Founders Review
Auxin Signaling1[OPEN]
Ottoline Leyser2
Sainsbury Laboratory, University of Cambridge, Cambridge CB2 1LR, United Kingdom
ORCID ID: 0000-0003-2161-3829 (O.L.).
Auxin acts as a general coordinator of plant growth
and development, transferring information over both
long and short ranges. Auxin famously appears to
be extraordinarily multifunctional, with different cells
responding very differently to changes in auxin levels.
There has been considerable progress over recent years
in understanding how this complexity is encoded in the
cellular auxin response machinery. Of central importance
is an elegantly short but versatile signaling pathway
through which auxin triggers changes in gene expression.
However, it is increasingly clear that this pathway is
not sufficient to explain all auxin responses, and other
auxin signaling systems are emerging.
AUXIN BY ANALOGY
Apparently, I have been writing reviews about auxin
for 20 years (Leyser,1997),and certainly I have been in very
good company. Auxin is a much-written-about molecule.
Entire books are dedicated to it (Estelle et al., 2011). From
all these column inches, there is a clear consensus that
auxin is extremely important, that it is involved in virtually
every aspect of plant biology, and that this baffling array of
functions makes the task of understanding auxin daunting
to say the least.
Many very helpful analogies have been developed in
an attempt to provide an intellectual framework within
which auxin biology can be adequately encapsulated. A
major driver for these analogies is to move away from
the constraints of auxin as a specific instructive signal
triggering specific and universal outcomes (Weyers and
Paterson, 2001). This idea is clearly inappropriate as far
as auxin is concerned, but nonetheless somehow inveigles
its way into the field due to prevalent paradigms in
signaling biology in general and hormone biology in
particular. For auxin, it is increasingly clear that the
specificity in the system is not in the signal but in the cells
that perceive it. Auxin does not instruct cells to do anything
in particular, but rather it influences the behavior
of cells according to their preexisting identity. For example,
I have previously suggested that auxin is like the
baton wielded by the conductor of an orchestra: “When
the auxin baton points your way, it’s your turn to play
whatever musical instrument you happen to be holding”
(Leyser, 2005, p. 821).
This analogy is of course deficient in many ways. In
particular, it brings into sharp relief the second and
perhaps more important challenge that must be embraced
to understand auxin. Although auxin can trigger
very specific changes in cells, this seldom involves step
changes in auxin concentration. The question of how
auxin works is not the question of what happens when
some auxin arrives at a cell. There is always auxin. The
absolute and relative amount of auxin at any one location
in the plant varies over time, and this tunes and retunes
the balance within a set of interlocking feedback loops
operating at subcellular, cellular, tissue, organ, and wholeplant
scales. Within the orchestral analogy, maybe some
of this can be captured by considering the dynamics of the
music. The baton is not a play-don’t play switch but can
also be used to modulate how loudly to play. But even
with this addition, the analogy does not allow sufficiently
for feedback, with the baton being modulated by the
players. Maybe if there is an unruly player in the orchestra,
the baton might become a little more insistent, and
certainly an orchestral performance is a collaboration between
the conductor and the players, with information
flow mediated by the baton and its behavior. The same
musical score can be interpreted differently by different
conductor-orchestra combinations. Nonetheless, in this
scenario, there is no doubt who is in charge. It’s the conductor.
In a plant, there is no central control. Rather, all
aspects of auxin biology are characterized by emergent
self-organizing properties that allow distributed rather
than centralized decision making.
These considerations have led to another prevalent
analogy or, rather, a useful comparison (Leyser, 2016).
Many of the functions performed by the auxin network
in plants are fulfilled by the nervous system in animals.
Here, a very nonspecific signal, an electrical impulse,
carries information through the organism modulating
diverse outputs depending on the receiving tissue. There
is extensive feedback, and in the brain in particular, dynamic
self-organization with competition and reinforcement
rewiring neural connections depending on their
frequency of use. However, there is also the obvious and
glaring difference, already mentioned above, of central
versus distributed processing. This difference is often
discussed in the context of the heterotrophic versus
autotrophic life styles of animals versus plants. While
1
O.L.’s research is funded by the Gatsby Charitable Foundation,
the European Research Council, and the UK Biotechnology and Biological
Sciences Research Council.
2
Address correspondence to ol235@cam.ac.uk.
[OPEN]
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heterotrophy is supported by locomotion allowing long
distance foraging for food, autotrophy on land requires
large surface areas for acquisition of dilute resources
such as light, carbon dioxide, and water. This latter involves
large underground surface areas, precluding locomotion.
As a consequence, plants are unable to escape
predation by locomotion and so must be robust to loss of
body parts. Central processing—a brain—is poorly
suited to this requirement and instead plants typically
have no unique parts.
Addressing these issues in their review, Stewart and
Nemhauser argue that auxin is a cellular currency:
“.think of it as money. Auxin does not have much intrinsic
value—it stores very little energy or raw materials.
However, like paper currency, it has great symbolic
value, as an easily circulated means of facilitating transactions
in the dynamic economy of plant life” (Stewart
and Nemhauser, 2010, p. 1). Like auxin, money does everything,
but what it does depends on who gives it to
whom and under what circumstances. And the economics
analogy certainly offers plenty of potential for feedback
and complex self-organizing dynamics.
The money analogy is also interesting from an evolutionary
perspective. Money was invented as a proxy
for the things people really need or want, to allow diverse
goods and services to be more readily exchanged
in more complex and less direct ways than a one-on-one
swap, for example, of eggs for bread. This is potentially
an interesting way to think about auxin. It is metabolically
close to an amino acid, Trp, and amino acid availability
via limitations in nitrogen supply is an important
constraint on plant growth (Elser et al., 2007). Auxin
could plausibly have evolved from a mechanism directly
linking amino acid availability to growth. In the context
of the increasing complexity of multicellular plant form,
with division of labor between different cell and tissue
types, a direct local link between resource availability
and growth does not allow the necessary coordination
and prioritization of growth across the organism. For
example, in most seed plants, shoots capture carbon and
roots capture mineral nutrients such as nitrate. Growth
of the root and shoot systems must therefore be prioritized
depending on the C:N ratio in the plant, rather than
the local availability of either fixed carbon or nitrogen.
The requirement to coordinate growth, not just regulate
it, necessitates dedicated signaling systems that can operate
both systemically and locally. As a general description,
auxin is one such growth coordinator—regulating
where, when, how much, and what sort of growth should
occur. We have previously argued that the apparent
diversity of auxin action makes sense in this context
(Bennett and Leyser, 2014). An early origin as a general
growth coordinator could be followed by cooption to
modulate wider coordinated activities. Once money has
been invented, it can be used and reused to buy things
that did not previously exist. In this review, I will focus on
how auxin as a currency is traded at the cellular level,
or put rather less fancifully, how cells recognize and
use information about changes in the amount of auxin
present.
AUXIN AND THE REGULATION OF TRANSCRIPTION
The major mechanism by which changes in auxin
levels are converted into cellular responses is via changes
in transcription. Hundreds of genes change their expression
rapidly in response to exogenous auxin supply
(Paponov et al., 2008). Auxin regulates transcription via
an elegantly short signal transduction pathway, which
has been extensively reviewed (Chapman and Estelle,
2009; Salehin et al., 2015) and is illustrated in Figure
1. In brief, auxin acts as molecular glue bringing together
F-box proteins of the TRANSPORT INHIBITOR
RESPONSE1/AUXIN SIGNALING F-BOX (TIR1/AFB)
family and members of the Aux/IAA transcriptional
repressor family (Tan et al., 2007). F-box proteins are the
substrate selection subunit of SCF-type ubiquitin protein
ligase complexes, named after three of their subunits:
Skp1, Cullin, and an F-box protein (Smalle and Vierstra,
2004). The F-box itself is a motif at the N-terminal end of
F-box proteins through which they interact with Skp1,
which also interacts with a dimer of Cullin and RBX1.
This dimer transfers activated ubiquitin from a ubiquitin
activating enzyme and conjugates it to target proteins.
The target protein is brought to the SCF by its interaction
with the C-terminal domain of the F-box protein. In the
case of TIR1/AFBs, this consists of Leu-rich repeats including
an auxin binding pocket (Tan et al., 2007). The
binding of auxin in this pocket is greatly stabilized by the
docking of the Aux/IAA protein across the pocket mouth,
mediated by a short protein motif in the Aux/IAA known
as domain II (Tan et al., 2007). For this reason, TIR1/AFBAux/IAA
pairs can be considered as coreceptors for
auxin. The auxin-mediated binding of Aux/IAAs to
TIR1/AFBs brings them to the SCF, allowing their ubiquitination
and subsequent degradation (Gray et al., 2001;
Maraschin et al., 2009). In this way, changes in auxin
levels are converted into changes in Aux/IAA levels.
There are 29 Aux/IAAs in Arabidopsis (Arabidopsis
thaliana; Paponov et al., 2008). Their half-lives and the
extent to which their half-lives reduce in response to
applied auxin vary greatly (Dreher et al., 2006). An important
determinant of these characteristics is the sequence
of domain II. This sequence acts as a degron and is
sufficient to confer auxin-triggered destruction on heterologous
proteins (Zenser et al., 2001). Aux/IAA half-lives
also depend on sequences outside the degron and the
AFB with which the Aux/IAA interacts (Moss et al.,
2015). There are six AFBs in Arabidopsis (Dharmasiri
et al., 2005; Prigge et al., 2016). Different TIR1/AFB-Aux/
IAA pairs have very different affinities for each other
and for different auxins, contributing to the wide range
of Aux/IAA half-lives and their auxin sensitivities
(Calderón Villalobos et al., 2012). Reconstitution of the
auxin response pathway in yeast demonstrates the diversity
in dynamics that can be achieved through this
mechanism, such that the same change in auxin input can
result in a wide range of changes in Aux/IAA output
(Havens et al., 2012). There is evidence that this variation
is functionally significant in planta. For example, in
Arabidopsis the Aux/IAA protein IAA14 is involved in
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the early stages of lateral root development. Stabilized
auxin-insensitive versions of IAA14 are well known to
block lateral root emergence completely. Point mutations
in this protein have been engineered to produce variants,
which in partnership with TIR1 have approximately
10 and 100 times lower affinity for auxin (Guseman et al.,
2015). This increases their half-lives in response to auxin
addition from approximately 20 min to more than 2 h.
When these variants are expressed in plants, they delay
lateral root emergence in proportion to their half-lives
(Guseman et al., 2015).
Aux/IAA proteins act as transcriptional repressors
(Ulmasov et al., 1997). They contain a conserved EAR
domain through which they can recruit corepressors of
the TOPLESS (TPL) family to promoters (Tiwari et al.,
2004; Szemenyei et al., 2008). In turn, TPLs can recruit
chromatin remodeling proteins that stabilize transcriptional
repression (Szemenyei et al., 2008). The Aux/IAAs
do not themselves bind DNA, but they can dimerize
with transcription factors of the AUXIN RESPONSE
FACTOR (ARF) family. Dimerization occurs through
C-terminal PB1 domains shared by both protein families
(Guilfoyle, 2015). The PB1 domain has an acidic and a
basic interaction surface through which dimerization can
occur. This arrangement can support Aux/IAA oligomerization,
like a stack of Lego bricks (Korasick et al., 2014;
Nanao et al., 2014; Dinesh et al., 2015). Point mutations
along one face of IAA19 that allow Aux/IAA dimerization
with ARFs but do not support subsequent oligomerization
with additional Aux/IAAs are nonfunctional in some
assays, suggesting that these higher order complexes can
contribute to effective transcriptional repression (Korasick
et al., 2014). However, transcription can be repressed
without such higher order complex formation (PierreJerome
et al., 2016).
ARF proteins can also homodimerize through their
N-terminal B3 domains, and it is through this homotypic
dimerization that they cooperatively bind DNA
Figure 1. The main pathway for regulation of transcription by auxin. Auxin-inducible genes have AREs in their promoters, which
are bound by dimers of the ARF protein family. Gene expression is prevented by recruitment of Aux/IAA transcriptional repressors
to these promoters via their interaction with the ARFs. Aux/IAAs recruit TPL family corepressors, which in turn recruit chromatin
modifying enzymes (not shown) that stabilize the repressed state. The steps in the auxin response pathway are indicated by the
numbered arrows. 1, Auxin acts as a molecular glue bringing together Aux/IAAs and F-box proteins of the TIR1/AFB family. 2,
These F-box proteins are part of an SCF-type E3 ubiquitin protein ligase complex that transfers activated ubiquitin (Ub) from an E1/E2
enzyme system. 3, Polyubiquitination of the Aux/IAAs results in their degradation. 4, This releases repression at ARE-containing
promoters.
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(Boer et al., 2014). ARFs bind to specific Auxin Response
Elements (AREs) with the consensus sequence
TGTCTC in the promoters of auxin-regulated genes
(Abel and Theologis, 1996; Mironova et al., 2014). A
subset of ARFs include a Q-rich middle domain between
the B3 and PB1 domains, and these ARFs can act
as transcriptional activators through recruitment of
chromatin remodeling enzymes (Ulmasov et al., 1999;
Wu et al., 2015). Oligomerization with Aux/IAAs prevents
this activation. Auxin modulates the level of expression
from ARF-bound promoters by triggering
Aux/IAA degradation.
Since the ability of Aux/IAAs to modulate transcription
is entirely dependent on ARFs, the presence of
different ARF complements in different cells can also
contribute to auxin signaling specificity (Rademacher
et al., 2012; Bargmann et al., 2013). There are 23 different
ARFs in Arabidopsis and evidence for differential affinities
between specific ARFs and Aux/IAAs (Vernoux
et al., 2011). Different homo and hetero oligomerizations
of Aux/IAAs at the promoter may add an additional
mechanism for auxin response diversity (Knox
et al., 2003; Rademacher et al., 2012; Bargmann et al.,
2013). For palindromic AREs, there is also evidence for
different affinities between different dimerized ARFs
and ARE-containing promoters, based on the spacing
of the ARE palindrome (Boer et al., 2014). Interestingly,
many ARFs lack the Q-rich region (Ulmasov et al.,
1999). They bind the same AREs, and in some cases
there is good evidence that they act as transcriptional
repressors (Ulmasov et al., 1999). It is likely that these
repressor ARFs compete with activator ARFs for occupancy
of the same promoters (Vert et al., 2008). The
complement of repressive ARFs in a cell therefore provides
an additional mechanism by which different cells
might respond differently to auxin. It is also possible that
these various alternative protein-DNA and proteinprotein
interactions at the promoters of auxin-regulated
genes are further influenced by protein-protein interactions
off the promoter. For example, off-promoter Aux/
IAA oligomerizations could sequester specific Aux/
IAAs, preventing their interactions with ARFs.
This system allows extremely rapid changes in transcription
in response to auxin. Changes in transcript
abundance can be detected within 3 to 5 min of auxin
treatments (McClure et al., 1989; Abel and Theologis,
1996). The half-life of many Aux/IAAs is very short
even in the absence of exogenously applied auxin (Abel
et al., 1994), and the genes encoding Aux/IAA proteins
are themselves rapidly up-regulated in response to
auxin (Abel and Theologis, 1996). Thus, cells continuously
make and degrade Aux/IAAs with the flux of
Aux/IAAs through this cycle modulated by auxin, potentially
reequilibrating the system at different steady
states depending on the auxin concentration. Consistent
with this behavior, auxin signaling is particularly sensitive
to mutation in apparently generic parts of the protein
degradation and protein synthesis machinery (del
Pozo et al., 2002; Gray et al., 2003; Hellmann et al., 2003;
Chuang et al., 2004; Stirnberg et al., 2012). This powerfully
illustrates the mismatch between auxin signaling and
the classical on-off switch paradigm. Auxin modulates
transcription through retuning the equilibrium in a
highly dynamic, feedback-regulated network. The
system is capable of acting as a bistable switch, but it
has a much wider range of possible behaviors, and
even stable high and low expression states are dependent
on flux through the Aux/IAA synthesis-degradation
cycle (Bridge et al., 2012).
Another important feature of this system is that auxin
acts by degrading a non-DNA binding inhibitor of
transcription. From an evolutionary perspective, this
immediately suggests the hypothesis that the ability of
auxin to regulate transcription could have been added
on to an existing system of transcriptional regulation. A
completely auxin-independent system, consisting only
of activator and repressor ARFs competing for the same
promoters with some mechanism for changing their
relative abundance, could provide a way to modulate
transcription from multiple genes, for example, tuning
growth in response to environmental factors. Adding in
Aux/IAA-mediated transcriptional repression and
auxin-triggered Aux/IAA degradation would make these
genes auxin responsive in the presence of activator ARFs.
Some evidence to support this order of events in the evolution
of auxin signaling comes from recent work on
basal land plant species: the liverwort Marchantia polymorpha
and the moss Physcomitrella patens.
The Marchantia genome has only three ARFs, one
Aux/IAA, and one TIR1/AFB. Evidence to date supports
the idea that this system works in the same way as
in angiosperms, described above (Flores-Sandoval et al.,
2015; Kato et al., 2015). The ARFs fall into three distinct
clades covering both activator and repressor ARFs, all
three of which are represented in angiosperm genomes
(Kato et al., 2015). This three-clade structure is therefore
the likely ancestral state for land plants. Analysis of gainof-function
and reduced function mutants in these
components has identified diverse auxin-regulated aspects
of growth and development such as cell expansion
in dorsal epidermal tissues of the Marchantia thallus
(Flores-Sandoval et al., 2015; Kato et al., 2015).
In P. patens, there are 16 ARFs, distributed across the
three clades described above, and three Aux/IAAs
(Prigge et al., 2010). Recently, a Physcomitrella line completely
lacking all these Aux/IAAs has been generated
(Lavy et al., 2016). These plants lack any detectable transcriptional
response to auxin and phenotypically resemble
moss plants treated with high levels of auxin. The
analysis of this line arguably provides some support for
the idea that auxin signaling evolved as a refinement of a
system based on environmental control of the ratio of
repressive and activating ARFs. First, overexpression of
a repressive ARF in the triple Aux/IAA mutant background
suppresses its constitutive high auxin phenotype,
and indeed confers a phenotype similar to that conditioned
by expression of stabilized auxin-resistant Aux/
IAA protein variants (Lavy et al., 2016). This suggests that
repressive and activating ARFs do indeed compete for
access to the same promoters, and shifting the ratios
468 Plant Physiol. Vol. 176, 2018
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between them can coordinately regulate transcription
from these promoters.
Second, examination of the auxin-Aux/IAA-ARFregulated
transcriptome in moss shows that the gene
set down-regulated by auxin and loss of Aux/IAAs is
enriched for genes involved in light responses, photosynthesis,
and carbon fixation, while the up-regulated
gene set is enriched for genes involved in transcription
and biosynthesis (Lavy et al., 2016). This correlates with
the ability of auxin to suppress the production of the
chloroplast-rich chloronemal moss filaments and promote
the production of relatively chloroplast-poor caulonemal
filaments, which extend the moss colony. Auxin
can therefore be seen as switching the moss colony from
prioritizing energy and carbon capture to prioritizing
growth and expansion. This might be associated with
differences in the C:N ratio, and could ancestrally have
been driven, for example, by light regulation of ARFs,
perhaps modulated in some way by nitrogen availability.
Consistent with this speculative idea, like auxin, high
light prioritizes caulonemal over chloronemal growth
(Thelander et al., 2005). However, unlike light, auxin is
potentially mobile in the plant and hence can coordinate
growth systemically. As described above, an early origin
as a general growth coordinator could be followed by
cooption to modulate and coordinate a wider range of
activities across the plant body, consistent with the lethality
often observed in higher plant auxin signaling
mutants, for example, lacking multiple members of the
TIR1/AFB family (Dharmasiri et al., 2005).
In this context, the recent discovery of an additional
ARF-dependent auxin-sensing mechanism is of particular
interest. ETTIN is a noncanonical ARF that lacks the PB1
Aux/IAA interaction domain. Its best characterized role is
in regulating patterning in the developing gynoecium
(Sessions et al., 1997; Nemhauser et al., 2000). Here, ETTIN
dimerizes with the basic helix-loop-helix transcription
factor INDEHISCENT (IND) to regulate transcription of
key target genes (Simonini et al., 2016). This dimerization
is affected by auxin binding, and this correlates with
auxin-dependent changes in the association of ETTIN
with the promoters of target genes and auxin-dependent
changes in the expression of these targets (Simonini et al.,
2016). This ability of auxin to modulate the interaction
between ETTIN and partner transcription factors extends
beyond IND, and indeed could be quite widespread
(Simonini et al., 2016). It will be interesting to explore the
evolutionary origin of this mechanism for auxin-regulated
transcription based on a noncanonical ARF.
IS THAT ALL THERE IS?
The TIR1/AFB-Aux/IAA-ARF signaling system provides
plenty of scope for diversity in auxin response, and
the dramatic phenotypes of mutants compromised in
this pathway attest to its importance in the coordination
of plant growth and development.
This begs the question as to whether this is the only
auxin detection system in plants. Several arguments
have been put forward in support of the existence of
additional auxin response systems. However, many of
these can easily be accommodated by the TIR1/AFBAux/IAA-ARF
system. One argument that there are
multiple systems comes from the very different auxin
sensitivities of different auxin responses, both with respect
to their dose-response kinetics and their specificity
for different auxins. For example, a much-studied
classical response to auxin is its ability to drive cell
elongation in hypocotyls, epicotyls, and coleoptiles, all
of which must elongate rapidly during early seedling
establishment after seed germination. This response is
complex. In pea (Pisum sativum), the natural auxin indole-
3-acetic acid (IAA) has a biphasic dose response curve
with maxima in the 1 mM and 1 mM ranges (Yamagami
et al., 2004). Meanwhile, the dose response curve for the
synthetic auxin 1-naphthaleneacetic acid (NAA) has only
one maximum in the 10 mM range (Yamagami et al.,
2004). Of these response maxima, only the high-affinity
IAA response is strongly sensitive to Ca2+
availability,
while the others are not (Yamagami et al., 2004). This
kind of diversity has been used to argue for two distinct
perception mechanisms with different affinities for IAA
and NAA. However, the diversity in auxin sensitivity of
TIR1/AFB-Aux/IAA pairs could account for these phenomena
given the wide range of Kms described for different
auxins and different AFB-Aux/IAA pairs (Calderón
Villalobos et al., 2012). The basis for the differences in Ca2+
sensitivity is less straightforward to accommodate through
the AFB-Aux/IAA system.
While the downstream effectors driving these elongation
responses are a matter for debate, there is substantial
evidence supporting a role for various ion fluxes
across the plasma membrane (Rück et al., 1993). Because
the activity of diverse membrane transporters is regulated
posttranscriptionally, this is a second argument
that has fueled speculation about additional response
pathways and, specifically, nontranscriptional auxin
effects. For example, auxin-induced elongation is associated
with stimulation of the plasma membrane protonpumping
ATPase (PM H+
ATPase), thereby acidifying
the apoplast (Hager, 2003; Takahashi et al., 2012). According
to the acid growth hypothesis, this in turn affects cell
wall extensibility leading to turgor-driven elongation
(Kutschera, 1994). This response is quite slow and so
could reasonably result from the very rapid changes in
gene expression triggered by auxin (Badescu and Napier,
2006). Indeed, recent data support this idea. In
particular, there is mounting evidence that auxin stimulates
the activity of the PM H+
ATPase by up-regulating
the transcription of members of the SMALL AUXIN
UP-REGULATED RNA (SAUR) gene family via the canonical
TIR1/AFB-Aux/IAA-ARF pathway (Chae et al.,
2012; Spartz et al., 2012).
As their name suggests, the SAUR genes were originally
identified because many of them are rapidly
up-regulated in response to auxin (Abel and Theologis,
1996). The promoters of the auxin-up-regulated SAUR
family members include the classical ARE ARF binding
motif, and changes in SAUR transcript abundance can
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be detected within 5 min of auxin application (McClure
et al., 1989; Abel and Theologis, 1996). SAUR genes are
plant-specific and typically present as multigene families,
making genetic analysis difficult. However, the use
of artificial microRNAs to target multiple SAURs simultaneously,
together with gain-of-function studies,
has demonstrated that in Arabidopsis the SAUR19SAUR24
subfamily are positive regulators of cell expansion
in diverse tissues, including the hypocotyl
(Spartz et al., 2012).
This work has been greatly helped by the discovery
that fusion proteins between these SAURs and a wide
range of tags result in the stabilization of the proteins,
providing overexpression lines (Spartz et al., 2012).
Arabidopsis lines overexpressing tagged SAUR19 have
long hypocotyls due to increased cell expansion. This
phenotype is associated with reduced apoplastic pH,
which is significantly attenuated in PM H+
ATPase
mutant backgrounds (Spartz et al., 2014). Consistent
with these results, the SAUR19 overaccumulating lines
show constitutive activity of the PM H+
ATPase, associated
with increased phosphorylation of Thr- 947 within
the pump’s C-terminal autoinhibitory domain. Phosphorylation
at this site is known to activate the PM
H+
ATPase by driving recruitment of a 14-3-3 protein,
which binds to the domain, alleviating its inhibitory influence
(Fuglsang et al., 1999; Kanczewska et al., 2005).
Fluorescent tags demonstrate that SAUR19 can localize
to the plasma membrane (Spartz et al., 2012). Furthermore,
SAUR19 and other SAUR family members interact
directly with type 2C protein phosphatase D (PP2C-D),
inhibiting its activity, and this PP2C-D interacts with and
inhibits the activity of the PM H+
ATPase (Spartz et al.,
2014; Sun et al., 2016). These results support a clear chain
of events through which auxin regulates PM H+
ATPase
activity via transcriptional up-regulation of SAUR19,
which directly inhibits PP2C-D activity, leading to the
accumulation of active Thr-947 phosphorylated PM
H+
ATPase, apoplast acidification, cell wall loosening,
and growth. In support of this model, Arabidopsis
SAUR19 and SAUR9 can inhibit tomato PP2C-Ds, and
overexpression of Arabidopsis SAUR19 in tomato bypasses
the requirement for auxin addition to trigger increased
cell wall extensibility and elongation in excised hypocotyl
segments (Spartz et al., 2017). Consistent with these
results, acid growth in Arabidopsis hypocotyls is dependent
on the TIR1/AFB-Aux/IAA-ARF signaling system
(Fendrych et al., 2016).
These results provide direct evidence that the TIR1/
AFB-Aux/IAA-ARF transcriptional pathway for auxin
response regulates PM H+
ATPase activity and that this
contributes to the ability of auxin to promote elongation.
As mentioned above, the response times for acidification
and growth in these tissues are in the order of 10 to
30 min, concordant with this idea. However, in other
situations much more rapid changes in ion fluxes and
growth are observed.
A particularly convincing example comes from
high spatiotemporal resolution analyses of root responses
to auxin. Addition of physiologically relevant
concentrations of auxin to Arabidopsis roots results
in a dose-dependent influx of Ca2+
across the plasma
membrane, increasing cytosolic Ca2+
concentrations
(Monshausen et al., 2011). This response occurs within
10 s and is accompanied by apoplastic alkalinization,
which can be measured as a change in root surface pH.
Pretreatment with the Ca2+
channel blocker La3+
inhibits
both the Ca2+
and pH responses, suggesting that
the pH changes are dependent on the increase in cytosolic
Ca2+
.
Such rapid changes in ion fluxes have been detected
previously in guard cells and particularly in protoplast
systems where their physiological significance has
remained unclear (Rück et al., 1993; Blatt and Thiel, 1994).
The high level of spatiotemporal resolution now possible
in growing roots, coupled with the power of Arabidopsis
genetics, has provided robust evidence linking these
fluxes to growth. Both the rapid auxin-induced cytosolic
Ca2+
elevation and the apoplastic alkalinization have
been shown to depend on a cyclic nucleotide-gated
channel, CNGC14 (Shih et al., 2015). Both responses are
completely abolished in cngc14 loss-of-function mutants
(Shih et al., 2015). Members of this family are known to
transport Ca2+
, and although this protein accumulates
only to very low levels in root tips, it can be detected at
the plasma membrane in at least some cell types (Shih
et al., 2015). Crucially, auxin-induced root growth inhibition
is significantly delayed in cngc14 roots (Shih et al.,
2015). In wild-type roots, a significant reduction in elongation
rate across the elongation zone can be detected
within 1 min of auxin addition. In cngc14 mutants, it is
not until 7 min after auxin application that the rate of root
growth slows significantly. These data suggest that auxin
stimulates the activity of CNGC14 triggering Ca2+
influx,
which in turn results in cell wall alkalinization, inhibiting
cell elongation.
This idea is supported by analysis of the root gravity
response, bypassing the possibility of any artifacts associated
with the use of applied auxin (Shih et al., 2015).
The ability of roots to reorient their growth in response
to changes in the gravity vector is dependent on the
asymmetric redistribution of auxin at the root tip.
Auxin transported toward the tip in the central stele is
recirculated back up the root through the lateral root
cap and root epidermis (Luschnig et al., 1998; Müller
et al., 1998; Swarup et al., 2005). This pattern of auxin
transport is mediated by polarly localized transporters
of the PIN-FORMED (PIN) family (Blilou et al., 2005). In
the columella root cap, auxin arriving through central
tissues is transported laterally by PIN3, where it enters
the peripheral shootward transport system. Tight control
over the flow of auxin through these tissues is
achieved by rapid removal from the apoplast mediated
by the AUX/LAX family of auxin importers (Swarup
et al., 2005). Upon gravistimulation detected in the
columella root cap, PIN3 is polarized in the root cap
cells, directing auxin disproportionately to the lower
root surface, resulting in asymmetric distribution of
auxin across the root and consequent asymmetric
growth (Friml et al., 2002; Harrison and Masson, 2008).
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Current methods for live imaging of auxin concentrations
are not sufficiently sensitive to detect directly
this auxin asymmetry rapidly after gravistimulation
(Band et al., 2012). The best available method at present
is based on the auxin-triggered degradation of a fluorescent
reporter protein fused to a relatively long halflife
Aux/IAA degron (Vernoux et al., 2011). A long
half-life degron is required to allow sufficient accumulation
of the fluorescent protein for detection, but comes
at the expense of temporal resolution. Using a parameterized
model for degradation of this protein in response
to auxin, it has been estimated that asymmetry is
established at 5 min after gravistimulation (Band et al.,
2012). This is broadly consistent with the asymmetric
distribution of Ca2+
, which was detected within 3 to
6 min of gravistimulation, with higher levels on the
lower surface of the root than the upper surface (Shih
et al., 2015). Similarly, when auxin is applied locally at
the root tip, a wave of elevated cytosolic Ca2+
is detected,
moving shootward at a rate 150 to 600 mm/min, consistent
with typical rates of polar auxin transport (Shih
et al., 2015). That this wave is auxin-mediated is supported
by the fact that it is dependent on the auxin influx
carrier AUX1. In the case of gravistimulation, the pH at
the upper surface reduces and at the lower surface increases
with similar kinetics. These rapid changes in ion
flux correlate with differential growth, which can be
detected after 4 min. In the cgnc14 mutant, all three of
these effects are delayed by approximately 6 min (Shih
et al., 2015).
These data provide compelling evidence for a mode
of auxin action too rapid for the transcriptional pathway.
A 10-s response timeframe does not seem plausible
for even the extremely rapid changes in gene
expression elicited by auxin. Consistent with this idea,
the ion flux changes are unaffected in a mutant lacking
three of the TIR1/AFB auxin receptors (Shih et al.,
2015), which is severely compromised in the transcriptional
response system (Dharmasiri et al., 2005). In
this context it is important to note that gravitropic root
growth is only very mildly affected in cgnc14 mutant
roots but is strongly compromised in mutants defective
in the transcriptional response system. This suggests
that, while the CNG14-dependent response contributes
to the very rapid initiation of root reorientation, the
transcriptional system is required for longer timeframe,
sustained reorientation responses.
The nontranscriptional detection of auxin extends
further the timeframes over which cells can directly
sense changes in auxin concentrations from seconds to
many hours. The importance of these diverse response
times is beautifully illustrated at the root tip. Here, there
is good evidence that a tip-high gradient of auxin patterns
root development (Sabatini et al., 1999; Galinha
et al.,2007;Mähönen et al.,2014),but also fine-tunes growth
rates, as in the example of gravitropism (Luschnig et al.,
1998; Müller et al., 1998; Swarup et al., 2005). A cluster
of initial cells form a stem cell niche at the root tip,
where the auxin concentration is high (Sabatini et al.,
1999). These cells give rise to a rapidly dividing population
of cells that make up the division zone. On exiting the
cell cycle, the cells expand rapidly, forming an elongation
zone, after which they mature and differentiate, for
example, as root hairs in the differentiation zone. There
is good evidence that the rate of progress of cells
through these stages is dependent on the tip-high auxin
concentration gradient mentioned above but interpreted
indirectly through the auxin-regulated expression
of a set of transcription factors of the PLETHORA
family (Galinha et al., 2007; Mähönen et al., 2014).
These proteins are expressed in response to the high
auxin concentrations of the root tip. They are very
stable, persisting through cell divisions. As a result, a
stable gradient of PLETHORA forms, buffered against
rapid changes in auxin concentration, for example,
due to gravistimulated auxin redistribution at the root
tip (Mähönen et al., 2014). Meanwhile, as described
above, these transient fluctuations in auxin level can
be interpreted over very short timescales by calcium
spiking, and over intermediate timescales by rapid
primary transcriptional changes, supporting dynamic
responses in root growth rate and direction. Thus,
multiple direct and indirect auxin-sensing systems
operating over different timescales deliver robust
multiscale growth coordination.
A POLARIZING ISSUE
The existence of a nontranscriptional auxin signaling
system is also supported by several theoretical considerations.
Transcriptional regulation has only a limited
ability to account for the apparent importance of auxin
in polarity and polarization. There is a considerable
body of evidence that auxin is intimately involved in
the regulation of polarity. The mechanisms by which
auxin contributes to cellular and tissue level polarity are
poorly understood, and consequently they are currently
a very active area of research. In some cases, it
seems likely that auxin acts to allow proteins to respond
to polarizing cues provided by other systems. For example,
auxin, acting through its transcriptional pathway,
influences polar PIN accumulation and activity
(Hazak et al., 2010). However, there is also evidence
that auxin can contribute to the polarizing cue itself.
Here again, in some cases, the canonical TIR1/AFBAux/IAA-ARF
pathway can account for auxin-mediated
polarization phenomena. For example, organ primordia
at the shoot apical meristem are formed at sites of locally
high auxin (Reinhardt et al., 2003; Heisler et al., 2005).
Auxin accumulates at these maxima through the action of
specific members of the PIN family of auxin efflux carrier.
In the meristem epidermis, these PINs are polarized toward
the neighboring cell inferred from auxin response
reporters to have the highest auxin concentration, in a
so-called “up the gradient” pattern (Reinhardt et al.,
2003; Heisler et al., 2005; Jönsson et al., 2006; Smith et al.,
2006). One mechanism that has been proposed to account
for this polarization is based on the ability of cells to
detect differences in physical forces across the cell wall
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(Heisler et al., 2010). If auxin promotes cell expansion in
the meristem epidermis, then the increased expansion of
cells with higher auxin levels than their neighbors will
create tensile stresses in the cell wall adjacent to the
expanding cell. If PIN proteins are delivered or retained
in the plasma membrane in proportion to adjacent cell
wall stresses, then this could drive up-the-gradient PIN
polarization, as observed. In this way, nuclear auxin
signaling, working via auxin-regulated gene expression,
could regulate polarization of neighboring cells.
There is now strong evidence to suggest that this
noncell-autonomous polarization depends on the TIR1/
AFB-Aux/IAA-ARF system (Bhatia et al., 2016). The ARF
MONOPTEROS (MP) is a central player in patterning
organ emergence at the shoot apical meristem. Meristems
deficient in MP are barren, producing no organs, nor can
they be induced to produce organs by local auxin application
(Reinhardt et al., 2000). When small clones of wildtype
MP were induced in such an mp mutant background,
PIN1 polarized toward the clones and organ-like outgrowths
were initiated at these sites (Bhatia et al., 2016).
This strongly suggests auxin signaling via the TIR1/AFBAux/IAA-ARF
system can act as a polarizing cue for
neighboring cells in the shoot apical meristem. This could
act via the wall stress mechanism described above, but
any directional signal from the MP-expressing cells could
contribute to orienting PINs in neighboring cells.
However, there are other examples of auxin-regulated
cell polarization that are more difficult to explain using
only nuclear auxin signaling. Cellular polarization fundamentally
requires symmetry breaking, and within a
cell it is difficult to see how this could be accomplished
by changes in transcription alone. During organ formation
at the shoot apical meristem, the role of auxin is
noncell autonomous, with asymmetry established in the
polarizing cell by auxin signaling in its neighbors. There
are situations where this type of explanation is more
difficult to reconcile with the observed phenomena.
A good example is the positioning of Arabidopsis root
hairs. As described above, various assays suggest that
there is a tip-high gradient of auxin that patterns the
Arabidopsis root, regulating the transition of cells between
the division, elongation, and differentiation
zones. This gradient also appears to play a role in positioning
root hairs. Root epidermal cells elongate highly
anisotropically in the elongation zone. In trichoblast cell
files, upon exit from the elongation zone, root hairs develop
as tip-growing projections from the epidermal
cells (Nakamura et al., 2012). The hairs are positioned
close to, but a little way back from, the rootward end of
the trichoblast. The site of root hair development is
predicted by accumulation of a patch of the type I Rho of
Plants (ROP) GTPase protein on the plasma membrane
(Molendijk et al., 2001; Jones et al., 2002). This patch acts
as an organizing center for the cytoskeletal changes associated
with root hair development.
The positioning of the ROP patch and therefore the
root hair along the trichoblast can be shifted by manipulation
of the auxin gradient along the root (Fig. 2;
Fischer et al., 2006; Ikeda et al., 2009). For example, when
this gradient is flattened as in the aux1 ethylene-insensitive2
(ein2) gnomeb (gneb) triple mutant, root hairs emerge at
variable positions along the trichoblast cells. Although
the hairs can emerge at any position, there is a bias toward
either the rootward or the shootward end of the
cell. Correct, rootward positioning can be restored by
application of auxin rootward of the differentiation zone
(Fischer et al., 2006). Strikingly, root hair positioning can
be biased to the shootward end by application of auxin
shootward of the differentiating hair (Fig. 2). These data
support the hypothesis that an auxin gradient contributes
to the positioning of the root hair, and consistent
with this idea, mutants with defects in root tip auxin
biosynthesis have a shootward shift in root hair positioning
(Ikeda et al., 2009).
Manipulation of ROP activity can also affect the position
of root hair emergence. For example, overexpression
of ROPs, or expression of a constitutively active form, can
trigger the development of multiple root hairs per cell
(Jones et al., 2002). Combined, this phenomenology, and
particularly the wide range of root hair positions observed
in response to auxin and ROP manipulation, is
difficult to explain with a system in which polarization is
driven noncell autonomously from neighboring cells.
An attractive hypothesis has been proposed to explain
these data. At its core is the ability of a gradient to
stabilize a self-organizing Turing pattern. Turing patterns
emerge from a regulatory architecture involving a
slow diffusing activator that promotes its own activity,
combined with a faster diffusing inhibitor of this activity
(Turing, 1952). Type I ROPs could act in a Turing
system since upon activation they can be S-acylated and
recruited into lipid rafts (Sorek et al., 2007). Thus, in
their inactive form they may diffuse much more rapidly
than in their active state. With the assumption of autocatalytic
activation, this can create patches of active
ROPs on the membrane (Payne and Grierson, 2009).
However, to position the patch at a specific site along
the cell, additional information is needed, and this is
hypothesized to derive from the auxin gradient. If local
auxin concentration tunes, for example, the rate of autocatalytic
activation of ROPs, then the site(s) of ROP
nucleation can depend on local auxin concentration
(Payne and Grierson, 2009). Consistent with this idea,
there is some evidence that auxin can rapidly activate
ROPs in a dose-dependent manner (Tao et al., 2002; Xu
et al., 2010). This can be detected using antibodies specific
to the active form of ROPs. Active ROPs are known
to regulate cytoskeletal organization, for example, via
RIC1 and RIC4 (Yalovsky et al., 2008), which could coordinate
the events necessary for root hair emergence.
This model is compelling because of its impressive
ability to generate the patterns of root hair emergence
observed as a result of various genetic and pharmacological
manipulations by mechanistically plausible model
parameter changes. The idea that symmetry breaking to
generate a root hair relies on a self-organizing Turing-like
patterning system for ROPs has an evidence base, particularly
with respect to polarization events in slime
mold and animal cells (Jilkine et al., 2007; Otsuji et al.,
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2007), as well as from more general considerations of the
requirements for self-organizing polarity. The role of
auxin is simply to bias this system, effectively providing
a quantitative positional cue. In particular, the model
involves sensing an intracellular, tip-high auxin gradient
in each trichoblast. Such a gradient is predicted to be fed
by the macroscopic tip-high auxin gradient but patterned
substantially by the polar shootward localization
of PIN2 efflux carriers in each trichoblast cell (Grieneisen
et al., 2007). The active efflux of auxin from the shootward
end of these cells is predicted to deplete the adjacent
cytoplasm of auxin, establishing a diffusion-limited
auxin gradient along the cell. Under this hypothesis, the
aux1 ein2 gneb triple mutant could reduce the steepness of
the intracellular gradient by reducing cellular auxin
supply and uptake (Fig. 2). Unfortunately, it is not currently
possible to measure intracellular gradients, so
these ideas cannot be directly tested.
If root hair position is indeed determined by this
mechanism, auxin’s role is effectively to convert an inherent
apical-basal polarity axis, reflected in PIN2 polarity,
into a new lateral growth axis for root hair emergence.
However, intracellular auxin gradients have also been
proposed as a mechanism to regulate polar PIN protein
distribution itself, raising the possibility of feedback between
PIN-generated auxin gradients, and PIN polarity.
This idea is particularly interesting in the context of auxin
transport canalization. Auxin transport canalization is a
hypothesis proposed by Tsvi Sachs to account for a variety
of observations associated with the regulation of
vascular strand patterning by auxin (Sachs, 1968, 1969,
1981). Vascular strands develop between auxin sources,
such as young expanding leaves, and auxin sinks, such as
existing vascular strands, which include files of cells with
highly polar rootward auxin transport. Vascular strand
development is preceded by the emergence of files of cells
expressing highly polar PIN transporters connecting the
auxin source to the sink (Sauer et al., 2006; Scarpella et al.,
2006; Sawchuk et al., 2013). This can be explained if an
initial passive flux of auxin between the source and the
sink is up-regulated and polarized in the direction of the
sink by the accumulation of PIN proteins at the plasma
membrane in proportion to net auxin flux across the
membrane (Mitchison, 1980; Sachs, 1981; Rolland-Lagan
and Prusinkiewicz, 2005). Although this idea can account
for a wide range of observations, the mechanism by
which PIN proteins can be allocated in proportion to flux
(so-called “with the flux” polarization) is entirely obscure
(Bennett et al., 2014). One mechanism that has been proposed
to account for this is the allocation of PIN proteins
to membranes dependent on intracellular auxin gradients
(Kramer, 2009). Modeling has demonstrated that if cells
with an intracellular auxin gradient greater than 1% allocate
PINs to the plasma membrane adjacent to the
lowest cytoplasmic auxin concentration, this can contribute
to the positive feedback necessary to drive canalization
of auxin transport between an auxin source and a
sink. Integral to this feedback is the amplification of the
intracellular gradient by PIN polarization.
Alternative or additional explanations for fluxcorrelated
PIN allocation involve positive feedback
between PIN accumulation and PIN activity, modified
by extracellular auxin concentration (Cieslak et al.,
2015). This requires an extracellular auxin sensor of
some kind, as well as the ability of PIN proteins to act,
in effect, as auxin sensors, for example, if PIN activity
inhibits their removal from the plasma membrane.
Interestingly, this kind of dual-sensing system can
generate either with-the-flux type patterns of PIN accumulation
or up-the-gradient type patterns depending on
whether extracellular auxin decreases or increases local PIN
accumulation (Cieslak et al., 2015).
An additional mechanism has been reported that can
generate either up-the-gradient or with-the-flux type
Figure 2. Hypothetical intracellular auxin gradients
and root hair position. A, Epidermal cells in
trichoblast cell files of the Arabidopsis root produce
a root hair toward the rootward end of the
cell. This is patterned by a root-tip-high auxin
gradient, which is predicted to feed a tip-high
intracellular auxin gradient in trichoblasts (blue
shading), reinforced by shootward localization of
the PIN2 auxin exporter (orange line). B, Triple
mutation in aux1 ein2 gneb flattens the tip-high
auxin gradient and presumably also the intracellular
auxin gradient. This is associated with shifts
in root hair placement. C, Application of auxin to
aux1 ein2 gneb mutants shootward of the root hair
differentiation zone shifts the root hair position to
the shootward end of the cell, presumably associated
with an inverted intracellular auxin gradient.
D, Application of auxin to aux1 ein2 gneb
mutants rootward of the root hair differentiation
zone restores a more wild-type root hair position,
presumably associated with restoration of the
tip-high intracellular auxin gradient.
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patterns of PIN polarization. This involves an auxinbiased
spontaneous polarization mechanism similar to
that described above for ROPs but operating at the
whole-cell level to create a single axis of polarity across
the cell rather than a small patch (Abley et al., 2013). If
this axis is oriented by extracellular auxin, even a very
shallow auxin gradient can create coordinated cell polarity
across a tissue oriented from high extracellular
auxin, as might be expected at an auxin source, to low
extracellular auxin, as might be expected at an auxin
sink. Since polarization depends on extracellular auxin,
orientation of PINs toward cells with high intracellular
auxin, as in up-the-gradient polarization patterns, can
be achieved if these cells express high levels of auxin
importers and therefore have low extracellular auxin. This
is consistent with the observation that up-the-gradient
PIN polarization is correlated with, and in some cases
dependent on, expression of auxin importers of the Aux/
LAX family (Abley et al., 2016). Unfortunately, as for
intracellular auxin gradients, there is currently no way
to measure extracellular auxin concentration at the
level of resolution necessary to assess correlations with
PIN accumulation.
Together, these considerations suggest that additional
auxin-sensing mechanisms beyond transcription
are necessary to explain the full range of auxin activities.
These could include an intracellular sensor that can
detect intracellular gradients and/or an extracellular
auxin sensor. In both cases, these could act to bias selforganizing
polarization systems, such as ROP partitioning.
There is some evidence that auxin can regulate
ROP activity and thus ROP partitioning (Tao et al.,
2002; Xu et al., 2010). The detection systems involved
are poorly understood but a small family of plasma
membrane-localized receptor kinases have been implicated
(Xu et al., 2014). Similarly, the relationship between
these hypothetical auxin detection systems and
the rapid auxin-induced calcium transients described
above is also unclear.
OTHER AUXIN RECEPTORS
Given the evidence for auxin signaling beyond
the TIR1/AFB-Aux/IAA-ARF system, including nontranscriptional
effects, there should be additional auxin
receptors/sensors. Indeed, several have been proposed,
with varying degrees of support and varying levels of
evidence for their functional significance. Prominent
among them is Auxin Binding Protein1 (ABP1), which
has been a constant and controversial feature on the
auxin signaling landscape throughout the full 20 years I
have been writing reviews about auxin. As its name
suggests, ABP1 was originally identified biochemically
through its ability to bind auxin (Löbler and Klämbt,
1985a, 1985b). A substantial body of data on the biochemistry
of ABP1 has accumulated since, including its
crystal structure (Woo et al., 2002). ABP1 binds the natural
auxin IAA with a Kd of 5 to 10 mM (Napier, 1995). It has a
much higher affinity, in the region of 100 nM, for the
synthetic auxin NAA. Binding of NAA to ABP1 is highly
pH sensitive, with an optimum of 5.0 to 5.5, with very little
binding at pH 7. The cell biology of ABP1 has also been
studied quite intensively because the majority of the protein
is retained in the endoplasmic reticulum (ER), where
the pH is such that auxin binding is predicted to be weak
(Napier, 1997; Klode et al., 2011). Although there is considerable
speculation about the role of ER-localized auxin
in nuclear and cytoplasmic auxin homeostasis (Friml and
Jones, 2010), there is currently no evidence that auxin signals
from the ER. However, some ABP1 appears to escape
ER retention and is secreted, where the pH matches better
the auxin binding maximum (Jones and Herman, 1993;
Napier, 1997).
The analysis of protoplast auxin responses provides
some evidence that ABP1 might function as an extracellular
auxin receptor (Rück et al., 1993; Steffens et al.,
2001; Badescu and Napier, 2006). For example, protoplasts
derived from pea epicotyl epidermal cells swell
slightly in response to IAA addition. Measured 90 min
after auxin treatment, this response is biphasic with
maxima at around 1 and 10 mM. The dose response
curve for NAA has only a single maximum at around
1 mM. When these assays are performed in the presence
of antibodies against ABP1, the response to NAA is
completely abolished, and the response to IAA is simplified,
with a single peak retained at 1 mM (Yamagami
et al., 2004; Badescu and Napier, 2006). One interpretation
of these results is that auxin can induce protoplast
swelling by two different mechanisms, one of
which is ABP1 dependent.
However, while these responses are intriguing, their
physiological significance is unclear. Robust in planta
analysis requires genetic resources, and it is here that the
ABP1 story becomes most problematic. Various methods
have been used to assess the effects of modulating ABP1
activity in planta. These include conventional antisense
expression, as well as overexpression of both the native
protein and a mutant version lacking the ER retention
sequence (Jones et al., 1998; Braun et al., 2008; Tromas
et al., 2009; Robert et al., 2010). In addition, lines overexpressing
the anti-ABP1 antibody have also been
widely used, again either ER retained or secreted (Venis
et al., 1992; Leblanc et al., 1999; Braun et al., 2008; Tromas
et al., 2009; Robert et al., 2010). These approaches come
with caveats because, for example, antibodies might
have off-target effects, and overexpression is prone to
neomorphism artifacts. The use of these lines has been a
mainstay of ABP1 research because of initial suggestions
that its stable loss of function resulted in early embryonic
lethality, precluding the use of clean null mutants for
analyzing the postembryonic roles of ABP1 (Chen et al.,
2001). To circumvent this problem, TILLING was used to
isolate weaker alleles, including the abp1-5 allele, which
carries a point mutation in the auxin binding pocket
(Robert et al., 2010; Xu et al., 2010). This was predicted to
prevent auxin binding without markedly altering other
ABP1 properties.
A number of auxin response defects have been
reported for these ABP1-perturbed lines. Most have
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focused on cellular-level responses, such as inhibition
of endocytosis (Robert et al., 2010; Chen et al., 2012) and
microtubule reorientation (Chen et al., 2014). These
phenotypes have been linked to ROP activity, which as
described above may be auxin regulated. ROPs, auxin,
and ABP1 have also all been implicated in pavement
cell morphogenesis in the leaf (Xu et al., 2010).
However, the interpretation of these results has been
called into question by the identification of new null
mutant alleles in the ABP1 gene (Gao et al., 2015). These
mutations have no obvious phenotypic differences
compared with wild-type plants. Meanwhile, the lethality
attributed to loss of ABP1 in the originally analyzed
line results from mutation in a closely linked gene
(Michalko et al., 2015). Thus, rather than being an essential
gene, ABP1 is apparently completely dispensable
for normal plant growth and development under
lab conditions. Compounding the problem, an isolate of
the abp1-5 allele was found to include thousands of
polymorphisms compared with the parental genetic
background, as well as a chromosomal segment derived
from a different background, consistent with errors
in the backcrossing regime that followed the
isolation of this allele (Enders et al., 2015). As a result,
the phenotypic differences between this line and its
wild-type previously attributed to the abp1-5 allele may
in fact result from other mutations in the background.
Together, these results require a major reexamination of
the evidence supporting a functionally significant role
for ABP1, including in endocytosis, cytoskeletal arrangement,
and ROP modulation. Clean null mutants
are now available, and assessment of these auxin responses
in the new mutant background will establish
whether they require ABP1.
ABP1 is highly conserved across the plant kingdom,
although interestingly it is apparently missing from the
M. polymorpha genome (Kato et al., 2015). Nonetheless,
this suggests that it confers a significant selective advantage,
despite the fact that it is not required for the
auxin-regulated processes so far examined, and to the
extent to which they have been examined in abp1 null
mutants. It may be that either fully parallel systems
and/or different auxin response systems regulate the
same processes, resulting in functional redundancy.
This is the case for auxin-regulated root growth described
above. Here, the very earliest stages of gravitropic bending
depend on auxin-stimulated Ca2+
influx, mediated independently
of the TIR1/AFB-Aux/IAA-ARF system.
However, the slightly later effects of the TIR1/AFBAux/IAA-ARF
system on gravitropism all but mask the
loss of the nontranscriptional pathway at a macroscopic
level. For this response, however, auxin-stimulated Ca2+
influx has been assessed in an abp1 null mutant background
and found to be normal (Shih et al., 2015).
It is noteworthy that for many of the cell biological
responses reported as being ABP1-dependent, associated
whole-plant level phenotypes have not been examined
in much detail, and where they have, relatively
modest effects are typically reported, despite strong
effects reported for the cellular level (Robert et al., 2010;
Chen et al., 2014; Baskin, 2015). This is consistent with
the idea that these cell biological responses are either
not very important at the organismal level, or there is
sufficient redundancy in their regulation to mask any
morphological effects of ABP1 manipulation. There are,
however, some examples of strong morphological effects
from perturbed ABP1 levels, including standard
ABP1 antisense expression (Braun et al., 2008; Tromas
et al., 2009). These experiments involve inducible antisense
expression, raising the possibility that sudden
changes in ABP1 levels have more significant effects
than stable loss of function. This could be consistent
with one or more redundant compensating systems
requiring time to reequilibrate in response to ABP1 loss.
Whatever the role of ABP1, it is clear that it is not
sufficient to explain the known and/or strongly suspected
nontranscriptional effects of auxin, for example,
on root Ca2+
spiking. There must therefore be other
auxin perception systems. There are certainly additional
biochemically identified auxin binding proteins,
although there is very limited evidence that they play
any role in auxin responses (Napier and Venis, 1995).
Perhaps the best-supported auxin receptor mediating
an at least partially nontranscriptional auxin response is
the Arabidopsis S-Phase Kinase-Associated Protein2A
(SKP2A). SKP2A is an F-box protein with structural
similarities to the TIR1 family (Jurado et al., 2008, 2010).
It regulates cell cycle progression by promoting degradation
of cell cycle transcriptional regulators including
E2FC and DPB (del Pozo et al., 2006). Auxin has
been shown to trigger these degradative events as well
as trigger degradation of SKP2A itself (Jurado et al.,
2010). Structural modeling against TIR1 suggested that
auxin could bind directly to SKP2A, and this was experimentally
validated. Furthermore, mutation of the
predicted auxin binding site compromised auxin binding,
as well as auxin-induced destabilization of SKP2A
and interaction with DPB. These data directly link auxin
to cell cycle progression, and the functional significance
of this link is supported by the auxin-resistant root
growth phenotype of plants in overexpressing SPK2A
that is unable to bind auxin.
SUMMARY AND PERSPECTIVES
Auxin can be considered as a general coordinator of
growth and development, used and reused throughout
the life cycle of plants to mediate communication between
cells and tissues at short and long ranges. How
this information is decoded at the receiving tissues is
unsurprisingly complex. Relevant information is present
in the absolute as well as spatially and temporally
relative levels of auxin. The appropriate response to all
this information depends on myriad other factors, requiring
an extensive and highly tunable informationprocessing
system in every cell. The TIR1/AFB-Aux/
IAA-ARF system provides impressive power to deliver
the necessary response properties, but there is mounting
evidence that it cannot and does not account for all auxin
Plant Physiol. Vol. 176, 2018 475
Auxin Signaling
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responses. Additional auxin binding activities, for example,
mediated by ETTIN and SKP2A, have been
linked to specific downstream responses, and specific
auxin responses unlikely to be mediated by known
auxin reception systems have been identified, such as
root Ca2+
transients and root hair positioning. Evolutionary
approaches are a promising route for the characterization
of these auxin responses, and integration of
computational modeling continues to make important
contributions. However, limitations in currently available
in vivo auxin detection systems are a major constraint
on progress.
Received June 7, 2017; accepted August 17, 2017; published August 17, 2017.
LITERATURE CITED
Abel S, Oeller PW, Theologis A (1994) Early auxin-induced genes encode
short-lived nuclear proteins. Proc Natl Acad Sci USA 91: 326–330
Abel S, Theologis A (1996) Early genes and auxin action. Plant Physiol 111: 9–17
Abley K, De Reuille PB, Strutt D, Bangham A, Prusinkiewicz P, Marée
AF, Grieneisen VA, Coen E (2013) An intracellular partitioning-based
framework for tissue cell polarity in plants and animals. Development
140: 2061–2074
Abley K, Sauret-Güeto S, Marée AFM, Coen E (2016) Formation of polarity
convergences underlying shoot outgrowths. eLife 5: e18165
Badescu GO, Napier RM (2006) Receptors for auxin: Will it all end in TIRs?
Trends Plant Sci 11: 217–223
Band LR, Wells DM, Larrieu A, Sun J, Middleton AM, French AP, Brunoud
G, Sato EM, Wilson MH, Péret B, et al (2012) Root gravitropism is regulated
by a transient lateral auxin gradient controlled by a tipping-point mechanism.
Proc Natl Acad Sci USA 109: 4668–4673
Bargmann BO, Vanneste S, Krouk G, Nawy T, Efroni I, Shani E, Choe G,
Friml J, Bergmann DC, Estelle M, et al (2013) A map of cell typespecific
auxin responses. Mol Syst Biol 9: 688
Baskin TI (2015) Auxin inhibits expansion rate independently of cortical
microtubules. Trends Plant Sci 20: 471–472
Bennett T, Hines G, Leyser O (2014) Canalization: what the flux? Trends
Genet 30: 41–48
Bennett T, Leyser O (2014) The auxin question: a philosophical overview.
In E Zazímalová, J Petrasek, E Benkova, eds, Auxin and Its Role in Plant
Development. Springer, Berlin, pp 3–20
Bhatia N, Bozorg B, Larsson A, Ohno C, Jönsson H, Heisler MG (2016)
Auxin acts through MONOPTEROS to regulate plant cell polarity and
pattern phyllotaxis. Curr Biol 26: 3202–3208
Blatt MR, Thiel G (1994) K+ channels of stomatal guard cells: bimodal
control of the K+ inward-rectifier evoked by auxin. Plant J 5: 55–68
Blilou I, Xu J, Wildwater M, Willemsen V, Paponov I, Friml J, Heidstra R,
Aida M, Palme K, Scheres B (2005) The PIN auxin efflux facilitator network
controls growth and patterning in Arabidopsis roots. Nature 433: 39–44
Boer DR, Freire-Rios A, van den Berg WA, Saaki T, Manfield IW, Kepinski S,
López-Vidrieo I, Franco-Zorrilla JM, de Vries SC, Solano R, et al (2014)
Structural basis for DNA binding specificity by the auxin-dependent ARF
transcription factors. Cell 156: 577–589
Braun N, Wyrzykowska J, Muller P, David K, Couch D, Perrot-Rechenmann
C, Fleming AJ (2008) Conditional repression of AUXIN BINDING PROTEIN1
reveals that it coordinates cell division and cell expansion during
postembryonic shoot development in Arabidopsis and tobacco. Plant Cell
20: 2746–2762
Bridge LJ, Mirams GR, Kieffer ML, King JR, Kepinski S (2012) Distinguishing
possible mechanisms for auxin-mediated developmental
control in Arabidopsis: models with two Aux/IAA and ARF proteins,
and two target gene-sets. Math Biosci 235: 32–44
Calderón Villalobos LI, Lee S, De Oliveira C, Ivetac A, Brandt W, Armitage L,
Sheard LB, Tan X, Parry G, Mao H, et al (2012) A combinatorial TIR1/AFBAux/IAA
co-receptor system for differential sensing of auxin. Nat Chem
Biol 8: 477–485
Chae K, Isaacs CG, Reeves PH, Maloney GS, Muday GK, Nagpal P, Reed
JW (2012) Arabidopsis SMALL AUXIN UP RNA63 promotes hypocotyl
and stamen filament elongation. Plant J 71: 684–697
Chapman EJ, Estelle M (2009) Mechanism of auxin-regulated gene expression
in plants. Annu Rev Genet 43: 265–285
Chen JG, Ullah H, Young JC, Sussman MR, Jones AM (2001) ABP1 is
required for organized cell elongation and division in Arabidopsis embryogenesis.
Genes Dev 15: 902–911
Chen X, Grandont L, Li H, Hauschild R, Paque S, Abuzeineh A, Rakusová H,
Benková E, Perrot-Rechenmann C, Friml J (2014) Inhibition of cell expansion
by rapid ABP1-mediated auxin effect on microtubules. Nature
516: 90–93
Chen X, Naramoto S, Robert S, Tejos R, Löfke C, Lin D, Yang Z, Friml J
(2012) ABP1 and ROP6 GTPase signaling regulate clathrin-mediated
endocytosis in Arabidopsis roots. Curr Biol 22: 1326–1332
Chuang HW, Zhang W, Gray WM (2004) Arabidopsis ETA2, an apparent ortholog
of the human cullin-interacting protein CAND1, is required for auxin
responses mediated by the SCFTIR1
ubiquitin ligase. Plant Cell 16: 1883–1897
Cieslak M, Runions A, Prusinkiewicz P (2015) Auxin-driven patterning
with unidirectional fluxes. J Exp Bot 66: 5083–5102
del Pozo JC, Dharmasiri S, Hellmann H, Walker L, Gray WM, Estelle M
(2002) AXR1-ECR1-dependent conjugation of RUB1 to the Arabidopsis
Cullin AtCUL1 is required for auxin response. Plant Cell 14: 421–433
del Pozo JC, Diaz-Trivino S, Cisneros N, Gutierrez C (2006) The balance
between cell division and endoreplication depends on E2FC-DPB,
transcription factors regulated by the ubiquitin-SCFSKP2A
pathway in
Arabidopsis. Plant Cell 18: 2224–2235
Dharmasiri N, Dharmasiri S, Weijers D, Lechner E, Yamada M, Hobbie L,
Ehrismann JS, Jürgens G, Estelle M (2005) Plant development is regulated
by a family of auxin receptor F box proteins. Dev Cell 9: 109–119
Dinesh DC, Kovermann M, Gopalswamy M, Hellmuth A, Calderón Villalobos
LI, Lilie H, Balbach J, Abel S (2015) Solution structure of the PsIAA4 oligomerization
domain reveals interaction modes for transcription factors in
early auxin response. Proc Natl Acad Sci USA 112: 6230–6235
Dreher KA, Brown J, Saw RE, Callis J (2006) The Arabidopsis Aux/IAA
protein family has diversified in degradation and auxin responsiveness.
Plant Cell 18: 699–714
Elser JJ, Bracken MES, Cleland EE, Gruner DS, Harpole WS, Hillebrand
H, Ngai JT, Seabloom EW, Shurin JB, Smith JE (2007) Global analysis
of nitrogen and phosphorus limitation of primary producers in freshwater,
marine and terrestrial ecosystems. Ecol Lett 10: 1135–1142
Enders TA, Oh S, Yang Z, Montgomery BL, Strader LC (2015) Genome
sequencing of Arabidopsis abp1-5 reveals second-site mutations that
may affect phenotypes. Plant Cell 27: 1820–1826
Estelle M, Weijers D, Ljung K, Leyser O, editors (2011) Auxin Signaling:
From Synthesis to Systems Biology. Cold Spring Harbor Press, Cold
Spring Harbor, NY
Fendrych M, Leung J, Friml J (2016) TIR1/AFB-Aux/IAA auxin perception
mediates rapid cell wall acidification and growth of Arabidopsis hypocotyls.
eLife 5: e19048
Fischer U, Ikeda Y, Ljung K, Serralbo O, Singh M, Heidstra R, Palme K,
Scheres B, Grebe M (2006) Vectorial information for Arabidopsis planar
polarity is mediated by combined AUX1, EIN2, and GNOM activity.
Curr Biol 16: 2143–2149
Flores-Sandoval E, Eklund DM, Bowman JL (2015) A simple auxin transcriptional
response system regulates multiple morphogenetic processes
in the liverwort Marchantia polymorpha. PLoS Genet 11: e1005207
Friml J, Jones AR (2010) Endoplasmic reticulum: the rising compartment in
auxin biology. Plant Physiol 154: 458–462
Friml J, Wisniewska J, Benková E, Mendgen K, Palme K (2002) Lateral
relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis.
Nature 415: 806–809
Fuglsang AT, Visconti S, Drumm K, Jahn T, Stensballe A, Mattei B,
Jensen ON, Aducci P, Palmgren MG (1999) Binding of 14-3-3 protein to
the plasma membrane H(+)-ATPase AHA2 involves the three C-terminal
residues Tyr(946)-Thr-Val and requires phosphorylation of Thr(947). J Biol
Chem 274: 36774–36780
Galinha C, Hofhuis H, Luijten M, Willemsen V, Blilou I, Heidstra R,
Scheres B (2007) PLETHORA proteins as dose-dependent master regulators
of Arabidopsis root development. Nature 449: 1053–1057
Gao Y, Zhang Y, Zhang D, Dai X, Estelle M, Zhao Y (2015) Auxin binding
protein 1 (ABP1) is not required for either auxin signaling or Arabidopsis
development. Proc Natl Acad Sci USA 112: 2275–2280
Gray WM, Kepinski S, Rouse D, Leyser O, Estelle M (2001) Auxin regulates
SCFTIR1
-dependent degradation of AUX/IAA proteins. Nature
414: 271–276
476 Plant Physiol. Vol. 176, 2018
Leyser
www.plantphysiol.orgon January 9, 2018 - Published byDownloaded from
Copyright © 2018 American Society of Plant Biologists. All rights reserved.
Gray WM, Muskett PR, Chuang HW, Parker JE (2003) Arabidopsis SGT1b
is required for SCF(TIR1)-mediated auxin response. Plant Cell 15: 1310–
1319
Grieneisen VA, Xu J, Marée AF, Hogeweg P, Scheres B (2007) Auxin
transport is sufficient to generate a maximum and gradient guiding root
growth. Nature 449: 1008–1013
Guilfoyle TJ (2015) The PB1 domain in auxin response factor and Aux/IAA
proteins: a versatile protein interaction module in the auxin response.
Plant Cell 27: 33–43
Guseman JM, Hellmuth A, Lanctot A, Feldman TP, Moss BL, Klavins E,
Calderón Villalobos LI, Nemhauser JL (2015) Auxin-induced degradation
dynamics set the pace for lateral root development. Development
142: 905–909
Hager A (2003) Role of the plasma membrane H+-ATPase in auxin-induced
elongation growth: historical and new aspects. J Plant Res 116: 483–505
Harrison BR, Masson PH (2008) ARL2, ARG1 and PIN3 define a gravity
signal transduction pathway in root statocytes. Plant J 53: 380–392
Havens KA, Guseman JM, Jang SS, Pierre-Jerome E, Bolten N, Klavins E,
Nemhauser JL (2012) A synthetic approach reveals extensive tunability
of auxin signaling. Plant Physiol 160: 135–142
Hazak O, Bloch D, Poraty L, Sternberg H, Zhang J, Friml J, Yalovsky S
(2010) A rho scaffold integrates the secretory system with feedback
mechanisms in regulation of auxin distribution. PLoS Biol 8: e1000282
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 Biol 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. Curr Biol 15: 1899–1911
Hellmann H, Hobbie L, Chapman A, Dharmasiri S, Dharmasiri N, del
Pozo C, Reinhardt D, Estelle M (2003) Arabidopsis AXR6 encodes CUL1
implicating SCF E3 ligases in auxin regulation of embryogenesis. EMBO
J 22: 3314–3325
Ikeda Y, Men S, Fischer U, Stepanova AN, Alonso JM, Ljung K, Grebe M
(2009) Local auxin biosynthesis modulates gradient-directed planar
polarity in Arabidopsis. Nat Cell Biol 11: 731–738
Jilkine A, Marée AF, Edelstein-Keshet L (2007) Mathematical model for
spatial segregation of the Rho-family GTPases based on inhibitory
crosstalk. Bull Math Biol 69: 1943–1978
Jones AM, Herman EM (1993) KDEL-containing auxin-binding protein is
secreted to the plasma membrane and cell wall. Plant Physiol 101: 595–
606
Jones AM, Im KH, Savka MA, Wu MJ, DeWitt NG, Shillito R, Binns AN
(1998) Auxin-dependent cell expansion mediated by overexpressed
auxin-binding protein 1. Science 282: 1114–1117
Jones MA, Shen J-J, Fu Y, Li H, Yang Z, Grierson CS (2002) The Arabidopsis
Rop2 GTPase is a positive regulator of both root hair initiation
and tip growth. Plant Cell 14: 763–776
Jönsson H, Heisler MG, Shapiro BE, Meyerowitz EM, Mjolsness E (2006)
An auxin-driven polarized transport model for phyllotaxis. Proc Natl
Acad Sci USA 103: 1633–1638
Jurado S, Abraham Z, Manzano C, López-Torrejón G, Pacios LF, Del Pozo
JC (2010) The Arabidopsis cell cycle F-box protein SKP2A binds to
auxin. Plant Cell 22: 3891–3904
Jurado S, Díaz-Triviño S, Abraham Z, Manzano C, Gutierrez C, del Pozo
C (2008) SKP2A, an F-box protein that regulates cell division, is degraded
via the ubiquitin pathway. Plant J 53: 828–841
Kanczewska J, Marco S, Vandermeeren C, Maudoux O, Rigaud JL, Boutry
M (2005) Activation of the plant plasma membrane H+-ATPase by
phosphorylation and binding of 14-3-3 proteins converts a dimer into a
hexamer. Proc Natl Acad Sci USA 102: 11675–11680
Kato H, Ishizaki K, Kouno M, Shirakawa M, Bowman JL, Nishihama R,
Kohchi T (2015) Auxin-mediated transcriptional system with a minimal
set of components is critical for morphogenesis through the life cycle in
Marchantia polymorpha. PLoS Genet 11: e1005084
Klode M, Dahlke RI, Sauter M, Steffens B (2011) Expression and subcellular
localization of Arabidopsis thaliana auxin-binding protein
1 (ABP1). J Plant Growth Regul 30: 416–424
Knox K, Grierson CS, Leyser O (2003) AXR3 and SHY2 interact to regulate
root hair development. Development 130: 5769–5777
Korasick DA, Westfall CS, Lee SG, Nanao MH, Dumas R, Hagen G,
Guilfoyle TJ, Jez JM, Strader LC (2014) Molecular basis for AUXIN
RESPONSE FACTOR protein interaction and the control of auxin response
repression. Proc Natl Acad Sci USA 111: 5427–5432
Kramer EM (2009) Auxin-regulated cell polarity: an inside job? Trends
Plant Sci 14: 242–247
Kutschera U (1994) The current status of the acid-growth hypothesis. New
Phytol 126: 549–569
Lavy M, Prigge MJ, Tao S, Shain S, Kuo A, Kirchsteiger K, Estelle M
(2016) Constitutive auxin response in Physcomitrella reveals complex
interactions between Aux/IAA and ARF proteins. eLife 5: e13325
Leblanc N, David K, Grosclaude J, Pradier JM, Barbier-Brygoo H, Labiau S,
Perrot-Rechenmann C (1999) A novel immunological approach establishes
that the auxin-binding protein, Nt-abp1, is an element involved in auxin
signaling at the plasma membrane. J Biol Chem 274: 28314–28320
Leyser HMO (1997) Auxin: lessons from a mutant weed. Physiol Plant 100:
407–414
Leyser O (2005) Auxin distribution and plant pattern formation: How
many angels can dance on the point of PIN? Cell 121: 819–822
Leyser O (2016) Angiosperm multicellularity: the whole, the parts and the
sum. In KJ Niklas, SA Newman, eds, The Evolution and Consequences
of Multicellularity. Vienna Series of Theoretical Biology. MIT Press,
Cambridge, MA, pp 87–102
Löbler M, Klämbt D (1985a) Auxin-binding protein from coleoptile
membranes of corn (Zea mays L.). I. Purification by immunological
methods and characterization. J Biol Chem 260: 9848–9853
Löbler M, Klämbt D (1985b) Auxin-binding protein from coleoptile
membranes of corn (Zea mays L.). II. Localization of a putative auxin
receptor. J Biol Chem 260: 9854–9859
Luschnig C, Gaxiola RA, Grisafi P, Fink GR (1998) EIR1, a root-specific
protein involved in auxin transport, is required for gravitropism in
Arabidopsis thaliana. Genes Dev 12: 2175–2187
Mähönen AP, Ten Tusscher K, Siligato R, Smetana O, Díaz-Triviño S,
Salojärvi J, Wachsman G, Prasad K, Heidstra R, Scheres B (2014)
PLETHORA gradient formation mechanism separates auxin responses.
Nature 515: 125–129
Maraschin FdosS, Memelink J, Offringa R (2009) Auxin-induced, SCF
(TIR1)-mediated poly-ubiquitination marks AUX/IAA proteins for
degradation. Plant J 59: 100–109
McClure BA, Hagen G, Brown CS, Gee MA, Guilfoyle TJ (1989) Transcription,
organization, and sequence of an auxin-regulated gene cluster
in soybean. Plant Cell 1: 229–239
Michalko J, Dravecká M, Bollenbach T, Friml J (2015) Embryo-lethal
phenotypes in early abp1 mutants are due to disruption of the neighboring
BSM gene. F1000 Res 4: 1104
Mironova VV, Omelyanchuk NA, Wiebe DS, Levitsky VG (2014) Computational
analysis of auxin responsive elements in the Arabidopsis
thaliana L. genome. BMC Genomics (Suppl 12) 15: S4
Mitchison GJ (1980) A model for vein formation in higher plants. Proc R
Soc Lond B Biol Sci 207: 79–109
Molendijk AJ, Bischoff F, Rajendrakumar CSV, Friml J, Braun M, Gilroy
S, Palme K (2001) Arabidopsis thaliana Rop GTPases are localized to tips
of root hairs and control polar growth. EMBO J 20: 2779–2788
Monshausen GB, Miller ND, Murphy AS, Gilroy S (2011) Dynamics of
auxin-dependent Ca2+ and pH signaling in root growth revealed by
integrating high-resolution imaging with automated computer visionbased
analysis. Plant J 65: 309–318
Moss BL, Mao H, Guseman JM, Hinds TR, Hellmuth A, Kovenock M,
Noorassa A, Lanctot A, Villalobos LI, Zheng N, et al (2015) Rate motifs
tune auxin/indole-3-acetic acid degradation dynamics. Plant Physiol
169: 803–813
Müller A, Guan C, Gälweiler L, Tänzler P, Huijser P, Marchant A, Parry
G, Bennett M, Wisman E, Palme K (1998) AtPIN2 defines a locus of
Arabidopsis for root gravitropism control. EMBO J 17: 6903–6911
Nakamura M, Kiefer CS, Grebe M (2012) Planar polarity, tissue polarity
and planar morphogenesis in plants. Curr Opin Plant Biol 15: 593–600
Nanao MH, Vinos-Poyo T, Brunoud G, Thévenon E, Mazzoleni M, Mast D,
Lainé S, Wang S, Hagen G, Li H, et al (2014) Structural basis for oligomerization
of auxin transcriptional regulators. Nat Commun 5: 3617
Napier R (1995) Towards an understanding of ABP1. J Exp Bot 46: 1787–
1795
Napier RM (1997) Trafficking of the auxin-binding protein. Trends Plant
Sci 2: 251–255
Plant Physiol. Vol. 176, 2018 477
Auxin Signaling
www.plantphysiol.orgon January 9, 2018 - Published byDownloaded from
Copyright © 2018 American Society of Plant Biologists. All rights reserved.
Napier RM, Venis MA (1995) Auxin action and auxin-binding proteins.
New Phytol 129: 167–201
Nemhauser JL, Feldman LJ, Zambryski PC (2000) Auxin and ETTIN in
Arabidopsis gynoecium morphogenesis. Development 127: 3877–3888
Otsuji M, Ishihara S, Co C, Kaibuchi K, Mochizuki A, Kuroda S (2007) A
mass conserved reaction-diffusion system captures properties of cell
polarity. PLOS Comput Biol 3: e108
Paponov IA, Paponov M, Teale W, Menges M, Chakrabortee S, Murray
JA, Palme K (2008) Comprehensive transcriptome analysis of auxin
responses in Arabidopsis. Mol Plant 1: 321–337
Payne RJH, Grierson CS (2009) A theoretical model for ROP localisation by
auxin in Arabidopsis root hair cells. PLoS One 4: e8337
Pierre-Jerome E, Moss BL, Lanctot A, Hageman A, Nemhauser JL (2016)
Functional analysis of molecular interactions in synthetic auxin response
circuits. Proc Natl Acad Sci USA 113: 11354–11359
Prigge MJ, Greenham K, Zhang Y, Santner A, Castillejo C, Mutka AM,
O’Malley RC, Ecker JR, Kunkel BN, Estelle M (2016) The Arabidopsis
auxin receptor F-box proteins AFB4 and AFB5 are required for response
to the synthetic auxin picloram. G3 (Bethesda) 6: 1383–1390
Prigge MJ, Lavy M, Ashton NW, Estelle M (2010) Physcomitrella patens
auxin-resistant mutants affect conserved elements of an auxin-signaling
pathway. Curr Biol 20: 1907–1912
Rademacher EH, Lokerse AS, Schlereth A, Llavata-Peris CI, Bayer M,
Kientz M, Freire Rios A, Borst JW, Lukowitz W, Jürgens G, et al (2012)
Different auxin response machineries control distinct cell fates in the
early plant embryo. Dev Cell 22: 211–222
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, Sauer M, Paciorek T, Baster P, Vanneste S,
Zhang J, Simon S, Čovanová M, et al (2010) ABP1 mediates auxin inhibition
of clathrin-dependent endocytosis in Arabidopsis. Cell 143:
111–121
Rolland-Lagan AG, Prusinkiewicz P (2005) Reviewing models of auxin
canalization in the context of leaf vein pattern formation in Arabidopsis.
Plant J 44: 854–865
Rück A, Palme K, Venis MA, Napier RM, Felle HH (1993) Patch-clamp
analysis establishes a role for an auxin binding protein in the auxin
stimulation of plasma membrane current in Zea mays protoplasts. Plant
J 4: 41–46
Sabatini S, Beis D, Wolkenfelt H, Murfett J, Guilfoyle T, Malamy J,
Benfey P, Leyser O, Bechtold N, Weisbeek P, Scheres B (1999) An
auxin-dependent distal organizer of pattern and polarity in the Arabidopsis
root. Cell 99: 463–472
Sachs T (1968) On the determination of the pattern of vascular tissue in
peas. Ann Bot (Lond) 32: 781–790
Sachs T (1969) Polarity and the induction of organized vascular tissues.
Ann Bot (Lond) 33: 263–275
Sachs T (1981) The control of the patterned differentiation of vascular tissues.
Adv Bot Res 9: 151–162
Salehin M, Bagchi R, Estelle M (2015) SCFTIR1/AFB-based auxin perception:
mechanism and role in plant growth and development. Plant
Cell 27: 9–19
Sauer M, Balla J, Luschnig C, Wisniewska J, Reinöhl V, Friml J, Benková
E (2006) Canalization of auxin flow by Aux/IAA-ARF-dependent
feedback regulation of PIN polarity. Genes Dev 20: 2902–2911
Sawchuk MG, Edgar A, Scarpella E (2013) Patterning of leaf vein networks
by convergent auxin transport pathways. PLoS Genet 9: e1003294
Scarpella E, Marcos D, Friml J, Berleth T (2006) Control of leaf vascular
patterning by polar auxin transport. Genes Dev 20: 1015–1027
Sessions A, Nemhauser JL, McColl A, Roe JL, Feldmann KA, Zambryski
PC (1997) ETTIN patterns the Arabidopsis floral meristem and reproductive
organs. Development 124: 4481–4491
Shih HW, DePew CL, Miller ND, Monshausen GB (2015) The cyclic
nucleotide-gated channel CNGC14 regulates root gravitropism in Arabidopsis
thaliana. Curr Biol 25: 3119–3125
Simonini S, Deb J, Moubayidin L, Stephenson P, Valluru M, Freire-Rios
A, Sorefan K, Weijers D, Friml J, Østergaard L (2016) A noncanonical
auxin-sensing mechanism is required for organ morphogenesis in Arabidopsis.
Genes Dev 30: 2286–2296
Smalle J, Vierstra RD (2004) The ubiquitin 26S proteasome proteolytic
pathway. Annu Rev Plant Biol 55: 555–590
Smith RS, Guyomarc’h S, Mandel T, Reinhardt D, Kuhlemeier C, Prusinkiewicz
P (2006) A plausible model of phyllotaxis. Proc Natl Acad Sci USA 103:
1301–1306
Sorek N, Poraty L, Sternberg H, Bar E, Lewinsohn E, Yalovsky S (2007) Activation
status-coupled transient S acylation determines membrane partitioning
of a plant Rho-related GTPase. Mol Cell Biol 27: 2144–2154; retraction
Sorek N, Poraty L, Sternberg H, Bar E, Lewinsohn E, Yalovsky S (2017)
Mol Cell Biol 37: e00321–17
Spartz AK, Lee SH, Wenger JP, Gonzalez N, Itoh H, Inzé D, Peer WA,
Murphy AS, Overvoorde PJ, Gray WM (2012) The SAUR19 subfamily
of SMALL AUXIN UP RNA genes promote cell expansion. Plant J 70:
978–990
Spartz AK, Lor VS, Ren H, Olszewski NE, Miller ND, Wu G, Spalding
EP, Gray WM (2017) Constitutive expression of Arabidopsis SMALL
AUXIN UP RNA19 (SAUR19) in tomato confers auxin-independent hypocotyl
elongation. Plant Physiol 173: 1453–1462
Spartz AK, Ren H, Park MY, Grandt KN, Lee SH, Murphy AS, Sussman
MR, Overvoorde PJ, Gray WM (2014) SAUR inhibition of PP2C-D
phosphatases activates plasma membrane H+-ATPases to promote cell
expansion in Arabidopsis. Plant Cell 26: 2129–2142
Steffens B, Feckler C, Palme K, Christian M, Böttger M, Lüthen H (2001)
The auxin signal for protoplast swelling is perceived by extracellular
ABP1. Plant J 27: 591–599
Stewart JL, Nemhauser JL (2010) Do trees grow on money? Auxin as the
currency of the cellular economy. Cold Spring Harb Perspect Biol 2:
a001420
Stirnberg P, Liu J-P, Ward S, Kendall SL, Leyser O (2012) Mutation of the
cytosolic ribosomal protein-encoding RPS10B gene affects shoot meristematic
function in Arabidopsis. BMC Plant Biol 12: 160
Sun N, Wang J, Gao Z, Dong J, He H, Terzaghi W, Wei N, Deng XW, Chen
H (2016) Arabidopsis SAURs are critical for differential light regulation of
the development of various organs. Proc Natl Acad Sci USA 113: 6071–
6076
Swarup R, Kramer EM, Perry P, Knox K, Leyser HM, Haseloff J, Beemster
GT, Bhalerao R, Bennett MJ (2005) Root gravitropism requires lateral
root cap and epidermal cells for transport and response to a mobile
auxin signal. Nat Cell Biol 7: 1057–1065
Szemenyei H, Hannon M, Long JA (2008) TOPLESS mediates auxindependent
transcriptional repression during Arabidopsis embryogenesis.
Science 319: 1384–1386
Takahashi K, Hayashi K, Kinoshita T (2012) Auxin activates the plasma
membrane H+-ATPase by phosphorylation during hypocotyl elongation
in Arabidopsis. Plant Physiol 159: 632–641
Tan X, Calderon-Villalobos LIA, Sharon M, Zheng C, Robinson CV,
Estelle M, Zheng N (2007) Mechanism of auxin perception by the TIR1
ubiquitin ligase. Nature 446: 640–645
Tao LZ, Cheung AY, Wu HM (2002) Plant Rac-like GTPases are activated
by auxin and mediate auxin-responsive gene expression. Plant Cell 14:
2745–2760
Thelander M, Olsson T, Ronne H (2005) Effect of the energy supply on
filamentous growth and development in Physcomitrella patens. J Exp Bot
56: 653–662
Tiwari SB, Hagen G, Guilfoyle TJ (2004) Aux/IAA proteins contain a
potent transcriptional repression domain. Plant Cell 16: 533–543
Tromas A, Braun N, Muller P, Khodus T, Paponov IA, Palme K, Ljung K,
Lee J-Y, Benfey P, Murray JAH, et al (2009) The AUXIN BINDING
PROTEIN 1 is required for differential auxin responses mediating root
growth. PLoS One 4: e6648
Turing AM (1952) The chemical basis of morphogenesis. Philos Trans R Soc
Lond B Biol Sci 237: 37–72
Ulmasov T, Hagen G, Guilfoyle TJ (1999) Activation and repression of
transcription by auxin-response factors. Proc Natl Acad Sci USA 96:
5844–5849
Ulmasov T, Murfett J, Hagen G, Guilfoyle TJ (1997) Aux/IAA proteins
repress expression of reporter genes containing natural and highly active
synthetic auxin response elements. Plant Cell 9: 1963–1971
Venis MA, Napier RM, Barbier-Brygoo H, Maurel C, Perrot-Rechenmann
C, Guern J (1992) Antibodies to a peptide from the maize auxin-binding
protein have auxin agonist activity. Proc Natl Acad Sci USA 89: 7208–
7212
478 Plant Physiol. Vol. 176, 2018
Leyser
www.plantphysiol.orgon January 9, 2018 - Published byDownloaded from
Copyright © 2018 American Society of Plant Biologists. All rights reserved.
Vernoux T, Brunoud G, Farcot E, Morin V, Van den Daele H, Legrand J,
Oliva M, Das P, Larrieu A, Wells D, et al (2011) The auxin signalling
network translates dynamic input into robust patterning at the shoot
apex. Mol Syst Biol 7: 508
Vert G, Walcher CL, Chory J, Nemhauser JL (2008) Integration of auxin
and brassinosteroid pathways by Auxin Response Factor 2. Proc Natl
Acad Sci USA 105: 9829–9834
Weyers JDB, Paterson NW (2001) Plant hormones and the control of
physiological processes. New Phytol 152: 375–407
Woo E-J, Marshall J, Bauly J, Chen J-G, Venis M, Napier RM, Pickersgill
RW (2002) Crystal structure of auxin-binding protein 1 in complex with
auxin. EMBO J 21: 2877–2885
Wu M-F, Yamaguchi N, Xiao J, Bargmann B, Estelle M, Sang Y, Wagner D
(2015) Auxin-regulated chromatin switch directs acquisition of flower
primordium founder fate. eLife 4: e09269
Xu T, Dai N, Chen J, Nagawa S, Cao M, Li H, Zhou Z, Chen X, De Rycke
R, Rakusová H, et al (2014) Cell surface ABP1-TMK auxin-sensing
complex activates ROP GTPase signaling. Science 343: 1025–1028
Xu T, Wen M, Nagawa S, Fu Y, Chen JG, Wu MJ, Perrot-Rechenmann C, Friml
J, Jones AM, Yang Z (2010) Cell surface- and rho GTPase-based auxin signaling
controls cellular interdigitation in Arabidopsis. Cell 143: 99–110
Yalovsky S, Bloch D, Sorek N, Kost B (2008) Regulation of membrane
trafficking, cytoskeleton dynamics, and cell polarity by ROP/RAC
GTPases. Plant Physiol 147: 1527–1543
Yamagami M, Haga K, Napier RM, Iino M (2004) Two distinct signaling
pathways participate in auxin-induced swelling of pea epidermal protoplasts.
Plant Physiol 134: 735–747
Zenser N, Ellsmore A, Leasure C, Callis J (2001) Auxin modulates the
degradation rate of Aux/IAA proteins. Proc Natl Acad Sci USA 98:
11795–11800
Plant Physiol. Vol. 176, 2018 479
Auxin Signaling
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