at specific locations of the PZ in the SAM triggers both TIR1/AFB- and ABP1-mediated signalling. The activation of cellular auxin
responses promotes: (1) PIN1-mediated auxin efflux that will in turn influence polar auxin transport patterns in the SAM; and (2) the
level of auxin biosynthesis in the SAM. These will in turn feed back on the patterns of auxin accumulation amplifying the local growth
responses that promote organ formation.
2586 | Sassi and Vernoux
The process of morphogenesis is not only determined by
the growth rates of the cells within a tissue, as oriented cell
expansion along a preferential axis (anisotropy) also plays
an important role in the acquisition of the final shape (Coen
et al., 2004; Baskin, 2005; Murray et al., 2012). At a cellular level, anisotropy is established via the spatial distribution of the microfibrillar component of the cell wall. In fact,
the ordered deposition of cellulose microfibrils in transverse
hoops around the cell will direct the growth towards the axis
that is perpendicular to the orientation of cellulose deposition (Lloyd and Chan, 2004). Compelling evidence indicates
that the microtubular cytoskeleton plays an important role in
determining cell anisotropy: microtubules control the insertion and movement of cellulose synthase complexes on the
plasma membrane, thereby guiding the spatial deposition
of cellulose microfibrils in the cell wall (Paredez et al., 2006;
Crowell et al., 2009; Gutierrez et al., 2009; Bringmann et al.,
2012; Li et al., 2012). The functional link between microtubules and cellulose deposition provides an easy way to investigate the contribution of anisotropy to plant morphogenesis.
Microtubules can easily be tagged with fluorescent proteins, allowing indirect visualization of cellulose deposition
and growth anisotropy in the SAM in live imaging experiments (Hamant et al., 2008; Heisler et al., 2010; Uyttewaal
et al., 2012). Moreover, the use of microtubule-depolymerizing drugs as well as of mutants with defective microtubule
dynamics allow easy manipulation of the growth anisotropy
(Hamant et al., 2008; Uyttewaal et al., 2012). By using a
combination of these experimental approaches, it was demonstrated that growth anisotropy plays a relevant role in
morphogenesis at the SAM. Ordered microtubule alignment
marks the boundary cells that physically separate the SAM
and the primordium (Hamant et al., 2008; Uyttewaal et al.,
2012). In the absence of microtubules, the tissue folding that
allows organ separation from the SAM is abolished, despite
a correct specification of the boundary cells (Hamant et al.,
2008). Growth anisotropy was also shown to contribute to the
pin-like shape of the SAM in the absence of organ formation.
In a naked meristem, microtubules align circumferentially
around the SAM periphery, imposing cell anisotropic growth
along the vertical axis of the stem. When microtubules are
depolymerized, growth becomes isotropic, leading the SAM
to acquire a globular shape that has the geometric properties
of soap froth (Grandjean et al., 2004; Hamant et al., 2008;
Corson et al., 2009).
Microtubules might mediate a mechanical feedback
during morphogenesis
While the mechanical changes associated with morphogenesis have classically been considered to be downstream of
the genetic and biochemical inputs, an opposite trend proposing a direct role of mechanical forces on developmental
patterning has been gaining strong support over the years
(Selker et al., 1992; Hamant et al., 2010; Uyttewaal et al.,
2010; Mirabet et al., 2011). The hypothesis of a biophysical
patterning mechanism is based on the assumption that the
SAM surface is under tension. This is because cells in the L1
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
explained by the acid growth hypothesis: auxin would promote the acidification of the cell wall, presumably by activating some plasma membrane H+-ATPases that release protons
outside the cell (Christian et al., 2006). The low cell-wall pH
would in turn increase the activity of expansins, causing cellwall relaxation (Cosgrove, 2005; Sánchez-Rodríguez et al.,
2010; Murray et al., 2012). While this mechanism would
account for very fast and transcription-independent loosening responses, evidence also shows that auxin regulates
the expression of a number of cell-wall-modifying proteins
including expansins and pectin methyl estherases (PMEs)
(Overvoorde et al., 2005), probably promoting sustained
loosening effects.
Concerning the SAM, an increasing amount of evidence
suggests that cell-wall modifications underlie organ formation. First, local application of exogenous expansins on the
SAM surface has been shown to promote the formation
of outgrowths that do not develop into proper organs but
have the potential to alter phyllotaxis (Fleming et al., 1997).
More recently, PMEs and their inhibitors (PMEIs) have been
involved in the control of SAM morphogenesis (Peaucelle
et al., 2008, 2011). PMEs and PMEIs are thought to modify
the fluidity of the pectin matrix and, in turn, the permeability
to wall-loosening agents, presumably leading to softer or stiffer
walls, respectively (Peaucelle et al., 2008). Accordingly, local
PME treatments on the SAM surface could promote organ
outgrowth, whereas PMEI overexpression inhibits organogenesis at the SAM, leading to pin-like phenotypes (Peaucelle
et al., 2008). Together, these pieces of evidence suggest a scenario in which auxin accumulation at the site of incipient
primordia triggers cell-wall loosening via acidic growth and
subsequent induction of wall-modifying enzymes, thereby
promoting organogenesis. This idea is further supported by
recent measurements of cell-wall mechanical properties on
the SAM surface, as well as by finite-element modelling, suggesting that the slow-growing cells at the top of the SAM
might have stiffer cell walls compared with the fast-growing
cells at the periphery (Milani et al., 2011; Kierzkowski et al.,
2012). However, cell-wall remodelling might not occur only at
the meristem surface. Changes in the cell-wall elastic properties of subepidermal cells at the site of incipient primordia
have also been reported, suggesting that cell expansion is initiated in the internal cell layers (Peaucelle et al., 2011). These
results are in line with previous works showing that transgenic expression of expansins in the inner tissues of the SAM
is capable of initiating the entire programme of leaf development in tomato (Solanum lycopersicon) apices (Pien et al.,
2001). While all these elements suggest an interesting framework that links auxin to cell-wall-controlled morphogenesis,
it has to be pointed out that clear genetic evidence causally
linking cell-wall loosening to in planta morphogenesis is still
missing. However, auxin action on cell growth at the SAM
might not be limited to cell-wall loosening, but might also
involve changes in turgor pressure (Christian et al., 2006). An
attractive hypothesis is that auxin could regulate the activity
of aquaporins, as has been demonstrated recently for lateral
root formation (Péret et al., 2012), although this remains to
be proven in the SAM.
Auxin at the shoot apical meristem | 2587
membrane have been shown to limit the lateral diffusion of
PIN proteins, thus contributing in the maintenance of PIN
polarity (Feraru et al., 2011). Along the same line, it has been
shown that in tomato (S. lycopersicon) apices, PIN1 levels and
intracellular localization can be altered by treatments that
cause cell-wall deformation or that modify plasma membrane
properties. These findings suggest that cell-wall deformations
caused by mechanical stress are sensed via the plasma membrane and affect PIN1 intracellular localization, possibly by
altering the endocytosis/exocytosis ratio (Nakayama et al.,
2012).
Taken together, these lines of evidence support a scenario
in which morphogenesis is the result of the coordinated action
not only of genetic and biochemical pathways but also of the
mechanical forces at the SAM surface (Fig. 2). This vision
is also supported by theoretical works that suggest a plausible scenario in which biophysical and biochemical mechanisms compete and cooperate to generate robust phyllotactic
patterns: fluctuations of auxin concentrations could modify
the mechanical properties of the L1, causing uneven tissue
growth that would further alter the stress distribution, eventually driving changes in auxin distribution (Newell et al.,
2008a,b). It is likely that only through the integration of both
biochemical and mechanical pathways will the mechanisms
controlling the dynamics of PIN polarization in SAM tissues
be elucidated.
Conclusions
From the body of work we have discussed in this review, the
SAM can be viewed as a typical complex system in which
interactions between elements that constitute the system at
a given scale (e.g. the cells) allow the emergence of a behaviour at a higher scale (e.g. the tissue). Phyllotaxis provides
a perfect illustration of this vision: the dynamics of tissue
­patterning emerge from repulsive interactions between organs
that are themselves determined by rules of signal exchange
between individual cells. Auxin is a central signal that acts
at these different scales, but it is probably not the only one.
We have indeed seen that different feedback mechanisms
are key to controlling the self-organization properties of the
SAM system: feedback between auxin concentration or flux
and the polarity of the PIN1 pumps, feedback between auxin
transport and auxin biosynthesis that remain to be fully
analysed, feedback between auxin and its signalling pathways, and feedback between auxin and the geometry of the
tissue through mechanics. At this point, the complexity of
this developmental system clearly exceeds what can be understood intuitively, explaining why modelling approaches have
been so instrumental in extracting plausible hypotheses from
the data. Thus, we anticipate that modelling will remain central in the analysis of SAM function and phyllotaxis, together
with further dissections of the molecular mechanisms implicated. We have already discussed the fact that understanding
the cellular mechanisms of PIN polarity and the control of
auxin biosynthesis are likely to be crucial to understand how
robust phyllotactic patterns can be generated. Concerning
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
layer are subject to an upward pressure caused by the turgor of inner cells. The unbalance between the inner turgor
pressure and surface tension leads to the buckling of the L1
that underlies the formation of an organ. The resulting physical deformation of the SAM surface generates folded areas
where organogenesis is prevented, thus generating physicsbased inhibitory fields (Selker et al., 1992; Green, 1996,
1999). Deformations of the SAM surface might arise at the
cellular level when neighbouring cells grow at very different
rates (i.e. boundary cells versus primordium cells), triggering
changes in surface tension that could feed back on local cellexpansion rates (Murray et al., 2012). Hamant et al. (2008)
indeed showed that microtubule alignment on the SAM surface matched the theoretical distribution of the main force
directions, calculated on a purely geometrical basis. This
would suggest that microtubules might sense, through an
unknown mechanism, the physical stress on the SAM surface
and align accordingly to guide the anisotropic cellulose deposition, thus counteracting the stress. As a result, the oriented
microtubule pattern in boundary cells that underlies organ
separation would be generated by the action of mechanical
forces (Hamant et al., 2008). The idea that microtubules react
to mechanical changes at the SAM surface is also supported
by SAM compressions, laser ablations, and pharmacological
treatments weakening the cell wall, which all trigger a reorientation within a few hours of the microtubules according
to the predicted stress patterns generated by the treatments
(Hamant et al., 2008; Heisler et al., 2010; Uyttewaal et al.,
2012). This stress-driven microtubule reorientation has been
shown to depend on the action of the microtubule-severing
protein katanin (AtKTN1) (Uyttewaal et al., 2012).
In addition, patterns of microtubule orientation at the SAM
surface have also been suggested to feed back on the auxin
transport system. Indeed, in SAM cells, a negative correlation
between microtubule alignment and PIN1 polarity has been
observed: PIN1 localizes to anticlinal walls parallel to the
main microtubule orientation (Heisler et al., 2010). Although
PIN localization is unlikely to be under the direct control
of microtubules (Geldner et al., 2001; Heisler et al., 2010),
PIN1 has been shown to lose its cellular polarity after long
treatments with microtubule-depolymerizing drugs (Heisler
et al., 2010). In addition, PIN1 also reorients its polarity
away from wounded cells in laser-ablation experiments with
a dynamic similar to that observed for microtubules (Heisler
et al., 2010). This would suggest that mechanical forces might
influence PIN polarity and, as a result, auxin transport patterns. Computer simulations showed that a mechanism that
distributes PIN1 polarity according to cell-wall stress behaves
like a concentration-based model for auxin transport and is
indeed capable of generating phyllotactic patterns (Heisler
et al., 2010). This is in contrast to the results of Wabnik et al.
(2010), which instead support canalization, thus questioning
again which mechanism best explains phyllotactic patterning.
However, further supporting the idea of a mechanical control of the auxin transport system, evidence that the cell wall
plays a role in determining PIN polarity has been reported. In
particular, cellulose-based mechanical connections between
the extracellular matrix and the polar domains of the plasma
2588 | Sassi and Vernoux
the role of mechanics, it should be pointed out that the contribution of biophysical pathways to plant morphogenesis
is still controversial, because of the difficulty of experimentally measuring mechanical forces in the SAM (Kuhlemeier,
2007; Besnard et al., 2011). New technological developments
will be necessary to convincingly demonstrate the role of
mechanical forces in phyllotaxis at the SAM alongside auxin.
Sensor mechanisms for mechanical forces are also required
and remain to be identified. Finally, it should be noted that
the theoretical approaches to phyllotaxis have led to an idealistic view of this developmental process. However, it can
be expected that the phyllotactic patterning will be prone to
noise like any developmental system and will thus show a
certain level of variability. Recent analyses have indeed demonstrated deviations from the Fibonacci angle in Arabidopsis
(Peaucelle et al., 2007, Refahi et al., 2011), which probably
reflect the impact of noise on phyllotaxis (Mirabet et al.,
2012). A proper quantification of the dynamics of organ initiation at the SAM using, for example, data obtained from
live imaging (Grandjean et al., 2004, Heisler et al., 2005,
Fernandez et al., 2010) is undoubtedly necessary to define
the characteristics of the phyllotactic system. Such an analysis will be crucial to evaluate the predictive capacities of the
models that will be used to understand how the different levels of control, from auxin to mechanics, are integrated in the
SAM to generate robust phyllotactic patterns.
Acknowledgements
We apologize to all colleagues whose original works could
not be cited for space constraints. We would like to thank
Olivier Hamant for critical reading of the manuscript. The
work of the authors is funded by the Agence National de la
Recherche and the iSAM ERASysBIO+ programme.
References
Abas L, Benjamins R, Malenica N, et al... 2006. Intracellular
trafficking and proteolysis of the Arabidopsis auxin-efflux facilitator
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Fig. 2. Cross-talks between auxin and mechanical signalling in the regulation of morphogenesis at the SAM. Auxin-induced cell-wall
loosening is thought to cause mechanical stress that in turn feeds back on PIN1-mediated auxin transport to amplify local growth
responses (see text for details). Dashed lines represent putative interactions: (1) auxin might also regulate the turgor pressure of the SAM
cells, probably by modulating aquaporins activity, as observed recently in the process of lateral root formation (Peret et al., 2012); (2)
the mechanical stresses that determine microtubule orientation (Hamant et al., 2008; Heisler et al., 2010; Uyttewaal et al., 2012) might
be sensed via the plasma membrane, as proposed recently for the mechanical stress-driven changes in PIN1 intracellular distribution
(Nakayama et al., 2012).
Auxin at the shoot apical meristem | 2589
PIN2 are involved in root gravitropism. Nature Cell Biology 8,
249–256.
Adler I, Barabe D, Jean R. 1997. A history of the study of
phyllotaxis. Annals of Botany 80, 231–244.
Calderón-Villalobos LI, Lee S, De Oliveira C, et al. 2012. A
combinatorial TIR1/AFB-Aux/IAA co-receptor system for differential
sensing of auxin. Nature Chemical Biology 8, 477–485.
Aida M, Tasaka M. 2006. Morphogenesis and patterning at the
organ boundaries in the higher plant shoot apex. Plant Molecular
Biology 60, 915–928.
Cheng Y, Dai X, Zhao Y. 2006. Auxin biosynthesis by the YUCCA
flavin monooxygenases controls the formation of floral organs
and vascular tissues in Arabidopsis. Genes & Development 20,
1790–1799.
Bainbridge K, Guyomarc’h S, Bayer E, Swarup R, Bennett
M, Mandel T, Kuhlemeier C. 2008. Auxin influx carriers stabilize
phyllotactic patterning. Genes & Development 22, 810–823.
Cheng Y, Dai X, Zhao Y. 2007. Auxin synthesized by the YUCCA
flavin monooxygenases is essential for embryogenesis and leaf
formation in Arabidopsis. Plant Cell 19, 2430–2439.
Barbier de Reuille P, Bohn-Courseau I, Ljung K, Morin H,
Carraro N, Godin C, Traas J. 2006. Computer simulations reveal
properties of the cell-cell signaling network at the shoot apex in
Arabidopsis. Proceedings of the National Academy of Sciences, USA
103, 1627–1632.
Christensen SK, Dagenais N, Chory J, Weigel D. 2000.
Regulation of auxin response by the protein kinase PINOID. Cell 100,
469–478.
Baskin TI. 2005. Anisotropic expansion of the plant cell wall. Annual
Review of Cell and Developmental Biology 21, 203–222.
Bayer EM, Smith RS, Mandel T, Nakayama N, Sauer M,
Prusinkiewicz P, Kuhlemeier C. 2009. Integration of transportbased models for phyllotaxis and midvein formation. Genes &
Development 23, 373–384.
Benjamins R, Scheres B. 2008. Auxin: the looping star in plant
development. Annual Review of Plant Biology 59, 443–465.
Benková E, Michniewicz M, Sauer M, Teichmann T, Seifertová
D, Jürgens G, Friml J. 2003. Local, efflux-dependent auxin gradients
as a common module for plant organ formation. Cell 115, 591–602.
Besnard F, Vernoux T, Hamant O. 2011. Organogenesis from stem
cells in planta: multiple feedback loops integrating molecular and
mechanical signals. Cellular and Molecular Life Sciences 68, 2885–2906.
Blakeslee JJ, Bandyopadhyay A, Lee OR, et al. 2007.
Interactions among PIN-FORMED and P-glycoprotein auxin
transporters in Arabidopsis. Plant Cell 19, 131–147.
Bowman JL, Eshed Y, Baum SF. 2002. Establishment of polarity in
angiosperm lateral organs. Trends in Genetics 18, 134–141.
Brand U, Fletcher JC, Hobe M, Meyerowitz EM, Simon R. 2000.
Dependence of stem cell fate in Arabidopsis on a feedback loop
regulated by CLV3 activity. Science 289, 617–619.
Bringmann M, Li E, Sampathkumar A, Kocabek T, Hauser MT,
Persson S. 2012. POM-POM2/cellulose synthase interacting1 is
essential for the functional association of cellulose synthase and
microtubules in Arabidopsis. Plant Cell 24, 163–177.
Brunoud G, Wells DM, Oliva M, et al. 2012. A novel sensor to map
auxin response and distribution at high spatio-temporal resolution.
Nature 482, 103–106.
Byrne ME, Barley R, Curtis M, Arroyo JM, Dunham M, Hudson A,
Martienssen RA. 2000. Asymmetric leaves1 mediates leaf patterning
and stem cell function in Arabidopsis. Nature 408, 967–971.
Byrne ME. 2006. Shoot meristem function and leaf polarity: the role
of class III HD-ZIP genes. PLoS Genetics 2, e89.
Coen E, Rolland-Lagan AG, Matthews M, Bangham JA,
Prusinkiewicz P. 2004. The genetics of geometry. Proceedings of
the National Academy of Sciences, USA 101, 4728–4735.
Corson F, Hamant O, Bohn S, Traas J, Boudaoud A, Couder Y.
2009. Turning a plant tissue into a living cell froth through isotropic
growth. Proceedings of the National Academy of Sciences, USA 106,
8453–8458.
Cosgrove DJ. 2005. Growth of the plant cell wall. Nature Reviews
Molecular Cell Biology 6, 850–861.
Crowell EF, Bischoff V, Desprez T, Rolland A, Stierhof YD,
Schumacher K, Gonneau M, Höfte H, Vernhettes S. 2009.
Pausing of Golgi bodies on microtubules regulates secretion
of cellulose synthase complexes in Arabidopsis. Plant Cell 21,
1141–1154.
Dharmasiri N, Dharmasiri S, Estelle M. 2005a. The F-box protein
TIR1 is an auxin receptor. Nature 435, 441–445.
Dharmasiri N, Dharmasiri S, Weijers D, Lechner E, Yamada M,
Hobbie L, Ehrismann JS, Jürgens G, Estelle M. 2005b. Plant
development is regulated by a family of auxin receptor F box proteins.
Developmental Cell 9, 109–119.
Dhonukshe P, Aniento F, Hwang I, Robinson DG, Mravec
J, Stierhof YD, Friml J. 2007. Clathrin-mediated constitutive
endocytosis of PIN auxin efflux carriers in Arabidopsis. Current Biology
17, 520–527.
Dhonukshe P, Huang F, Galvan-Ampudia CS, et al. 2010. Plasma
membrane-bound AGC3 kinases phosphorylate PIN auxin carriers at
TPRXS(N/S) motifs to direct apical PIN recycling. Development 137,
3245–3255.
Dhonukshe P, Tanaka H, Goh T, et al. 2008. Generation of cell
polarity in plants links endocytosis, auxin distribution and cell fate
decisions. Nature 456, 962–966.
Dodsworth S. 2009. A diverse and intricate signalling network
regulates stem cell fate in the shoot apical meristem. Developmental
Biology 336, 1–9.
Douady S, Couder Y. 1996a. Phyllotaxis as a dynamical self
organizing process. Part I: the spiral modes resulting from timeperiodic iterations. Journal of Theoretical Biology 178, 255–274.
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Baster P, Robert S, Kleine-Vehn J, Vanneste S, Kania U,
Grunewald W, De Rybel B, Beeckman T, Friml J. 2012.
SCFTIR1/AFB-auxin signalling regulates PIN vacuolar trafficking and
auxin fluxes during root gravitropism. EMBO Journal 32, 260–274.
Christian M, Steffens B, Schenck D, Burmester S, Böttger M,
Lüthen H. 2006. How does auxin enhance cell elongation? Roles
of auxin-binding proteins and potassium channels in growth control.
Plant Biology 8, 346–352.
2590 | Sassi and Vernoux
Douady S, Couder Y. 1996b. Phyllotaxis as a dynamical selforganizing process. Part II: the spontaneous formation of a periodicity
and the coexistance of spiral and whorled patterns. Journal of
Theoretical Biology 178, 275–294.
Douady S, Couder Y. 1996c. Phyllotaxis as a dynamical self
organizing process. Part III: the simulation of the transient regimes of
ontogeny. Journal of Theoretical Biology 178, 295–312.
Endrizzi K, Moussian B, Haecker A, Levin JZ, Laux T. 1996.
The SHOOT MERISTEMLESS gene is required for maintenance of
undifferentiated cells in Arabidopsis shoot and floral meristems and
acts at a different regulatory level than the meristem genes WUSCHEL
and ZWILLE. The Plant Journal 10, 967–979.
Feraru E, Feraru MI, Kleine-Vehn J, Martinière A, Mouille G,
Vanneste S, Vernhettes S, Runions J, Friml J. 2011. PIN polarity
maintenance by the cell wall in Arabidopsis. Current Biology 21,
338–343.
Fleming AJ, McQueen-Mason S, Mandel T, Kuhlemeier C. 1997.
Induction of leaf primordia by the cell wall protein expansin. Science
276, 1415–1418.
Friml J. 2010. Subcellular trafficking of PIN auxin efflux carriers in
auxin transport. European Journal of Cell Biology 89, 231–235.
Friml J, Yang X, Michniewicz M, et al. 2004. A PINOID-dependent
binary switch in apical-basal PIN polar targeting directs auxin efflux.
Science 306, 862–865.
Furutani M, Vernoux T, Traas J, Kato T, Tasaka M, Aida M.
2004. PIN-FORMED1 and PINOID regulate boundary formation and
cotyledon development in Arabidopsis embryogenesis. Development
131, 5021–5030.
Gälweiler L, Guan C, Müller A, Wisman E, Mendgen K,
Yephremov A, Palme K. 1998. Regulation of polar auxin transport
by AtPIN1 in Arabidopsis vascular tissue. Science 282, 2226–2230.
Geldner N, Friml J, Stierhof YD, Jürgens G, Palme K. 2001. Auxin
transport inhibitors block PIN1 cycling and vesicle trafficking. Nature
413, 425–428.
Grandjean O, Vernoux T, Laufs P, Belcram K, Mizukami Y, Traas
J. 2004. In vivo analysis of cell division, cell growth, and differentiation
at the shoot apical meristem in Arabidopsis. Plant Cell 16, 74–87.
Green PB. 1996. Transductions to generate plant form and pattern:
an essay on cause and effect. Annals of Botany 78, 269–281.
Green PB. 1999. Expression of pattern in plants: combining
molecular and calculus-based biophysical paradigms. American
Journal of Botany 86, 1059–1076.
Guenot B, Bayer E, Kierzkowski D, Smith RS, Mandel T,
Žádníková P, Benková E, Kuhlemeier C. 2012. PIN1-independent
leaf initiation in Arabidopsis. Plant Physiology 159, 1501–1510.
Gutierrez R, Lindeboom JJ, Paredez AR, Emons AM, Ehrhardt
DW. 2009. Arabidopsis cortical microtubules position cellulose
synthase delivery to the plasma membrane and interact with
cellulose synthase trafficking compartments. Nature Cell Biology 11,
797–806.
Hamant O, Traas J, Boudaoud A. 2010. Regulation of shape
and patterning in plant development. Current Opinion in Genetics &
Development 20, 454–459.
Hardtke CS, Berleth T. 1998. The Arabidopsis gene MONOPTEROS
encodes a transcription factor mediating embryo axis formation and
vascular development. EMBO Journal 17, 1405–1411.
Heisler MG, Hamant O, Krupinski P, Uyttewaal M, Ohno C,
Jönsson H, Traas J, Meyerowitz EM. 2010. Alignment between
PIN1 polarity and microtubule orientation in the shoot apical meristem
reveals a tight coupling between morphogenesis and auxin transport.
PLoS Biology 8, e1000516.
Heisler MG, Ohno C, Das P, Sieber P, Reddy GV, Long JA,
Meyerowitz EM. 2005. Patterns of auxin transport and gene
expression during primordium development revealed by live imaging
of the Arabidopsis inflorescence meristem. Current Biology 15,
1899–1911.
Höfmeister W. 1868. Allgemeine Morphologie der Gewachse. In:
de Bary A, Irmisch TH, Sachs J, eds. Handbuch der Physiologischen
Botanik . Leipzig: Engelmann, 405–664.
Jönsson H, Heisler MG, Shapiro BE, Meyerowitz EM, Mjolsness
E. 2006. An auxin-driven polarized transport model for phyllotaxis.
Proceedings of the National Academy of Sciences, USA 103,
1633–1638.
Kepinski S, Leyser O. 2005. The Arabidopsis F-box protein TIR1 is
an auxin receptor. Nature 435, 446–451.
Kicheva A, Cohen M, Briscoe J. 2012. Developmental pattern
formation: insights from physics and biology. Science 338,
210–212.
Kierzkowski D, Nakayama N, Routier-Kierzkowska AL, Weber
A, Bayer E, Schorderet M, Reinhardt D, Kuhlemeier C, Smith
RS. 2012. Elastic domains regulate growth and organogenesis in the
plant shoot apical meristem. Science 335, 1096–1099.
Kleine-Vehn J, Dhonukshe P, Sauer M, Brewer PB, Wiśniewska
J, Paciorek T, Benková E, Friml J. 2008a. ARF GEF-dependent
transcytosis and polar delivery of PIN auxin carriers in Arabidopsis.
Current Biology 18, 526–531.
Kleine-Vehn J, Ding Z, Jones AR, Tasaka M, Morita MT, Friml J.
2010. Gravity-induced PIN transcytosis for polarization of auxin fluxes
in gravity-sensing root cells. Proceedings of the National Academy of
Sciences, USA 107, 22344–22349.
Kleine-Vehn J, Leitner J, Zwiewka M, Sauer M, Abas L,
Luschnig C, Friml J. 2008b. Differential degradation of PIN2 auxin
efflux carrier by retromer-dependent vacuolar targeting. Proceedings
of the National Academy of Sciences, USA 105, 17812–17817.
Kuhlemeier C. 2007. Phyllotaxis. Trends in Plant Science 12,
143–150.
Laux T, Mayer KF, Berger J, Jürgens G. 1996. The WUSCHEL
gene is required for shoot and floral meristem integrity in Arabidopsis.
Development 122, 87–96.
Leyser O. 2005. Auxin distribution and plant pattern formation: how
many angels can dance on the point of PIN? Cell 121, 819–822.
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Fernandez R, Das P, Mirabet V, Moscardi E, Traas J, Verdeil JL,
Malandain G, Godin C. 2010. Imaging plant growth in 4D: robust
tissue reconstruction and lineaging at cell resolution. Nature Methods
7, 547–553.
Hamant O, Heisler MG, Jönsson H, et al. 2008. Developmental
patterning by mechanical signals in Arabidopsis. Science 322,
1650–1655.
Auxin at the shoot apical meristem | 2591
Li S, Lei L, Somerville CR, Gu Y. 2012. Cellulose synthase
interactive protein 1 (CSI1) links microtubules and cellulose synthase
complexes. Proceedings of the National Academy of Sciences, USA
109, 185–190.
Ljung K, Bhalerao RP, Sandberg G. 2001. Sites and homeostatic
control of auxin biosynthesis in Arabidopsis during vegetative growth.
The Plant Journal 28, 465–474.
Ogawa M, Shinohara H, Sakagami Y, Matsubayashi Y. 2008.
Arabidopsis CLV3 peptide directly binds CLV1 ectodomain. Science
319, 294.
Okada K, Uedalb J, Komaki MK, Bell CJ, Shimura Y. 1991.
Requirement of the auxin polar transport system in early stages of
Arabidopsis floral bud formation. Plant Cell 3, 677–684.
Overvoorde PJ, Okushima Y, Alonso JM, et al. 2005. Functional
genomic analysis of the AUXIN/INDOLE-3-ACETIC ACID gene family
members in Arabidopsis thaliana. Plant Cell 17, 3282–3300.
Mashiguchi K, Tanaka K, Sakai T, et al. 2011. The main auxin
biosynthesis pathway in Arabidopsis. Proceedings of the National
Academy of Sciences USA 108, 18512–18517.
Paciorek T, Zazímalová E, Ruthardt N, et al. 2005. Auxin inhibits
endocytosis and promotes its own efflux from cells. Nature 435,
1251–1256.
Mayer KF, Schoof H, Haecker A, Lenhard M, Jürgens G, Laux T.
1998. Role of WUSCHEL in regulating stem cell fate in the Arabidopsis
shoot meristem. Cell 95, 805–815.
Paredez AR, Somerville CR, Ehrhardt DW. 2006. Visualization
of cellulose synthase demonstrates functional association with
microtubules. Science 312, 1491–1495.
Michniewicz M, Zago MK, Abas L, et al. 2007. Antagonistic
regulation of PIN phosphorylation by PP2A and PINOID directs auxin
flux. Cell 130, 1044–1056.
Peaucelle A, Braybrook SA, Le Guillou L, Bron E, Kuhlemeier C,
Höfte H. 2011. Pectin-induced changes in cell wall mechanics underlie
organ initiation in Arabidopsis. Current Biology 21, 1720–1726.
Milani P, Gholamirad M, Traas J, Arnéodo A, Boudaoud A,
Argoul F, Hamant O. 2011. In vivo analysis of local wall stiffness
at the shoot apical meristem in Arabidopsis using atomic force
microscopy. The Plant Journal 67, 1116–1123.
Peaucelle A, Louvet R, Johansen JN, Höfte H, Laufs P, Pelloux
J, Mouille G. 2008. Arabidopsis phyllotaxis is controlled by the
methyl-esterification status of cell-wall pectins. Current Biology 18,
1943–1948.
Mirabet V, Besnard F, Vernoux T, Boudaoud A. 2012. Noise and
robustness in phyllotaxis. PLoS Computational Biology 8, e1002389.
Peaucelle A, Morin H, Traas J, Laufs P. 2007. Plants expressing
a mir164-resistant CUC2 gene reveal the importance of postmeristematic maintenance of phyllotaxy in Arabidopsis. Development
134, 1045–1050.
Mirabet V, Das P, Boudaoud A, Hamant O. 2011. The role of
mechanical forces in plant morphogenesis. Annual Review of Plant
Biology 62, 365–385.
Mitchison GJ. 1977. Phyllotaxis and the Fibonacci series. Science
196, 270–275.
Mitchison GJ, Hanke DE, Sheldrake AR. 1981. The polar transport
of auxin and venation patterns in plants. Philosophical Transactions of
the Royal Society of London B 295, 461–471.
Mizukami Y, Fischer RL. 2000. Plant organ size control:
AINTEGUMENTA regulates growth and cell numbers during
organogenesis. Proceedings of the National Academy of Sciences,
USA 97, 942–947.
Perales M, Reddy GV. 2012. Stem cell maintenance in shoot apical
meristems. Current Opinion in Plant Biology 15, 10–16.
Péret B, Li G, Zhao J, et al. 2012. Auxin regulates aquaporin
function to facilitate lateral root emergence. Nature Cell Biology 14,
991–998.
Petrásek J, Mravec J, Bouchard R, et al. 2006. PIN proteins
perform a rate-limiting function in cellular auxin efflux. Science 312,
914–918.
Moyroud E, Kusters E, Monniaux M, Koes R, Parcy F. 2010.
LEAFY blossoms. Trends in Plant Science 15, 346–352.
Pien S, Wyrzykowska J, McQueen-Mason S, Smart C, Fleming
A. 2001. Local expression of expansin induces the entire process of
leaf development and modifies leaf shape. Proceedings of the National
Academy of Sciences, USA 98, 11812–11817.
Mravec J, Kubes M, Bielach A, Gaykova V, Petrásek J, Skůpa P,
Chand S, Benková E, Zazimalova E, Friml J. 2008. Interaction of
PIN and PGP transport mechanisms in auxin distribution-dependent
development. Development 135, 3345–3354.
Pinon V, Prasad K, Grigg SP, Sanchez-Perez GF, Scheres B.
2013. Local auxin biosynthesis regulation by PLETHORA transcription
factors controls phyllotaxis in Arabidopsis. Proceedings of the National
Academy of Sciences, USA 110, 1107–1112.
Murray JAH, Jones A, Godin C, Traas J. 2012. Systems Analysis
of Shoot Apical Meristem Growth and Development: Integrating
Hormonal and Mechanical Signaling. The Plant Cell 24, 3907–3919.
Rast MI, Simon R. 2008. The meristem-to-organ boundary:
more than an extremity of anything. Current Opinion in Genetics &
Development 18, 287–294.
Nakayama N, Smith RS, Mandel T, Robinson S, Kimura S,
Boudaoud A, Kuhlemeier C. 2012. Mechanical regulation of auxinmediated growth. Current Biology 22, 1468–1476.
Raven JA. 1975. Transport of indoleacetic acid in plant cells in
relation to pH and electrical potential gradients, and its significance for
polar IAA transport. New Phytologist 74, 163–172.
Newell AC, Shipman PD, Sun Z. 2008a. Phyllotaxis: cooperation
and competition between mechanical and biochemical processes.
Journal of Theoretical Biology 251, 421–439.
Refahi Y, Farcot E, Guedon Y, Besnard F, Vernoux T, Godin
C. (2011) A combinatorial model of phyllotaxis perturbations in
Arabidopsis thaliana. In: 22nd Annual Symposium on Combinatorial
Pattern Matching , Palermo, Italy, vol. 6661, 323–335. http://
www-sop.inria.fr/virtualplants/Publications/2011/RFGBVG11/. Last
accessed date : March 26, 2013.
Newell AC, Shipman PD, Sun Z. 2008b. Phyllotaxis as an example
of the symbiosis of mechanical forces and biochemical processes in
living tissue. Plant Signaling & Behavior 3, 586–589.
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Lloyd C, Chan J. 2004. Microtubules and the shape of plants to
come. Nature Reviews Molecular Cell Biology 5, 13–22.
2592 | Sassi and Vernoux
Reinhardt D, Frenz M, Mandel T, Kuhlemeier C. 2005.
Microsurgical and laser ablation analysis of leaf positioning and
dorsoventral patterning in tomato. Development 132, 15–26.
Reinhardt D. 2005a. Regulation of phyllotaxis. International Journal of
Developmental Biology 49, 539–46.
Reinhardt D. 2005b. Phyllotaxis—a new chapter in an old tale about
beauty and magic numbers. Current Opinion in Plant Biology 8,
487–493.
Reinhardt D, Mandel T, Kuhlemeier C. 2000. Auxin regulates
the initiation and radial position of plant lateral organs. Plant Cell 12,
507–518.
Reinhardt D, Pesce ER, Stieger P, Mandel T, Baltensperger K,
Bennett M, Traas J, Friml J, Kuhlemeier C. 2003. Regulation of
phyllotaxis by polar auxin transport. Nature 426, 255–260.
Robert S, Kleine-Vehn J, Barbez E, et al. 2010. ABP1 mediates
auxin inhibition of clathrin-dependent endocytosis in Arabidopsis. Cell
143, 111–121.
Rubery PH, Sheldrake AR. 1974. Carrier-mediated auxin transport.
Planta 121, 101–121.
Sablowski R. 2007. Flowering and determinacy in Arabidopsis.
Journal of Experimental Botany 58, 899–907.
Sachs T. 1969. Polarity and the induction of organized vascular
tissues. Annals of Botany 33, 263–275.
Sánchez-Rodríguez C, Rubio-Somoza I, Sibout R, Persson S.
2010. Phytohormones and the cell wall in Arabidopsis during seedling
growth. Trends in Plant Science 15, 291–301.
Sauer M, Balla J, Luschni, C, Wisniewska J, Reinöhl V, Friml
J. Benková E. 2006. Canalization of auxin flow by Aux/IAA-ARFdependent feedback regulation of PIN polarity. Genes & Development
20, 2902–2911.
Scarpella E, Marcos D, Friml J, Berleth T. 2006. Control of leaf
vascular patterning by polar auxin transport. Genes & Development
20, 1015–1027.
Schoof H, Lenhard M, Haecker A, Mayer KF, Jürgens G, Laux
T. 2000. The stem cell population of Arabidopsis shoot meristems
in maintained by a regulatory loop between the CLAVATA and
WUSCHEL genes. Cell 100, 635–644.
Selker JM, Steucek GL, Green PB. 1992. Biophysical mechanisms
for morphogenetic progressions at the shoot apex. Developmental
Biology 153, 29–43.
Shipman PD, Newell AC. 2005. Polygonal planforms and phyllotaxis
on plants. Journal of Theoretical Biology 236, 154–197.
Smith RS, Guyomarc’h S, Mandel T, Reinhardt D, Kuhlemeier C,
Prusinkiewicz P. 2006. A plausible model of phyllotaxis. Proceedings
of the National Academy of Sciences, USA 103, 1301–1306.
Snow M, Snow R. 1931. Experiments on phyllotaxis. I. The effect of
isolating a primordium. Philosophical Transactions of the Royal Society
London B 221, 1–43.
Somerville C, Bauer S, Brininstool G, et al. 2004. Toward a systems
approach to understanding plant cell walls. Science 306, 2206–11.
Tan X, Calderón-Villalobos LI, Sharon M, Zheng C, Robinson
CV, Estelle M, Zheng N. 2007. Mechanism of auxin perception by
the TIR1 ubiquitin ligase. Nature 446, 640–645.
Traas J, Doonan H. 2001. Cellular basis of shoot apical meristem
development. International Review of Cytology 208, 161–206.
Uyttewaal M, Burian A, Alim K, et al. 2012. Mechanical stress
acts via katanin to amplify differences in growth rate between adjacent
cells in Arabidopsis. Cell 149, 439–451.
Uyttewaal M, Traas J, Hamant O. 2010. Integrating physical stress,
growth, and development. Current Opinion in Plant Biology 13, 46–52.
Vanneste S, Friml J. 2009. Auxin: a trigger for change in plant
development. Cell 136, 1005–1016.
Vernoux T, Besnard F, Traas J. 2010. Auxin at the shoot apical
meristem. Cold Spring Harbor Perspectives in Biology 2, a001487.
Vernoux T, Brunoud G, Farcot E, et al. 2011. The auxin signalling
network translates dynamic input into robust patterning at the shoot
apex. Molecular Systems Biology 7, 508.
Vernoux T, Kronenberger J, Grandjean O, Laufs P, Traas J.
2000. PIN-FORMED 1 regulates cell fate at the periphery of the shoot
apical meristem. Development 127, 5157–5165.
Vieten A, Sauer M, Brewer PB, Friml J. 2007. Molecular and
cellular aspects of auxin-transport-mediated development. Trends in
Plant Science 12, 160–168.
Vieten A, Vanneste S, Wisniewska J, Benková E, Benjamins R,
Beeckman T, Luschnig C, Friml J. 2005. Functional redundancy of
PIN proteins is accompanied by auxin-dependent cross-regulation of
PIN expression. Development 132, 4521–4531.
Wabnik K, Govaerts W, Friml J, Kleine-Vehn J. 2011. Feedback
models for polarized auxin transport: an emerging trend. Molecular
BioSystems 7, 2352–2359.
Wabnik K, Kleine-Vehn J, Balla J, Sauer M, Naramoto S,
Reinöhl V, Merks RMH, Goaverts W, Friml J. 2010. Emergence of
tissue polarization from synergy of intracellular and extracellular auxin
signaling. Molecular Systems Biology 6, 447.
Wisniewska J, Xu J, Seifertová D, Brewer PB, Ruzicka K, Blilou
I, Rouquié D, Benková E, Scheres B, Friml J. 2006. Polar PIN
localization directs auxin flow in plants. Science 312, 883.
Won C, Shen X, Mashiguchi K, Zheng Z, Dai X, Cheng Y,
Kasahara H, Kamiya Y, Chory J, Zhao Y. 2011. Conversion
of tryptophan to indole-3-acetic acid by TRYPTOPHAN
AMINOTRANSFERASES OF ARABIDOPSIS and YUCCAs in
Arabidopsis. Proceedings of the National Academy of Sciences, USA
108, 18518–18523.
Yadav RK, Perales M, Gruel J, Girke T, Jönsson H, Reddy GV.
2011. WUSCHEL protein movement mediates stem cell homeostasis
in the Arabidopsis shoot apex. Genes & Development 25, 2025–2030.
Yamaguchi N, Wu MF, Winter CM, Berns MC, Nole-Wilson
S, Yamaguchi A, Coupland G, Krizek BA, Wagner D. 2013. A
molecular framework for auxin-mediated initiation of flower primordia.
Developmental Cell 24, 271–282.
Downloaded from http://jxb.oxfordjournals.org/ at Smith College on July 9, 2014
Rolland-Lagan AG, Prusinkiewicz P. 2005. Reviewing models
of auxin canalization in the context of leaf vein pattern formation in
Arabidopsis. The Plant Journal 44, 854–865.
Stoma S, Lucas M, Chopard J, Schaedel M, Traas J, Godin
C. 2008. Flux-based transport enhancement as a plausible unifying
mechanism for auxin transport in meristem development. PLoS
Computational Biology 4, e1000207.