Buschmann et Borchers 2019 Phytologist Tansley review

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Tansley review
Handedness in plant cell expansion: a mutant
perspective on helical growth
Author for correspondence:
Henrik Buschmann
Tel: +49 541 9692248
Email: henrik.buschmann@biologie.
uni-osnabrueck.de
Received: 19 March 2019
Accepted: 4 June 2019
Henrik Buschmann and Agnes Borchers
Botanical Institute, Biology and Chemistry Department, University of Osnabr€uck, 49076, Osnabr€uck, Germany
Contents
Summary 1
I. Introduction 2
II. The cell wall’s function in diffuse growth 2
III. Microtubules have a say: the controlled movement of
cellulose synthase 3
IV. Artificial helical growth exhibited by plant mutants 5
V. The mechanics behind twisting: from cells to organs 5
VI. Evidence for microtubules having a profound role in
helical growth 8
VII. Insight from epistasis 9
VIII. The microtubule plus tip and helical growth 9
IX. Helical growth and the salt stress syndrome 10
X.
The emerging role of calcium-mediated signaling in
twisted growth 11
XI. What is the significance of twisted cell wall microfibrils? 11
XII. Further evidence for a directional influence of the cell wall 11
XIII. Tropism-induced helical growth and TORTIFOLIA1/SPIRAL2 12
XIV. Conclusions and outlook 13
Acknowledgements 13
References 13
New Phytologist (2019)
doi: 10.1111/nph.16034
Key words: calcium, cell expansion, cell wall
integrity, handedness, helical growth,
microtubule, symmetry, tropism.
Summary
Many plant mutants are known that exhibit some degree of helical growth. This ‘twisted’
phenotype has arisen frequently in mutant screens of model organisms, but it is also found in
cultivars of ornamental plants, including trees. The phenomenon, in many cases, is based on
defects in cell expansion symmetry. Any complete model which explains the anisotropy of plant
cell growth must ultimately explain how helical cell expansion comes into existence –and how it
is normally avoided. While the mutations observed in model plants mainly point to the
microtubule system, additional affected components involve cell wall functions, auxin transport
and more. Evaluation of published data suggests a two-way mechanism underlying the helical
growth phenomenon: there is, apparently, a microtubular component that determines
handedness, but there is also an influence arising in the cell wall that feeds back into the
cytoplasm and affects cellular handedness. This idea is supported by recent reports
demonstrating the involvement of the cell wall integrity pathway. In addition, there is mounting
evidence that calcium is an important relayer of signals relating to the symmetry of cell
expansion. These concepts suggest experimental approaches to untangle the phenomenon of
helical cell expansion in plant mutants.
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I. Introduction
There are many examples of handedness in plants. Perhaps most
prominent are the twining plants that grow by coiling around a
support, among these bindweed (several species that belong to
the genus Convolvulus), honeysuckle (several species in the genus
Lonicera) and American bittersweet (Celastrus scandens; Fig. 1a).
Most twining plants show either left-handed or right-handed
coils, and the handedness of the coil is a fixed property of a given
species (Edwards et al., 2007; Smyth, 2016). Striking handedness
has been observed at the single-cell level as well, for example in
the tip growing protonema of the moss Ceratodon (Kern et al.,
2005) and in the internode cells of charophyceaen algae (Fig. 1b;
Green, 1954). Other well-known examples of handedness in
plants include spiral phyllotaxis (the handedness by which apical
meristems initiate primordia is usually not inherited) and the
spiral grain of ‘twisted’ stems exhibited by certain trees. Spiral
grain is often stress-induced and can reduce the value of trees if
wood is to be produced from them (Harris, 1989). These
different expressions of handedness likely involve independent
mechanisms (Forterre & Dumais, 2011; Smyth, 2016). In this
review we will discuss plant mutants showing helical handedness
in organs that normally grow straight (Fig. 1c–h). The phe-
nomenon is frequently referred to as helical growth and is
essentially derived from defects in cell expansion (Fig. 1i). This
phenotype results in artificial cell or even organ rotation and
may best be considered an atypical nastic movement (compa-
rable, for example, to induced epinasty). Mutation-induced
helical growth may impinge on the expression of naturally
occurring helical growth, for example during helical tropism. We
aim to summarize the literature on helical growth exhibited by
plant mutants, to discuss some of the controversies present in the
field and, if possible, to establish the cell biological mechanisms
underlying the twisted phenotype. Because the discussion relies
on a set of cell biological paradigms, we will begin with a short
overview on plant growth anisotropy.
II. The cell wall’s function in diffuse growth
The extent and symmetry of plant cell growth is regulated by the cell
wall. In diffuse growth (in contrast to polar growth) extensibility of
the cell wall is present across broader areas of the cell. The wall is a
complex matrix of polysaccharides and proteins. The polysaccha-
rides comprise cellulose, hemicellulose and pectin (Burton et al.,
2010). Cellulose consists of glucose subunits linked by b-1,4-
glycosidic bonds. This polymer is created by a family of mem brane-
localized glycosyltransferases –the so-called cellulose synthases.
Cellulose synthase (CesA) is organized in hexameric rosettes that
were first visualized by electron microscopy (Robi nson et al., 1972).
The enzyme utilizes UDP-glucose sequestered from cytoplasmic
metabolism and extrudes the synthesized cellulose chain into the
extracellular space. Once in the extracellular space, it is now
assumed that 18 glucose chains come together to form the so-called
cellulose microfibril (Lei et al., 2012; Cosgrove, 2018). Curiously,
the formation of the microfibril produces enough force to propel
the cellulase synthase forward (Fig. 2). In this way, the enzyme
continuously travels through the plasma membrane parallel to the
cell’s surface (Bringmann et al., 2012b).
(a)
(e) (g) (i)(h)
(h)
(h)
(f)
(b) (c)
(c)
(c) (d)
α
α
Ψ
250 µm
Fig. 1 Plant helical growth as a result of
handedness in cell and organ expansion. (a)
The right-handed twining plant American
bittersweet (Celastrus scandens) using itself as
support. Bar, 5 mm. (b) The striated
Nitellopsis obtusa giant internode cell as an
example of single-cell helical growth in algae.
(c) Needles of white pine (Pinus strobus). (d)
Examples of needles from Pinus strobus var.
‘Contorta’. The needles tend to be right-
handed, but the strength of the phenotype
varies between different dwarf shoots. (e)
Three-week-old wild-type Arabidopsis plant
with straight petioles. (f) Mutant Arabidopsis
plant (tortifolia1/spiral2) with twisted
petioles. Bars, 1 cm. (g) Cortical microtubules
in petioles of Arabidopsis wild-type (labelled
by a green fluorescent protein fusion to ß-
tubulin). Bar, 5 lm. (h) Left-handed cortical
microtubule arrays in the right-handed mutant
tortifolia1/spiral2. Bar, 5 lm. (i) Model of
helically growing cell. The average
handedness of the structural elements
(cellulose microfibrils) is opposite to the
direction of cell twisting. In this simplified
model the microfibril angle wequals the angle
of growth, a. The model was originally
published by Roelofsen (1965), and was
slightly modified and redrawn for this
publication.
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In diffuse growth, cellulose microfibril alignment is of pivotal
importance for growth anisotropy (i.e. for the direction of growth).
Pioneering work on the internode cells of charophyceaen algae has
shown that cellulose microfibrils are not randomly oriented in the
wall, but are aligned transversely to the axis of cell expansion
(Green, 1962). Generally, growth occurs mainly perpendicularly to
the alignment of the microfibrils because the cellulosic microfibrils
are strong enough to resist the stress produced by turgor pressure.
During growth the microfibrils will be separated in the axial
direction instead (Roelofsen, 1965). When microfibril alignment is
randomized by applying chemical agents, the growth is no longer
channeled in one direction but becomes isotropic. This means that,
because the force generated by turgor is non-directional, the cell
starts swelling and becomes spherical (Baskin, 2005). Typically, the
primary wall of growing cells consists of several layers of
microfibrils, and even strictly anisotropically expanding cells can
have longitudinal microfibrils in their outer layers. This apparent
contradiction is explained by the multinet growth hypothesis. It
states that while the inner microfibrils are load bearing and give
directionality to expansion, the outer microfibrils will passively
reorient and may therefore show deviating angles of orientation
(Roelofsen, 1965).
While the cellulose microfibrils control anisotropy, the question
of whether there is expansion or not depends on other factors
(Szymanski & Cosgrove, 2009). Hemicelluloses (e.g. xyloglucans)
are interwoven with cellulose, and manipulating this interaction is
one way of controlling the rate and extent of cell expansion. For
example, the cutting of xyloglucans by endoglucanases or their
ligation through endotransglycosylases are ways of controlling cell
wall loosening. Pectin also influences the extensibility of the cell
wall, and pectin methylesterases have a function of controlling the
stiffness of the pectin gels in the matrix (Peaucelle et al., 2015). The
secretion of expansins further controls cell wall loosening, albeit by
an unknown mechanism. Expansins (and additional cell wall
enzymes) are pH regulated, which is likely the basis of the ‘acid
growth’ phenomenon. In addition, activation of plasma membrane
NADH oxidase resulting in the redox-dependent regulation of cell
wall loosening or stiffening plays a part in the regulation of cell
expansion (Voxeur & Hofte, 2016).
III. Microtubules have a say: the controlled movement
of cellulose synthase
It was recognized early that colchicine, a poison directed against
spindle fibers (i.e. microtubules), disturbs the cellulose microfibril
patterns of plants (Green, 1962). Soon after, microtubules were
discovered and found to lie just underneath the plant plasma
membrane, forming extended arrays (Ledbetter & Porter, 1963).
These findings together formed the basis for the ‘alignment
hypothesis’ (Fig. 2), which posits that the alignment of intracellular
Plus tip
CC
CMU
CSI
KOR
CC
CMU
CSI
KOR
Microtubule
Hexameric
CesA rosette
Cellulose microfibril
(right-handed twist)
Plasma membrane
COB
Fig. 2 The cellulose microfibril alignment hypothesis and its current model, according to the findings of recent studies (Bringmann et al., 2012b; Lei et al., 2012;
Kesten et al., 2019). Cellulose (blue fibrils) is produced by cellulose synthase rosettes (light red) that are integral to the plasma membrane (yellow). The rosettes
move along microtubules (red bar) an d are propelled forward through the force generated by cellulose polymerization. The microtubules serve as guiding tracks
for the rosettes and determine the direction of cellulose microfibril alignment. The CELLULOSE SYNTHASE INTERACTING (CSI) protein (green) is a linker
protein that connects cellulose synthase rosettes with microtubules. Another recently discovered linker protein is named COMPANION OF CELLULOSE
SYNTHASE (CC) (light green). Further microtubule-associated proteins (MAPs) are involved in the guidance process (e.g. CELLULOSE SYNTHASE-
MICROTUBULE UNCOUPLINGS (CMU; blue circles), but how this is achieved remains unclear. Additional proteins known to be involved in cellulose
biosynthesis are COBRA (COB) and KORRIGAN (KOR). Cellulose microfibrils were repeatedly reported to have a regular (frequently right-handed) twist, which
seems to imply that the rosettes are rotating while spinning out the nascent fibril. How linker proteins CSI and CC would respond to the hypothetical rotation is
currently unclear.
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microtubules controls the patterning of extracellular microfibrils
(Heath, 1974; Giddings & Staehelin, 1991). The hypothesis
became widely accepted but was disputed by some (reviewed by
Baskin, 2001), until eventually fluorescent protein microscopy
allowed for the simultaneous observation of microtubules and
moving cellulose synthase complexes in living Arabidopsis thaliana
cells (Paredez et al., 2006). The analysis showed that cellulose
synthase predominantly travels along the routes delineated by
microtubules (Fig. 2). This holds true even when microtubules are
induced to re-orient by blue light. At the time it remained
enigmatic how cellulose synthases were able to trace the micro-
tubules; however, proteins were meanwhile discovered that appear
to be involved in the guidance process. Arabidopsis CELLULOSE
SYNTHASE INTERACTING1 (CSI1) is a large armadillo repeat
protein that interacts with microtubules and cellulose synthases.
Importantly, it moves together with cellulose synthases, and in the
csi1 knock-out mutant microtubule guidance of cellulose synthase
is reduced (Bringmann et al., 2012a, b; Li et al., 2012). Another
group of proteins that seems to be involved in the guidance process
are the CELLULOSE SYNTHASE-MICROTUBULE
UNCOUPLINGS (CMU) proteins. These proteins, which are
related to kinesin light chain (Burstenbinder et al., 2013), bind to
microtubules of the cortical array but, in contrast to CSI, remain
static on the array. While CMU proteins seem to have a function in
enhancing the rigidity of the microtubule track, it remains to be
determined how exactly they contribute to cellulose synthase
guidance (Liu et al., 2016). Recent research has uncovered an
additional family of proteins involved in cellulose synthase
function –the COMPANION OF CELLULOSE SYNTHASE
(CC) proteins. These proteins, like CSI, bind to CesA as well as
microtubules. CC proteins have a function for continued cellulose
synthesis and microtubule array recovery after salt stress (Endler
et al., 2015; Kesten et al., 2019). Two additional genes involved in
cellulose biosynthesis will be mentioned here. KORRIGAN (KOR)
encodes an endoglucanase that is now assumed to be part of the
cellulose synthase complex. The COBRA (COB) gene encodes an
extracellular GPI-anchored protein (Fig. 2). Mutants of both these
genes have severely reduced cellulose content (Schindelman et al.,
2001; Vain et al., 2014).
Plant cortical microtubules are highly dynamic and array
orientation may change quickly, thereby enabling the formation
of at times complicated cell wall patterns (Lloyd, 2011).
Microtubules are polymers assembled from tubulin heterodimers
that each consist of one polypeptide a-tubulin plus one
polypeptide b-tubulin. The tubulin heterodimers are added to
the microtubule in consistent orientation, providing the growing
polymer with strict polarity (Howard & Hyman, 2003). In vivo
microtubules add tubulin heterodimers predominantly from the
plus end, while the minus end may be stabilized or may be
shrinking through the loss of tubulins (which in the case of
ongoing growth at the plus end will result in ‘treadmilling’). In
the microtubule lattice the tubulin heterodimers are arranged in
linear files so that protofilaments are formed. The protofilaments
interact laterally and form the wall of the tube: usually 13 of
these make up one straight tube with a resulting diameter of c.
25 nm (Nogales et al., 1999; L€owe et al., 2001). b-tubulin is a
GTPase, and this capacity controls the stability of the micro-
tubule. When a tubulin dimer is added to the plus end, the b-
tubulin subunit will carry one molecule of GTP. After attach-
ment to the polymer the GTPase function eventually converts
GTP into GDP. This means that most of the b-tubulin in a
microtubule carries GDP, while at the microtubule plus tip a cap
of GTP is present (Howard & Hyman, 2009). This GTP cap is
important, because loss of the cap will impinge on microtubule
stability and likely cause shrinkage (catastrophe). It is assumed
that the GTP cap reduces the tendency of the protofilament for
outward curvature, while the curvature renders the microtubule
unstable. The function of the GTP cap is therefore the basis of a
phenomenon termed ‘dynamic instability’. This dynamic behav-
ior allows in vivo microtubule plus ends to continuously explore
the cytoplasm and establish meaningful connections, for example
to the kinetochores of mitotic chromosomes (Howard &
Hyman, 2009). Microtubule array reorientations depend on this
dynamic behaviour (Sambade et al., 2012; Lindeboom et al.,
2013).
Plant microtubules are populated by myriads of microtubule-
associated proteins. Microtubule-associated proteins (MAPs)
facilitate and regulate the diverse functions that microtubules have
in plants (Buschmann & Lloyd, 2008). Tasks fulfilled by MAPs
include the regulation of microtubule end dynamics, microtubule
nucleation, bundling, reorientation and severing, intracellular
vesicle transport and the interaction with other physical entities,
including F-actin, kinetochores and membranes. MAPs are
structurally diverse and do not belong to a single protein family
defined by homology. But they do have in common a microtubule
interaction domain (or several of these), which is frequently
characterized by positively charged amino acids (Buschmann et al.,
2007). This allows contact with the acidic surface of microtubules
based on electrostatic interaction. Plants have evolved several MAP
families not present in animals, fungi or protozoa, and it is assumed
that hundreds if not thousands of proteins associate with micro-
tubules in vascular plants (Korolev et al., 2005; Hamada et al.,
2013; Derbyshire et al., 2015).
It is now widely accepted that microtubules control cellulose
microfibril orientation in the cell wall; however, there appears to be
a feedback mechanism at work that transfers information con-
cerning the anisotropy of the cell wall back to the microtubule
array. Every turgescent cell experiences stress, and the direction of
maximal stress depends on cell shape, which (as explained above) is
influenced by the alignment of cellulose microfibrils. Generally,
cortical microtubules align with the direction of dominant stress
(Fischer & Schopfer, 1998; Hejnowicz et al., 2000). This was later
confirmed through physically manipulating the stress pattern
experienced by epidermal cells of the Arabidopsis shoot apical
meristem (Hamant et al., 2008). When cortical microtubules align
according to stress patterns, they will orient cellulose microfibrils in
the same directions, allowing the cells to resist the stress. Unless this
feedback is interrupted by a symmetry breaking event, cells will
therefore grow in a direction perpendicular to the main stress axis,
reinforcing the stress pattern (Uyttewaal et al., 2012). Precisely how
the cortical microtubule cytoskeleton is informed concerning stress
patterns at the cell’s surface remains unknown; however, it is now
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clear that microtubule severing by KATANIN plays an important
role here. Upstream signaling may involve the cell wall integrity
pathway and mechanosensitive ion channels, including calcium
channels (Hamant & Haswell, 2017).
IV. Artificial helical growth exhibited by plant mutants
Mutation-induced helical growth has mainly been investigated and
discussed on the basis of the model plant Arabidopsis. But can we
expect to see similar features in plants with diverse genetic
backgrounds, or even in large, woody plants? Before we discuss this,
we should perhaps define what is considered ‘helical growth’ here.
We consider helical growth as distinct from growth th at has a strong
random component, as seen, for example, in famous garden plants
like Corylus avellana ‘Contorta’ (the corkscrew hazel) or Salix
matsudana ‘Tortuosa’ (corkscrew willow) (Tangchun et al., 2018).
According to our definition, helical growth shows a recognizable
helical pattern which one can easily characterize as being left-
handed or right-handed (even though both forms may be expressed
in a single plant, or even organ). Whether these varieties are caused
by stable mutations (natural or induced) or perhaps by epigenetic
modification is not relevant here. A variety of Pinus strobus (white
pine) named ‘Contorta’ grows into large trees while showing clearly
helical features. Kr€ussmann (1983) reports that it shows twisted
stems and branches. We have investigated a large specimen in the
Arnold Arboretum (Boston, MA, USA), and it is clear that its
needles are strictly right-handed (Fig. 1c,d). The Sugi tree
(Cryptomerica japonica) also shows helical growth in some of its
varieties. A strong growing tree in the B€urgerpark (Osnabr€uck,
Germany), most likely the variety ‘Rasen’, shows left-handed and
right-handed segments in its branches. Within those segments all
the needles follow a given handedness until, perhaps in the next
segment of the same branch, the handedness may change again
(data not shown). These observations suggest that mutation-
induced helical growth is not restricted to inbred lines, nor is it
restricted to a specific taxonomic group or to plants with rather
small organs, like those of Arabidopsis.
Arabidopsis wild-type does not show obvious helical growth,
except for the mild left-handed twist of agar-grown roots and
etiolated hypocotyls seen in certain ecotypes (Simmons et al., 1995;
Migliaccio & Piconese, 2001). However, mutant screens have
revealed plants that show pronounced helical organ growth under
normal growth conditions. Most effective were screens using
slanted agar plates, because this easily demonstrates variations in the
root growth vector, usually referred to as root skewing (Rutherford
et al., 1998). Most, if not all, skewing mutants show abnormal root
twisting (Ishida et al., 2007b; Oliva & Dunand, 2007; Vaughn
et al., 2011; Roy & Bassham, 2014). But other morphological
screens focusing, for example, on leaf development, also uncovered
genes involved in controlling growth direction (Fig. 1e,f;
Buschmann et al., 2004). Eventually helical growth was observed
in rice (Sunohara et al., 2009; Cheng et al., 2017) and the analysis
of herbicide resistance revealed helical growth in the monocot
Lolium rigidum (Chu et al., 2018). Table 1 lists detailed informa-
tion concerning plants showing helical growth as mentioned in this
review.
V. The mechanics behind twisting: from cells to
organs
The Arabidopsis mutants show different expressions of the helical
growth phenotype. Many mutants show strictly left-handed or
right-handed growth, depending on the allele. This is perhaps the
most simple situation, even though the strength of the phenotype
may vary from organ to organ (following, for example, an
ontogenetic series). In some mutants there is organ specificity in
a sense that certain organs show one type of handedness, whereas
other organs show the other. This is seen in csi1 and wave-
dampened2-1 (wvd2-1) plants (Yuen et al., 2003; Bringmann et al.,
2012b), but this condition seems to be rather rare. A different class
of twisters shows right-handed and left-handed torsions alternating
in a fairly unpredictable manner, which is seen, for example, in
Arabidopsis lopped and tornado (trn) mutants (Carland & McHale,
1996; Cnops et al., 2000), as well as in Arabidopsis twisted dwarf1
(twd1)andstrubbelig (sub) (Perez-Perez et al., 2001; Geisler et al.,
2003; Vaddepalli et al., 2011). In the beginning it was not clear
whether helical growth in organs of Arabidopsis mutants is based on
the twisting of individual cells or whether helical growth is the
consequence of rather sophisticated cellular interactions within
organs. One idea was that helical growth is derived from a decrease
in longitudinal cell expansion of inner tissues as compared to outer
tissues, which would force outer tissues to tilt (Hashimoto, 2002).
Another idea was that altered division patterns in meristems (which
are actually helical by default in roots –this would require some
compensatory mechanism in the wild-type (Baum & Rost, 1996;
Wasteneys & Collings, 2004)) are responsible for helical organ
growth. The issue was solved, at least for the mutant tortifolia2
(tor2). In tor2 helical organ growth can be traced back to the
twisting of individual cells, as it is seen in the single-celled leaf
trichomes (Fig. 3a,b) as well as in suspension cell lines derived from
the same mutant background (Buschmann et al., 2009). Recently it
was found that a mutant defective in UDP-L-rhamnose synthetase
(termed rhm1), which results in altered pectin composition, shows
left-handed helical growth in petals (tortifolia2 also shows twisted
petals, though right-handed; Fig. 3c). Close examination of the
rhm1 petals showed that epidermal cells exhibit twisting of the
cellular dome that protrudes out of the plane of the organ, while this
not seen in the wild-type (Saffer et al., 2017). This indicates that the
twisting of individual cells can translate into the twisting of entire
organs (see also Schulgasser & Witztum, 2004; Verger et al., 2019),
but it also suggests that models describing single cell twisting may
serve as a proxy through which we can improve our understanding
of helical growth as a whole.
Mechanistically most important in relation to the mutation-
induced helical growth of model plants like Arabidopsis, therefore,
is the naturally occurring twisted growth of charophycean algae (the
giant cells of Nitella were the main subject of study; Fig. 1b shows
the similar Nitellopsis obtusa) and of the sporangiophore of the
fungus Phycomyces. These organisms were used as early models of
growth anisotropy and helical growth in particular (Roelofsen,
1965). The research showed that the helical growth of these cells is
based on diffusely growing cells that exhibit a helical arrangement
of microfibrils in the wall. Because growth typically occurs
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Table 1. Plant mutants, varieties and transgenics with helical growth phenotypes as mentioned in this review.
Mutant, variety or
transgene Homology Plant species Cellular component Handedness Helical phenotype seen in*Miscellaneous References
Contorta Unknown Pinus strobus Unknown R Needles, branches Kr€
ussmann (1983)
DC 2.15 Proline-rich Daucus carrota Cell wall n/a Leaves Antisense Holk et al. (2002)
Hong Mang Mai Unknown Triticum
aestivum
Unknown R First internode Mediated by
gibberellic acid
Chen et al. (2003)
kegne a-tubulin isotype Eragrostis tef Microtubule R Leaves, coleoptiles tua
T198I
Jost et al. (2015)
‘R’ a-tubulin isotype Lolium rigidum Microtubule R Leaves tua
R243M
Chu et al. (2018)
Rasen Unknown Cryptomeria
japonica
Unknown L/R alt. Needles, branches This paper
SUN IQ67 domain family Solanum
lycopersicum
Microtubule L? Leaves oe Wu et al. (2011)
tid1 a-tubulin isotype Oryza sativa Microtubule R Leaves, stem tua
T56I
Sunohara et al. (2009)
WINDING1 Bric-a-Brac/NPH3 O. sativa Plasma membrane L/R alt. Leaves oe Cheng et al. (2017)
agb1 Heterotrimeric G protein Arabidopsis
thaliana
Endo-membrane suppr. Roots Propyzamide
dependent
Pandey et al. (2008)
ark2 Kinesin A. thaliana MAP R Roots Sakai et al. (2008)
aux1 Transporter, APC group A. thaliana Plasma and endo-
membrane
L Roots May be R in
hypocotyls
Mirza (1987), Okada &
Shimura (1990)
cmu (1,2) Kinesin-light-chain
related
A. thaliana Microtubule L Etiolated hypocotyls Double mutant Liu et al. (2016)
cob GPI-anchored A. thaliana Cell wall L Roots Yuen et al. (2005)
csi1/pom2 ARM repeats A. thaliana MAP L/R org. Roots, etiolated hypocotyls,
inflorescence stems, leaves
Bringmann et al. (2012b)
eb1 (a,b,c) End Binding1 family A. thaliana MAP L Roots Triple mutant Bisgrove et al. (2008)
fer Receptor-like kinase A. thaliana Plasma membrane R Roots Shih et al. (2014)
IQD11 IQ67 domain family A. thaliana Microtubule L Petioles oe Burstenbinder et al. (2017)
IQD16 IQ67 domain family A. thaliana Microtubule L Petioles oe (Burstenbinder et al. (2017)
IQD18 IQ67 domain family A. thaliana Microtubule R Petioles oe (Wendrich et al., 2018)
MAP18 DREPP family A. thaliana Microtubule L Roots oe C. Wang et al. (2007), X. Wang
et al. (2007)
mik2/lrr-kiss Receptor-like kinase A. thaliana Plasma membrane L Roots Van der Does et al. (2017)
mor1 XMAP215 A. thaliana MAP L Roots, hypocotyls Conditional mutant Whittington et al. (2001)
mur8 Unknown A. thaliana Cell wall L Roots Rhamnose
decreased
Saffer et al. (2017)
Murine MAP4 Murine MAP4 A. thaliana Microtubule R Roots, hypocotyls, leaves oe Hashimoto (2002)
rcn1 Subunit A of PP2A A. thaliana Plasma membrane R Roots Deruere et al. (1999)
rhd3 GTP-binding A. thaliana Endo-membrane suppr. Roots Yuen et al. (2005)
rhm1 UDP-L-rhamnose
synthetase
A. thaliana Cell wall L Roots, petals and their epidermal cells Saffer et al. (2017)
sku5 GPI-anchored,
multicopper oxidase
A. thaliana Cell wall L Roots Sedbrook et al. (2002)
sos1 Na+/H+ antiporter A. thaliana plasma membrane suppr. Roots Shoji et al. (2006)
spr1/sku6 Plant-specific A. thaliana MAP R Roots Furutani et al. (2000)
sub Receptor-like kinase A. thaliana Plasma membrane L/R alt. Inflorescence stems, carpels, leaves,
petals
Chevalier et al. (2005)
tch2 Calmodulin-like 24 A. thaliana Microtubule L Roots Missense allele
cml24-4
Wang et al. (2011)
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Table 1. (Continued)
Mutant, variety or
transgene Homology Plant species Cellular component Handedness Helical phenotype seen in*Miscellaneous References
tch2 Calmodulin-like 24 A. thaliana Microtubule R Roots Missense allele
cml24-2
Wang et al. (2011)
tor1/spr2 HEAT repeats A. thaliana MAP R Roots, hypocotyls, inflorescence
stems, leaves
Buschmann et al. (2004)
tor2 a-tubulin isotype A. thaliana Microtubule R Roots, hypocotyls, leaves, trichomes,
petals
tua4
R2K
Buschmann et al. (2009)
trn1 Plant-specific A. thaliana Unknown L/R alt. Roots, inflorescence stems, leaf
petioles, carpels, petals
Allelic with lopped Carland & McHale (1996)
trn2 TETRASPANIN A. thaliana Plasma membrane L/R alt. Roots, inflorescence stems, leaf
petioles, carpels, petals
Cnops et al. (2000)
tua a-tubulin isotype A. thaliana Microtubule L Roots e.g. tua6
S180F
(left-
y1)
Thitamadee et al. (2002)
tua a-tubulin isotype A. thaliana Microtubule R Roots e.g. tua6
S277F
Ishida et al. (2007a)
tub b-tubulin isotype A. thaliana Microtubule L Roots e.g. tub4
T178I
Ishida et al. (2007a,b)
tub b-tubulin isotype A. thaliana Microtubule R Roots e.g. tub3
P287L
Ishida et al. (2007a,b)
twd1/ucu2 FKBP-type
immunophilin
A. thaliana Plasma membrane L/R alt. Roots, hypocotyls, stems, carpels Defective auxin
flow
Perez-Perez et al. (2001)
wvd2-1 TPX2 family A. thaliana Microtubule L/R org. Roots, hypocotyls, leaves oe (activation tag) Yuen et al. (2003)
The table additionally presents some known suppressors of helical growth. It first shows examples from plants other than Arabidopsis thaliana (alphabetically) and then Arabidopsis itself (also
alphabetically). L, left-handed; R, right-handed; alt., L and R alternating in individual organs; org., L or R depending on organtype; suppr., suppressor of helical growth; oe, overexpressor; *additional
organs or cell types may be affected.
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perpendicular to the arrangement of microfibrils, cells with left-
handed microfibrils will show right-handed growth, and vice versa
(Green, 1954, 1962). The steepness of the angle of helical growth,
in a quantitative manner, is therefore related to the steepness of
microfibril alignment (Fig. 1i). Interestingly, such a correlation has
also been suggested for Arabidopsis (Ishida et al., 2007a), though
this was done for cells growing in a tissue context and analyzing
cortical microtubules rather than cellulose microfibrils themselves.
Details of the mechanism of single cell twisting were presented in a
recent paper (Wada, 2012).
VI. Evidence for microtubules having a profound role
in helical growth
The molecular-genetic pathways involved in mutant helical growth
primarily point towards microtubule function (Table 1). Many
mutants were found to exhibit dominant-negative missense
mutations of a-tubulin or b-tubulin genes. The mutations were
predicted to affect the GTPase function of tubulin or, in other cases,
the interaction surface between tubulin subunits when inserted into
the microtubule (Ishida et al., 2007a; Buschmann et al., 2009). In
addition, helical growth significantly contributed to the identifi-
cation of novel plant MAPs. Eventually, it was found that
knocking out MAPs tends to result in helical growth, as
observed for TOR1/SPR2, SPIRAL1 (SPR1), EB1 (END
BINDING1), MOR1 (MICROTUBULE ORGANIZATION1),
ARK2 (ARMADILLO REPEAT KINESIN2) (Whittington et al.,
2001; Buschmann et al., 2004; Nakajima et al., 2004; Bisgrove
et al., 2008; Sakai et al., 2008) and others. Still, there are mutants
with microtubule-related functions that have no clear helical growth
phenotype, for example mutants of KATANIN or mutants of FASS
(neither weak nor strong alleles). But helical growth can also be
found in mutants of genes with functions that are auxin-related: the
aux1 mutant shows left-handed helical growth in roots (Mirza,
1987; Okada & Shimura, 1990) and the roots curl in NPA (rcn1)
mutant has a tendency for right-handed root growth (Garbers et al.,
1996; Deruere et al., 1999). Importantly, several helical growth
mutants point to the cell wall (see sections XI and XII).
What is the reason for microtubules having such a profound role
in helical growth? In roots of Arabidopsis wild-type (and in maize)
it was found that during late cell expansion cortical microtubule
arrays rotate out of the transverse and that these array reorientations
occur with consistent handedness (Liang et al., 1996; Sugimoto
et al., 2000). It was speculated that these innate features of
handedness of the wild-type may explain the mild helical growth
seen in certain Arabidopsis ecotypes (Vaughn et al., 2011).
Importantly, comparable observations were made using the
(usually more striking) helical mutants: these tend to show
pronounced microtubule alignment defects involving helical
configurations (left- or right-handed) rather than transverse
microtubules (Fig. 1e,f; (Furutani et al., 2000; Thitamadee et al.,
2002)). The microtubule orientation defect becomes visible early
during cell expansion and in some reports even before the onset of
morphological twisting (Yuen et al., 2003; Buschmann et al., 2004;
Ishida et al., 2007a). As microtubules are known to have a function
in directing cellulose microfibril alignment, it was assumed that
these mutants would also possess helical cell walls, which in
consequence would lead to twisted growth (Hashimoto, 2002;
Lloyd & Chan, 2002). Surprisingly, helical cellulose microfibrils
were only reported in one case so far (for the mutant csi1/pom2)
(Landrein et al., 2013). This scarcity of data leaves room for
interpretation, but it may for now be taken as support for helical cell
walls being present in Arabidopsis helical growth mutants.
However, it is by no means clear why the microtubules should
adopt helical orientations in the first place.
Several investigations showed that there is a strong correlation
between alterations in the dynamics of cortical microtubules and
helical growth, even though the mechanistic implications are not
obvious. Arabidopsis mutants with right-handed growth frequently
show microtubule dynamics that are consistent with microtubule
stabilization (Ishida et al., 2007a; Korolev et al., 2007; Buschmann
et al., 2009). Likewise, wild-type plants overexpressing murine
MAP4 coupled to GFP tend to show right-handed growth. MAP4
expression is believed to stabilize microtubules in plants
(Hashimoto, 2002). On the other hand, mutants with destabilized
microtubules tend to produce left-handed growth (Furutani et al.,
2000; Thitamadee et al., 2002). The latter is also suggested by the
application of low doses of destabilizing drugs to Arabidopsis. It is
noteworthy that there are exceptions to these general rules: taxol,
for example, a microtubule stabilizing drug, also results in left-
(a) (b)
(c)
Fig. 3 Single-cell twisting and petal phenotype of the Arabidopsis tor2
mutant. (a) Wild-type and (b) tortifolia2 (tor2) leaf trichomes. Trichomes of
tor2 have a right-handed twist and are also larger on average. (c) Twisted
flowers of tor2. The images were originally published in: (a) Sambade et al.
(2014); (b) Buschmann et al. (2009).
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handed growth (some insight regarding the taxol effect is presented
in sections X, XI and XII). The results from studies of microtubule
dynamics in helical growth mutants seem to suggest that micro-
tubules are capable of controlling their own alignment through
microtubule stability. A well justified model for how this works is
currently not at hand, but the sections below (sections X and XII)
on the cell wall and on calcium provide some hints.
Another attempt to explain the role of microtubules in twisted
growth was based on the assumption of an altered protofilament
number in mutant microtubules. Abnormal microtubules with 12
or 14 protofilaments have helical protofilaments producing a
microtubule with a supertwist (Ishida et al., 2007b; Pampaloni &
Florin, 2008). Cortical arrays built from such microtubules could
potentially tilt and assume left-handed or right-handed orienta-
tions. However, transmission electron microscopy revealed only 13
protofilament microtubules in Arabidopsis twisters carrying tubu-
lin missense mutations, and the question of whether the respective
microtubules had a supertwist remained unresolved (Ishida et al.,
2007a). It therefore remained unclear whether microtubules
themselves represent the structural basis of handedness.
Recent data suggest that left-handed growth may be explained
through destabilized microtubules having a reduced capacity for
the direction of the movement of cellulose synthases in the plasma
membrane: the mutants csi1/pom2 and cmu1 cmu2, which were
reported to affect the functional linkage between microtubules and
cellulose synthase (Bringmann et al., 2012b; Liu et al., 2016), both
show left-handed growth (note, however, that apparently there is a
mild right-handed twist in csi1/pom2 stems (Landrein et al.,
2013)). This could mean that cell wall assembly under conditions
of reduced guidance of the cellulose synthase enzyme exerts a
certain intrinsic handedness (i.e. leading to left-handed growth),
which is normally ‘overwritten’ by functional microtubules.
VII. Insight from epistasis
That the cell wall or the process of cell wall assembly might impose a
certain handedness was speculated previously based on results
obtained from double mutant analyses using helical growth
mutants of Arabidopsis. These analyses showed that left-handed
growth in many cases is dominant (epistatic) over right-handed
growth (e.g. Buschmann, 2002; Thitamadee et al., 2002). Similar
results were obtained when treating right-handed mutants with
anti-microtubule drugs (Furutani et al., 2000). If one assumes that
left-handed growth is a direct consequence of the handedness
imposed upon the cell by some property of the cellulosic wall (and
independent of microtubules per se), then it follows that left-
handedness should be epistatic (Buschmann, 2002). But again,
there are exceptions to this pattern (e.g. Galva et al., 2014), and it is
clear that a complete theory of helical growth must accommodate
these exceptions. So what can we really learn from epistasis? It is
convenient to think of left and right as opposing forces that in an
ideal world are in complete balance. In normal organ expansion
that would lead to straight growth. A double mutant created from
left-handed and right-handed twisters should therefore restore
balance once again (or should be at least close to it). In many cases
this does not hold true. Instead we observe epistasis (concerning
handedness –sometimes left, sometimes right) and often exagger-
ation of the twisting itself (Buschmann, 2002; Galva et al., 2014).
This shows that left and right are not separate pathways that are
normally counteracting each other, but that they are intercon-
nected and that the direction of growth is a sophisticated result of
different aspects of microtubule function.
Further epistatic interactions were reported to result in the
suppression of twisting. Mutants of ROOT HAIR DEFECTIVE3
(RHD3), a gene that was speculated to function in plasma
membrane-directed vesicle trafficking, suppressed wild-type root
skewing (Arabidopsis No-0 naturally shows mild root skewing
when grown on slanted agar plates) and, additionally, the left-
handed growth response exerted by the anti-microtubule drug
propyzamide (Yuen et al., 2005). A mutation in the large G protein
AGB1 also suppresses the helical-growth inducing effect of anti-
microtubule drugs (Pandey et al., 2008). Similar to No-0,
Landsberg erecta roots tend to show mild twisting, and this is
suppressed by ethylene (Buer et al., 2003). Moreover, it was
reported that gadolinium ions suppress the helical growth of
hypocotyls of Arabidopsis tubulin mutants. This was true at norma l
gravity (1 g) and hypergravity (300 g) and for both left-handed and
right-handed twisters (Matsumoto et al., 2010). It was shown
previously that gadolinium is a potent blocker of calcium channels
in plants (Sivaguru et al., 2003), and this is consistent with these
channels having a role in mechanosensing. Interesting in this regard
is the finding that organ twisting of the climbing vine Bryonia dioica
is sensitive to blocking calcium signals. In this system tendril coiling
is fully suppressed by the application of gadolinium (Klusener et al.,
1995).
VIII. The microtubule plus tip and helical growth
Some of the MAPs that were found to have a role in helical growth
are microtubule plus tip localized. This is true for EB1 and SPR1,
for example, while TOR1/SPR2 is enriched at the plus tip and also
at several other locations on the cortical array (Galva et al., 2014;
Fan et al., 2018; Nakamura et al., 2018). It is quite likely that these
MAPs have a function in microtubule dynamics, and so it is
difficult to distinguish between a general role in microtubule
stability as opposed to a specific role at the microtubule plus tip. In
this sense, a specific role at the plus tip could involve interaction
with polarity determinants, for example plasma membrane-
localized receptors or ion channels. There is evidence that
microtubules and EB1 regulate calcium influx into the cytosol of
animal cells (Grigoriev et al., 2008; Pozo-Guisado et al., 2013).
Even though the hypothesis of a specific plus end function in helical
growth is attractive, there is only indirect support so far. There has
been speculation that some of the tubulin mutations carried by
helical growth mutants affect the GTPase function of tubulin, and
thereby the microtubule plus end. It has been noted that (in such
right-handed twisters) the size of the observable EB1 comet is
increased (Abe & Hashimoto, 2005; Buschmann et al., 2009). A
larger comet is likely to show altered interaction dynamics with the
proposed polarity determinant. Furthermore, and as mentioned in
section VI, the microtubule drug taxol, perhaps surprisingly, causes
left-handed growth rather than right-handed growth. Interestingly,
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taxol treatment is known to abolish EB1 association with the
microtubule plus tip, at least in cultured animal cells (Morrison
et al., 1998). This may explain why microtubule-stabilizing taxol
does not produce right-handed growth. And there is also the
puzzling observation that the two right-handed mutants spr1 and
tor1/spr2 differ in their response to salt stress: while 50 mM NaCl
can restore the spr1 root growth defect to wild-type, this does not
affect the tor1/spr2 root growth vector (Shoji et al., 2006). Is this
because only SPR1 function is limited to the microtubule plus tip?
IX. Helical growth and the salt stress syndrome
While destabilized microtubules have been reported to result in
left-handed root growth in Arabidopsis, here comes an apparent
exception. Growth medium containing NaCl can produce right-
handed root growth (right-handed twisting) in wild-type Ara-
bidopsis (C. Wang et al., 2007). This is accompanied by micro-
tubule depolymerization, even though at non-lethal NaCl
concentrations microtubule arrays recover after several hours. This
process of microtubule disassembly and recovery is actually
required for surviving the salt stress. The salt overly sensitive1 (sos1)
mutant of Arabidopsis –which accumulates intracellular sodium
due to a mutation in an Na
+
/H
+
antiporter and normally shows
fairly straight roots –responds (to the same amount of NaCl in the
medium) with increased right-handed growth and enhanced
microtubule perturbation (C. Wang et al., 2007). Interestingly,
in another study (and using slightly different plant growth
conditions), sos1 mutations were reported to be suppressors of
the right-handed root twisting of spiral1. When sos1 (and sos2)
mutants were grown on microtubule drugs taxol, propyzamide or
oryzalin, they showed right-handed growth, while the same drugs
produce left-handed growth in the wild-type (Shoji et al., 2006).
Taken together, these studies suggest that salt stress can invert the
innate handedness of Arabidopsis root growth. How can that be? A
role for sodium ions in twisted growth may stem from its capacity to
activate phospholipase D, which is known to bind and potentially
stabilize microtubules (Gardiner et al., 2001). Recently, a role for
phospholipase D was suggested in the salt-stress-induced formation
of phosphatidic acid, which in turn activated MAP65, resulting in
the stabilization of microtubules (Zhang et al., 2012). While this
could lead to twisted growth indirectly, it seems rather likely that
the effect of salt stress on handedness results from calcium signals
cytoplasm
cell wall
oriented and dynamic Microtubules
CesA
MAPs
MAPs
Light
SKU5 COB RA
?
?
?
?
oriented and twisted Cellulose Microfibrils
auxin
gibberellic acid
ethylene
Pectin
(RHM1; MUR8)
Phosphorylation
and Ca2+ influx
Na+
(salt stre ss)
FERONIA
MIK2/ KISS
STRUBBELIG
Receptor?
Fig. 4 A two-way mechanism controls the handedness of cell expansion. The arrows indicate routes of information flow and may only in certain cases represent
actual physical interaction. Cell expansion is dependent on cellulose microfibril alignment; however, microfibril alignment requires a feedback loop involving
cytoplasmic and apoplastic players. The cell wall integrity pathwayis deeply involved in controlling the handedness of cell expansion, and this is probably relayed
by phosphorylation and calcium signals.Cortical microtubule alignment is controlled on several levels. It is speculated that microtubules additionally controltheir
own alignment, perhaps by influencing the same signals that are elicited by cell wall integrity signaling (red arrows). The handedness of cell expansion is also
influenced by phytohormone signaling, but how this works is currently unclear. Twisted cellulose microfibrils may also impact on handedness, but this is highly
speculative at the moment. It should be noted that STRUBBELIG is an atypical (inactive) protein kinase. How the function of specific cell wall proteins likeCOBRA
and RHM1 (RHAMNOSE BIOSYNTHESIS1) are integrated into the process is currently unclear. The processes depicted are derived from investigations involving
various organs (mainly roots, but also stems, leaves and petals) and it may therefore be necessary to demonstrate that a process of interest is really active in a
given organ or even cell.
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affecting the direction of cell expansion (Fig. 4). It is known that an
increase of intracellular sodium will result in the release of a calciu m
signal, which also triggers the salt overly sensitive pathway (Ji et al.,
2013). The effect on calcium channels may be due to salt-induced
microtubule depolymerization. Microtubule depolymerization has
been reported to activate such channels (Thion et al., 1996, 1998).
According to this view the salt stress induced calcium signal would
divert from the SOS pathway and affect the handedness of cell
expansion, possibly through an effect on microtubules (see next
section).
X. The emerging role of calcium-mediated signaling in
twisted growth
Calcium serves as an important second messenger in plants, with
established roles in cell division and cell expansion, gravipercep-
tion, salt stress and cold stress (Hepler, 2016; Lazzaro et al., 2018;
Kolling et al., 2019). These effects will involve, at least in part, an
impact on microtubule function (Wang & Nick, 2017). Cell wall
integrity signaling and mechanosensing also involve calcium fluxes
(Monshausen & Haswell, 2013; Voxeur & Hofte, 2016). In the
above discussion on helical growth, calcium was mentioned
occasionally, and perhaps most significant is the ability of
gadolinium ions to suppress helical growth in Arabidopsis mutants
and in Bryonia (Klusener et al., 1995; Matsumoto et al., 2010). Is
there more evidence to support a role for this messenger in twisted
growth? Two separate point mutations in the calmodulin-like
touch gene TCH2 (CML24) result in Arabidopsis plants with
altered root growth vectors and defects in cell file rotation.
Importantly, the observed phenotypes were shown to involve an
effect on microtubules (Wang et al., 2011). Overexpression of
isoforms 11 and 16 of the plant IQD family of proteins results in a
set of developmental defects and in left-handed petiole growth of
Arabidopsis (Burstenbinder et al., 2017). Several IQD proteins
localize to microtubules and are known to interact with calmod-
ulin. IQD isoform 1 also interacts with CMU, whose inactivation
results in left-handed helical growth (Burstenbinder et al., 2013;
Liu et al., 2016). A recent paper reports that Arabidopsis IQD18
interacts with TOR1/SPR2, and this interaction was modulated by
the addition of calcium in vitro. The overexpression of IQD18
tagged with GFP resulted in right-handed helical petiole growth in
Arabidopsis (Wendrich et al., 2018). In addition, overexpression of
IQD isotype SUN of tomato leads to twisted growth in tomato
plants (Wu et al., 2011). Finally, overexpression of another protein,
MAP18, which belongs to a class of proteins known to interact with
calcium and calmodulin, was reported to result in left-handed
helical root growth in Arabidopsis. However, knock-down plants
of MAP18 were reported not to result in helical growth (X. Wang
et al., 2007; Kolling et al., 2019).
Taken together, these results seem to support a role for calcium
in plant helical growth (Fig. 4). However, when dealing with
overexpression one should consider that artificial expression of a
MAP is likely to influence the dynamics of cortical microtubules in
some way, which could lead to helical growth indirectly. It is
interesting, therefore, to consider a related process in another
organism. Hyphae of the fungus Candida grow in a sinusoidal and
helical manner when cultivated on horizontal agar plates. It was
shown that calcium is important for this touch-dependent growth
behavior, as suppression of calcium influx into the cytosol (using
calcium channel mutants) as well as interfering with calcium
signaling resulted in the attenuation of the twisted growth behavior
(Brand et al., 2009).
XI. What is the significance of twisted cell wall
microfibrils?
Could it be that the cell wall itself exerts some degree of handedness?
Handedness intrinsic to the wall could stem, for example, from the
cellulose microfibrils themselves (Fig. 2), which according to some
reports, including modeling approaches, show twisting (Fernandes
et al., 2011). If such twisted microfibrils are wrapped around a
cylindrical cell, cell elongation may inevitably result in helical
growth, even if the initial orientation was transverse (Landrein
et al., 2013). Indeed, microfibrils were frequently observed to be
twisted, and there seems to be a tendency towards the twist being
right-handed (Kannam et al., 2017), but early investigations also
found left-handed microfibrils (Ruben et al., 1989). Evaluating the
possible impact of twisted microfibrils on helical growth may
require a distinction to be made between different types of cellulose
and between primary vs secondary cell walls. Assuming that the
microfibrils are twisted, it was speculated previously that the
cellulose synthase complexes should rotate (Fig. 2; Somerville,
2006). Interestingly, the green alga Micrasterias is also reported to
have right-handed cellulose microfibrils in its (secondary) cell walls
(Hanley et al., 1997). The alga is closely related to land plants (as
part of the Zygnematophyceae), and its secondary wall cellulose is
known to be produced by hexameric rosettes that travel the
membrane in tight assemblies (Giddings & Staehelin, 1991). It
remains unknown whether the cellulose synthase rosettes of
Micrasterias could be rotating while being kept in the strict order
of the assembly.
There is another interesting type of asymmetry related to
cellulose microfibril formation. CesA complexes are known to
travel along microtubules bi-directionally. However, it was found
that CesA phosporylation patterns impact CesA’s capacity for
efficient bidirectional translocation (Chen et al., 2010). In these
studies it was seen that, depending on the direction of movement,
phosphorylation patterns could alter CesA velocity. The upstream
kinases are unknown but were recently speculated to involve those
known from cell wall integrity signaling (Speicher et al., 2018).
Whether the CesA phosphorylation mutants showed helical
growth remained unclear (Chen et al., 2010); nevertheless, the
results suggest that phosphorylation controls the symmetry of the
interaction between microtubules and CesA complexes (see also
model in Fig. 4).
XII. Further evidence for a directional influence of the
cell wall
If the cell wall had an autonomous influence on handedness, one
would expect mutant screens to reveal at least some genes with cell
wall related functions. Indeed, there is some support coming from
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mutants (see also Table 1). The Arabidopsis sku5 mutant has left-
handed twisted roots. SKU5 encodes a GPI-anchored protein with
a supposed in muro function (Sedbrook et al., 2002). Mutants of
COB, which codes for another GPI-anchored protein, are defective
in cellulose biosynthesis and were reported to have (a fairly mild)
aberrant root skewing phenotype (Yuen et al., 2005). A recent
screen for mutants with defective flowers revealed an Arabidopsis
plant with left-handed helical petals. This study was remarkable
also because the research showed that in this case (like in tor2) organ
twisting is rooted in the twisting of single cells. The gene affected
was identified to be RHAMNOSE BIOSYNTHESIS1 (RHM1) and
an additional cell wall gene (MUR8) was shown to be involved as
well. The authors propose that rhamnose-containing cell wall
polymers have a function in suppressing helical cell expansion
(Saffer et al., 2017). Another study (not based on an initial mutant
screen) pointing to an independent function of the cell wall in
helical growth comes from carrots. The study showed that
overexpression of a cell wall protein can induce helical handedness
in leaves (Holk et al., 2002). Finally, there are some recent reports
on Arabidopsis helical growth mutants that point to the cell wall
integrity pathway. Twisting is seen in several organs of strubbelig
(sub) mutants (Chevalier et al., 2005). The twisting direction is
seemingly random, but such a conclusion may require further
investigation. SUB codes for an atypical leucine-rich repeat
receptor-like kinase (Eyuboglu et al., 2007). Right-handed root
skewing (likely indicating a helical growth phenotype) was seen in
feronia (fer) mutants. FER codes for a receptor-like kinase and is
expressed throughout the plant (except in pollen). Mutants of FER
are known to be defective in calcium signaling and mechanoper-
ception (Haruta et al., 2014; Ngo et al., 2014; Shih et al., 2014).
Importantly, the FER receptor was also identified as a pectin
receptor (Feng et al., 2018). A recent study showed that the MIK2/
LRR-KISS gene, another receptor-like kinase, is required for a
normal root growth vector. So-called mik2 mutants show leftward
root skewing and increased salt stress sensitivity. Interestingly,
MIK2/LRR-KIS regulates root growth direction in a CesA6/
isoxaben dependent manner. It is certainly important to mention
that the mik2 mutant, even though its roots are twisted, did not
seem to have a cellulose microfibril orientation defect (Van der
Does et al., 2017). Taken together, the receptor-like kinase mu tants
suggest that in the wild-type, information that arrives from the cell
wall needs to be transmitted to the cytoplasm to facilitate straight
growth (see model in Fig. 4).
XIII. Tropism-induced helical growth and
TORTIFOLIA1/SPIRAL2
Erna Reinholz utilized the striking petiole phenotype of so-called
tortifolia (tor) mutants to determine the rate of spontaneous
mutation in untreated Arabidopsis seeds (Reinholz, 1947a,b).
Albert Kranz introduced tortifolia1-1 (F104; Enkheim2 back-
ground) to NASC in 1992, and the corresponding gene symbol
TOR was registered by Toni Sch€affner in 1993 (B€urger, 1971;
Kranz & Kirchheim, 1987; Mainke & Koornneef, 1997). Even
though the mutant was used in subsequent investigations (Fabri &
Sch€affner, 1994; Serrano-Cartagena et al., 1999), the name of
allelic spiral2 (spr2) (Furutani et al., 2000) became rather popular,
and several recent publications use the name SPIRAL2 only. To
resolve this confusion we suggest here that the community may use
a combined designator such as tor1/spr2.
The tor1/spr2 phenotype is produced by recessive knock-out
mutations. The mutant is fully right-handed and the phenotype is
seen in all major organs. The 94 kDa TOR1/SPR2 protein shows
N-terminal HEAT repeats and a predicted (and phylogenetically
conserved) central coiled-coil domain (Buschmann et al., 2004). In
planta TOR1/SPR2 shows oligomerization. TOR1/SPR2 pre-
sented the first known plant-specific MAP: the protein labeled
microtubules in vitro and the GFP fusion (which complements the
mutant when labeled C-terminally) decorated microtubules in vivo
(Buschmann et al., 2004; Shoji et al., 2004).
A fluorescent protein study by Yao et al., 2008 revealed many
details of the subcellular localization of TOR1/SPR2. The protein
was found to localize to plus ends and to microtubule crossover
sites. Additionally, fluorescent foci were seen to associate with
depolymerizing microtubules (Yao et al., 2008). According to two
recent studies these foci were probably microtubule minus ends,
and TOR1/SPR2 protects microtubules from minus end-derived
depolymerization (Fan et al., 2018; Nakamura et al., 2018).
TOR1/SPR2 binds minus ends even in vitro, suggesting that this
property is intrinsic to TOR1. However, in vitro the protein does
not recognize the plus tip, suggesting that this may be facilitated
through an interacting protein. Interestingly, we have identified a
sequence in TOR1/SPR2 that closely matches the EB1 binding
motif (Honnappa et al., 2009).
A number of studies have explored the microtubule dynamics of
tor1/spr2 in comparison to wild-type (Yao et al., 2008; Wightman
et al., 2013; Fan et al., 2018; Nakamura et al., 2018). This research
was carried out mainly in Arabidopsis, but recently also in
Physcomitrella knock-outs (Leong et al., 2018). Unfortunately, the
Arabidopsis results are not fully congruent. Two recent studies,
however, agree on a set of details (Fan et al., 2018; Nakamura et al.,
2018). TOR1/SPR2 destabilizes the microtubule plus end to some
extent, while the treadmilling minus end is protected from
catastrophe. It is now clear that tor1/spr2 mutants are defective in
microtubule reorientation, be it induced by blue light or by
externally applied auxin (Borchers et al., 2018; Fan et al., 2018;
Nakamura et al., 2018), and that the severing protein KATANIN is
involved (at least in blue-light reorientation). Interestingly, the
protein TOR1/SPR2 appears to function in increasing the
likelihood of KATANIN-mediated severing at microtubule
crossover sites. The papers by Nakamura et al. (2018) and Fan
et al. (2018) suggest that a specific step in building a differentially
oriented microtubule array is defective in tor1/spr2. Accordingly,
the step of amplifying already present discordant microtubules is
abnormal and inefficient in tor1/spr2. This is because increased
minus-end depolymerization removes crossovers frequently,
reducing the chance that severing at crossovers could serve as
amplification points for additional microtubules with a similar
orientation. One study confirmed this aspect of the tor1/spr2 defect
through modeling (Nakamura et al., 2018). The observed pheno-
types of tor1/spr2 may explain at least partially the inability of the
microtubule array to reorient in a timely manner. Such an array
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may therefore be considered ‘stabilized’. It is not evident from these
analyses of microtubule dynamics why the microtubule reorienta-
tion defect should produce helical growth in the mutant.
Recent research shows that the right-handed tor1/spr2 mutant is
defective in helical tropisms of the petiole. In plants, tropisms often
lead to organ bending, and the underlying mechanisms have been
extensively studied in the laboratory (Darwin & Darwin, 1880;
Fankhauser & Christie, 2015). Under natural conditions, however,
tropisms frequently result in organ twisting (i.e. helical growth;
Snow, 1950, 1962). Illuminating wild-type Arabidopsis leaves
from the side leads to helical growth of the petiole, apparently
serving maximum illumination. When tor1/spr2 is exposed to
lateral light it can be seen that the mutant is in principle capable of
twisting its petioles to the left, but this occurs at a lower rate.
However, the mutant is faster (than the wild-type) when moving to
the right. It is noteworthy that the angular velocity of the
constitutive twisting of tor1/spr2 petioles (under vertical light) is
much lower than the tropism-induced twistings in either direction.
The lateral light experiments on tor1/spr2 suggested that the MAP is
somehow involved in light-induced helical tropisms of the petiole.
Further experiments showed that tor1/spr2 mutants are incapable of
performing normal auxin-induced petiole torsions, and the
microtubules of excised tor1/spr2 petioles cannot reorient normally
when stimulated with auxin (Borchers et al., 2018). Taken
together, the results suggest that TOR1/SPR2 is required for auxin
and blue-light induced microtubule reorientations, and they
provide evidence that MAPs and microtubules are important for
helical tropisms of plant organs.
XIV. Conclusions and outlook
Helical plant growth is not an anomaly. Rather, it is shown by many
wild-type species as part as their developmental program (Smyth,
2016) or by wild-type species as part of their response to the
environment (Chen et al., 2003; Bisgrove et al., 2008; Borchers
et al., 2018). In this review we focused on helical growth that is
induced by mutation. We hope that this may contribute to a deeper
understanding of anisotropic growth. However, the discussion of
mutant phenotypes should also help to better understand naturally
occurring helical growth, including helical tropisms. The working
model on helical growth shown in Fig. 4 suggests routes of
information flow (arrows), and only some of the steps indicated
may involve actual physical interaction. Microtubule orientation
undoubtedly plays a major role in helical growth and most (but not
all!) twisting mutants present helical microtubule arrays. The
model suggests that handedness information leaves the symplast
and arrives at the apoplast, and vice versa. Accordingly, this is a two-
way system that functions in adjusting the handedness of cell
expansion. Phosphorylation events (in particular by receptor-like
kinases) and calcium influx together present a hub of information
flow. Other pathways known to intermingle with helical growth
may connect to this hub, for example gravitropism and touch (not
shown). It appears that microtubules can regulate their own
orientation, and this capacity can be mimicked by manipulating
microtubule stability. How this works is unclear; however, it is
speculated here that microtubule stability somehow feeds into the
same hub involving calcium (Thion et al., 1998) and/or phospho-
rylation (these possibilities are indicated by the red arrows in the
figure). Importantly, the actual structural basis of handedness in
helical growth is unresolved. It has been speculated that the
microtubule itself forms that basis, but proof is pending. While it is
likely that the cellulose microfibrils of primary Arabidopsis cell
walls are twisted, it is not clear whether this could affect the
direction of cell expansion. These aspects should be explored in
future experimentation and computational modeling. Further
experiments can reveal the details of cell wall integrity signaling in
helical growth and the spatio-temporal role of calcium in regulating
microtubule behavior and orientation.
The model in Fig. 4 may also help us to identify novel routes for
improving crops. Many helical growth mutants are smaller than the
wild-type. Teff (Eragrostis tef) is an important food crop in eastern
Africa. The plant’s productivity is severely decreased by its tendency
for lodging. Recent research has revealed that a semi-dwarfed teff
plant with right-handed twisted growth (named kegne)carriesa
missense mutation in a-tubulin. Importantly, this plant shows
lodging tolerance (Jost et al., 2015). This effect could be due to plant
size only; however, it was demonstrated that in long, flat leaves
twisting significantlycontributes to mechanical stability (Schulgasser
& Witztum, 2004). That twisting may increase plant organ stability
was also suggested in the case of the deep-sowing tolerant Triticum
aestivum cultivar ‘Hong Mang Mai’. This wheat plant shows an
unusual degree of gibberellic acid sensitivity of the first internode.
The first internode of this cultivar is capable of extreme elongation
growth and also shows right-handed twisting. The developing twist
may assist the plant in pushing its way through the soil (Chen et al.,
2003). Given these examples in teff and wheat, it is probably not too
far-fetched to speculate that manipulating root growth behavior
using twisting pathways could help the breeder to modify and
eventually improve the capacity of crop plants to invade the soil.
Acknowledgements
We are grateful to four anonymous reviewers for their productive
comments. Sabine Zachgo is thanked for helpful discussions and
hosting this research in Osnabr€uck. We are also grateful to Lothar
Krienitz, who was involved in the sampling and classification of
Nitellopsis obtusa. The Botanical Garden in Osnabr€uck is thanked
for providing plant material. We further acknowledge Toni
Sch€affner for his support of tortifolia projects. We acknowledge a
DFG (Deutsche Forschungsgemeinschaft) grant to HB (BU 2301/
2-1) and support by the local SFB944 research consortium.
ORCID
Henrik Buschmann https://orcid.org/0000-0003-3022-6150
References
Abe T, Hashimoto T. 2005. Altered microtubule dynamics by expression of
modified alpha-tubulin protein causes right-handed helical growth in transgenic
Arabidopsis plants. The Plant Journal 43: 191–204.
Ó2019 The Authors
New Phytologist Ó2019 New Phytologist Trust
New Phytologist (2019)
www.newphytologist.com
New
Phytologist Tansley review Review 13

Baskin TI. 2001. On the alignment of cellulose microfibrils by cortical
microtubules: a review and a model. Protoplasma 215: 150–171.
Baskin TI. 2005. Anisotropic expansion of the plant cell wall. Annual Review of Cell
and Developmental Biology 21: 203–222.
Baum SF, Rost TL. 1996. Root apical organization in Arabidopsis thaliana. 1. Root
cap and protoderm. Protoplasma 192: 178–188.
Bisgrove SR, Lee YR, Liu B, Peters NT, Kropf DL. 2008. The microtubule plus-
end binding protein EB1 functions in root responses to touch and gravity signals
in Arabidopsis.Plant Cell 20: 396–410.
Borchers A, Deckena M, Buschmann H. 2018. Arabidopsis petiole torsions induced
by lateral light or externally supplied auxin require microtubule-associated
TORTIFOLIA1/SPIRAL2. Protoplasma 255: 1505–1515.
Brand A, Lee K, Veses V, Gow NA. 2009. Calcium homeostasis is required for
contact-dependent helical and sinusoidal tip growth in Candida albicans hyphae.
Molecular Microbiology 71: 1155–1164.
Bringmann M, Landrein B, Schudoma C, Hamant O, Hauser MT, Persson S.
2012a. Cracking the elusive alignment hypothesis: the microtubule-cellulose
synthase nexus unraveled. Trends in Plant Science 17: 666–674.
Bringmann M, Li E, Sampathkumar A, Kocabek T, Hauser MT, Persson S. 2012b.
POM-POM2/cellulose synthase interacting1 is essential for the functional
association of cellulose synthase and microtubules in Arabidopsis.Plant Cell 24:
163–177.
Buer CS, Wasteneys GO, Masle J. 2003. Ethylene modulates root-wave responses
in Arabidopsis.Plant Physiology 132: 1085–1096.
B€urger D. 1971. Die morphologischen Mutanten des G€ottinger Arabidopsis-
Sortiments, einschließlich der Mutanten mit abweichender Samenfarbe.
Arabidopsis Information Service 8:36–42.
Burstenbinder K, Moller B, Plotner R, Stamm G, Hause G, Mitra D, Abel S. 2017.
The IQD family of calmodulin-binding proteins links calcium signaling to
microtubules, membrane subdomains, and the nucleus. Plant Physiology 173:
1692–1708.
Burstenbinder K, Savchenko T, Muller J, Adamson AW, Stamm G, Kwong R,
Zipp BJ, Dinesh DC, Abel S. 2013. Arabidopsis calmodulin-binding protein
IQ67-domain 1 localizes to microtubules and interacts with kinesin light chain-
related protein-1. Journal of Biological Chemistry 288: 1871–1882.
Burton RA, Gidley MJ, Fincher GB. 2010. Heterogeneity in the chemistry,
structure and function of plant cell walls. Nature Chemical Biology. 6: 724.
Buschmann H 2002. TORTIFOLIA Gene kontrollieren das Streckungswachstum in
Arabidopsis. Klonierung von TORTIFOLIA1. PhD thesis. Fakult€at f€ur Chemie
und Pharmacie. University of Munich LMU, Munich, Germany.
Buschmann H, Fabri CO, Hauptmann M, Hutzler P, Laux T, Lloyd CW,
Sch€affner AR. 2004. Helical growth of the Arabidopsis mutant tortifolia1
reveals a plant-specific microtubule-associated protein. Current Biology 14:
1515–1521.
Buschmann H, Hauptmann M, Niessing D, Lloyd CW, Schaffner AR. 2009.
Helical growth of the Arabidopsis mutant tortifolia2 does not depend on cell
division patterns but involves handed twisting of isolated cells. Plant Cell 21:
2090–2106.
Buschmann H, Lloyd CW. 2008. Arabidopsis mutants and the network of
microtubule-associated functions. Molecular Plant 1: 888–898.
Buschmann H, Sanchez-Pulido L, Andrade-Navarro MA, Lloyd CW. 2007.
Homologues of Arabidopsis microtubule-associated AIR9 in trypanosomatid
parasites: hints on evolution and function. Plant Signaling & Behavior 2: 296–
299.
Carland FM, McHale NA. 1996. LOP1: a gene involved in auxin transport and
vascular patterning in Arabidopsis.Development 122: 1811–1819.
Chen S, Ehrhardt DW, Somerville CR. 2010. Mutations of cellulose synthase
(CESA1) phosphorylation sites modulate anisotropic cell expansion and
bidirectional mobility of cellulose synthase. Proceedings of the National Academy of
Sciences, USA 107: 17188–17193.
Cheng ML, Lo SF, Hsiao AS, Hong YF, Yu SM, Ho TD. 2017. Ectopic expression
of WINDING 1 leads to asymmetrical distribution of auxin and a spiral phenotype
in rice. Plant and Cell Physiology 58: 1494–1506.
Chen L, Higashitani A, Suge H, Takeda K, Takahashi H. 2003. Spiral growth and
cell wall properties of the gibberellin-treated first internodes in the seedlings of a
wheat cultivar tolerant to deep-sowing conditions. Physiologia Plantarum 118:
147–155.
Chevalier D, Batoux M, Fulton L, Pfister K, Yadav RK, Schellenberg M, Schneitz
K. 2005. STRUBBELIG defines a receptor kinase-mediated signaling pathway
regulating organ development in Arabidopsis.Proceedings of the National Academy
of Sciences, USA 102: 9074–9079.
Chu Z, Chen J, Nyporko A, Han H, Yu Q, Powles S. 2018. Novel alpha-tubulin
mutations conferring resistance to dinitroaniline herbicides in Lolium rigidum.
Frontiers in Plant Science 9: 97.
Cnops G, Wang X, Linstead P, Van Montagu M, Van Lijsebettens M, Dolan L.
2000. TORNADO1 and TORNADO2 are required for the specification of radial
and circumferential pattern in the Arabidopsis root. Development 127: 3385–
3394.
Cosgrove DJ. 2018. Nanoscale structure, mechanics and growth of epidermal cell
walls. Current Opinion in Plant Biology 46:77–86.
Darwin C, Darwin F. 1880. The power of movement in plants. London, UK: John
Murray.
Derbyshire P, Menard D, Green P, Saalbach G, Buschmann H, Lloyd CW,
Pesquet E. 2015. Proteomic analysis of microtubule interacting proteins over the
course of xylem tracheary element formation in Arabidopsis.Plant Cell 27: 2709–
2726.
Deruere J, Jackson K, Garbers C, Soll D, Delong A. 1999. The RCN1-encoded A
subunit of protein phosphatase 2A increases phosphatase activity in vivo.The
Plant Journal 20: 389–399.
Edwards W, Moles AT, Franks P. 2007. The global trend in plant twining direction.
Global Ecology and Biogeography 16: 795–800.
Endler A, Kesten C, Schneider R, Zhang Y, Ivakov A, Froehlich A, Funke N,
Persson S. 2015. A mechanism for sustained cellulose synthesis during salt stress.
Cell 162: 1353–1364.
Eyuboglu B, Pfister K, Haberer G, Chevalier D, Fuchs A, Mayer KFX, Schneitz K.
2007. Molecular characterisation of the STRUBBELIG-RECEPTOR FAMILY of
genes encoding putative leucine-rich repeat receptor-like kinases in Arabidopsis
thaliana.BMC Plant Biology 7: 16.
Fabri CO, Sch€affner AR. 1994. An Arabidopsis thaliana RFLP mapping set to
localize mutations to chromosome regions. The Plant Journal 5: 149–156.
Fan Y, Burkart GM, Dixit R. 2018. The Arabidopsis SPIRAL2 protein targets and
stabilizes microtubule minus ends. Current Biology 28: 987–994. e983
Fankhauser C, Christie JM. 2015. Plant phototropic growth. Current Biology 25:
R384–R389.
Feng W, Kita D, Peaucelle A, Cartwright HN, Doan V, Duan Q, Liu MC, Maman
J, Steinhorst L, Schmitz-Thom I et al. 2018. The FERONIA receptor kinase
maintains cell-wall integrity during salt stress through Ca
2+
Signaling. Current
Biology 28: 666–675. e665
Fernandes AN, Thomas LH, Altaner CM, Callow P, Forsyth VT, Apperley DC,
Kennedy CJ, Jarvis MC. 2011. Nanostructure of cellulose microfibrils in spruce
wood. Proceedings of the National Academy of Sciences, USA 108: E1195–E1203.
Fischer K, Schopfer P. 1998. Physical strain-mediated microtubule reorientation in
the epidermis of gravitropically or phototropically stimulated maize coleoptiles.
The Plant Journal 15: 119–123.
Forterre Y, Dumais J. 2011. Materials science. Generating helices in nature. Science
333: 1715–1716.
Furutani I, Watanabe Y, Prieto R, Masukawa M, Suzuki K, Naoi K, Thitamadee S,
Shikanai T, Hashimoto T. 2000. The SPIRAL genes are required for directional
control of cell elongation in Arabidopsis thaliana.Development 127: 4443–4453.
Galva C, Kirik V, Lindeboom JJ, Kaloriti D, Rancour DM, Hussey PJ, Bednarek
SY, Ehrhardt DW, Sedbrook JC. 2014. The microtubule plus-end tracking
proteins SPR1 and EB1b interact to maintain polar cell elongation and directional
organ growth in Arabidopsis.Plant Cell 26: 4409–4425.
Garbers C, DeLong A, Deruere J, Bernasconi P, Soll D. 1996. A mutation in
protein phosphatase 2A regulatory subunit A affects auxin transport in
Arabidopsis.EMBO Journal 15: 2115–2124.
Gardiner JC, Harper JD, Weerakoon ND, Collings DA, Ritchie S, Gilroy S, Cyr
RJ, Marc J. 2001. A 90-kD phospholipase D from tobacco binds to microtubules
and the plasma membrane. Plant Cell 13: 2143–2158.
Geisler M, Kolukisaoglu HU, Bouchard R, Billion K, Berger J, Saal B, Frangne N,
Koncz-Kalman Z, Koncz C, Dudler R et al. 2003. TWISTED DWARF1, a
unique plasma membrane-anchored immunophilin-like protein, interacts with
Arabidopsis multidrug resistance-like transporters AtPGP1 and AtPGP19.
Molecular Biology of the Cell 14: 4238–4249.
New Phytologist (2019) Ó2019 The Authors
New Phytologist Ó2019 New Phytologist Trust
www.newphytologist.com
Review Tansley review
New
Phytologist
14

Giddings TH, Staehelin LA. 1991. Microtubule-mediated control of microfibril
deposition: a re-examination of the hypothesis. In: Lloyd CW, ed. The cytoskeletal
basis of plant growth and form. London, UK: Academic, 85–99.
Green PB. 1954. The spiral growth pattern of the cell wall in Nitella axillaris.
American Journal of Botany 41: 403–409.
Green PB. 1962. Mechanism for plant cellular morphogenesis. Science 138: 1404–
1405.
Grigoriev I, Gouveia SM, van der Vaart B, Demmers J, Smyth JT, Honnappa S,
Splinter D, Steinmetz MO, Putney JW Jr, Hoogenraad CC et al. 2008. STIM1
is a MT-plus-end-tracking protein involved in remodeling of the ER. Current
Biology 18: 177–182.
Hamada T, Nagasaki-Takeuchi N, Kato T, Fujiwara M, Sonobe S, Fukao Y,
Hashimoto T. 2013. Purification and characterization of novel microtubule-
associated proteins from Arabidopsis cell suspension cultures. Plant Physiology 163:
1804–1816.
Hamant O, Haswell ES. 2017. Life behind the wall: sensing mechanical cues in
plants. BMC Biology 15: 59.
Hamant O, Heisler MG, Jonsson H, Krupinski P, Uyttewaal M, Bokov P, Corson
F, Sahlin P, Boudaoud A, Meyerowitz EM et al. 2008. Developmental
patterning by mechanical signals in Arabidopsis.Science 322: 1650–1655.
Hanley SJ, Revol JF, Godbout L, Gray DG. 1997. Atomic force microscopy and
transmission electron microscopy of cellulose from Micrasterias denticulata;
evidence for a chiral helical microfibril twist. Cellulose 4: 209–220.
Harris JM. 1989. Spiral grain and wave phenomena in wood formation. Berlin,
Germany: Springer.
Haruta M, Sabat G, Stecker K, Minkoff BB, Sussman MR. 2014. A peptide
hormone and its receptor protein kinase regulate plant cell expansion. Science 343:
408–411.
Hashimoto T. 2002. Molecular genetic analysis of left-right handedness in plants.
Philosophical Transactions of the Royal Society of London. Series B,Biological Sciences
357: 799–808.
Heath IB. 1974. Unified hypothesis for role of membrane-bound enzyme
complexes and microtubules in plant-cell wall synthesis. Journal of Theoretical
Biology 48: 445–449.
Hejnowicz Z, Rusin A, Rusin T. 2000. Tensile tissue stress affects the orientation of
cortical microtubules in the epidermis of sunflower hypocotyl. Journal of Plant
Growth Regulation 19:31–44.
Hepler PK. 2016. The cytoskeleton and its regulation by calcium and protons. Plant
Physiology 170:3–22.
Holk A, Klumpp L, Scherer GFE. 2002. A cell wall protein down-regulated by auxin
suppresses cell expansion in Daucus carota (L.). Plant Molecular Biology 50: 295–
305.
Honnappa S, Gouveia SM, Weisbrich A, Damberger FF, Bhavesh NS, Jawhari H,
Grigoriev I, van Rijssel FJ, Buey RM, Lawera A et al. 2009. An EB1-binding
motif acts as a microtubule tip localization signal. Cell 138: 366–376.
Howard J, Hyman AA. 2003. Dynamics and mechanics of the microtubule plus
end. Nature 422: 753–758.
Howard J, Hyman AA. 2009. Growth, fluctuation and switching at microtubule
plus ends. Nature Reviews Molecular Cell Biology 10: 569–574.
Ishida T, Kaneko Y, Iwano M, Hashimoto T. 2007a. Helical microtubule arrays in
a collection of twisting tubulin mutants of Arabidopsis thaliana.Proceedings of the
National Academy of Sciences, USA 104: 8544–8549.
Ishida T, Thitamadee S, Hashimoto T. 2007b. Twisted growth and organization of
cortical microtubules. Journal of Plant Research 120:61–70.
Ji H, Pardo JM, Batelli G, Van Oosten MJ, Bressan RA, Li X. 2013. The Salt Overly
Sensitive (SOS) pathway: established and emerging roles. Molecular Plant 6: 275–
286.
Jost M, Esfeld K, Burian A, Cannarozzi G, Chanyalew S, Kuhlemeier C, Assefa K,
Tadele Z. 2015. Semi-dwarfism and lodging tolerance in tef (Eragrostis tef )is
linked to a mutation in the alpha-Tubulin 1 gene. Journal of Experimental Botany
66: 933–944.
Kannam SK, Oehme DP, Doblin MS, Gidley MJ, Bacic A, Downton MT. 2017.
Hydrogen bonds and twist in cellulose microfibrils. Carbohydrate Polymers 175:
433–439.
Kern VD, Schwuchow JM, Reed DW, Nadeau JA, Lucas J, Skripnikov A, Sack FD.
2005. Gravitropic moss cells default to spiral growth on the clinostat and in
microgravity during spaceflight. Planta 221: 149–157.
Kesten C, Wallmann A, Schneider R, McFarlane HE, Diehl A, Khan GA, van
Rossum BJ, Lampugnani ER, Szymanski WG, Cremer N et al. 2019. The
companion of cellulose synthase 1 confers salt tolerance through a Tau-like
mechanism in plants. Nature Communications 10: 857.
Klusener B, Boheim G, Liss H, Engelberth J, Weiler EW. 1995. Gadolinium-
sensitive, voltage-dependent calcium release channels in the endoplasmic
reticulum of a higher plant mechanoreceptor organ. EMBO Journal 14: 2708–
2714.
Kolling M, Kumari P, B€urstenbinder K. 2019. Calcium- and calmodulin-
regulated microtubule-associated proteins as signal-integration hubs at the
plasma membrane-cytoskeleton nexus. Journal of Experimental Botany 70:
387–396.
Korolev AV, Buschmann H, Doonan JH, Lloyd CW. 2007. AtMAP70-5, a
divergent member of the MAP70 family of microtubule-associated proteins, is
required for anisotropic cell growth in Arabidopsis.Journal of Cell Science 120:
2241–2247.
Korolev AV, Chan J, Naldrett MJ, Doonan JH, Lloyd CW. 2005. Identification of
a novel family of 70 kDa microtubule-associated proteins in Arabidopsis cells. The
Plant Journal 42: 547–555.
Kranz AR, Kirchheim B. 1987. Genetic resources in Arabidopsis.Arabidopsis
Information Service 24: 3.3.24.
Kr€ussmann G. 1983. Handbuch der Nadelgeh€olze. Berlin, Hamburg, Germany:
Parey.
Landrein B, Lathe R, Bringmann M, Vouillot C, Ivakov A, Boudaoud A, Persson S,
Hamant O. 2013. Impaired cellulose synthase guidance leads to stem torsion and
twists phyllotactic patterns in Arabidopsis.Current Biology 23: 895–900.
Lazzaro MD, Wu S, Snouffer A, Wang Y, van der Knaap E. 2018. Plant organ
shapes are regulated by protein interactions and associations with microtubules.
Frontiers in Plant Science 9: 1766.
Ledbetter MC, Porter KR. 1963. A “microtubule” in plant cell fine structure.
Journal of Cell Biology 19: 239–250.
Lei L, Li S, Gu Y. 2012. Cellulose synthase complexes: composition and regulation.
Frontiers in Plant Science 3: 75.
Leong SY, Yamada M, Yanagisawa N, Goshima G. 2018. SPIRAL2 stabilises
endoplasmic microtubule minus ends in the moss Physcomitrella patens.Cell
Structure and Function 43:53–60.
Li S, Lei L, Somerville CR, Gu Y. 2012. Cellulose synthase interactive protein 1
(CSI1) links microtubules and cellulose synthase complexes. Proceedings of the
National Academy of Sciences, USA 109: 185–190.
Liang BM, Dennings AM, Sharp RE, Baskin TI. 1996. Consistent handedness of
microtubule arrays in maize and Arabidopsis primary roots. Protoplasma 190:8–
15.
Lindeboom JJ, Nakamura M, Hibbel A, Shundyak K, Gutierrez R, Ketelaar T,
Emons AMC, Mulder BM, Kirik V, Ehrhardt DW. 2013. A mechanism for
reorientation of cortical microtubule arrays driven by microtubule severing.
Science 342: 1202–1205.
Liu Z, Schneider R, Kesten C, Zhang Y, Somssich M, Zhang Y, Fernie AR, Persson
S. 2016. Cellulose-microtubule uncoupling proteins prevent lateral displacement
of microtubules during cellulose synthesis in Arabidopsis.Developmental Cell 38:
305–315.
Lloyd C. 2011. Dynamic microtubules and the texture of plant cell walls.
International Review of Cell and Molecular Biology 287: 287–329.
Lloyd C, Chan J. 2002. Helical microtubule arrays and spiral growth. Plant Cell 14:
2319–2324.
L€owe J, Li H, Downing KH, Nogales E. 2001. Refined structure of alpha beta-
tubulin at 3.5 A resolution. Journal of Molecular Biology 313: 1045–1057.
Mainke D, Koornneef M. 1997. Community standards for Arabidopsis genetics.
The Plant Journal 12: 247–253.
Matsumoto S, Kumasaki S, Soga K, Wakabayashi K, Hashimoto T, Hoson T.
2010. Gravity-induced modifications to development in hypocotyls of
Arabidopsis tubulin mutants. Plant Physiology 152: 918–926.
Migliaccio F, Piconese S. 2001. Spiralizations and tropisms in Arabidopsis roots.
Trends in Plant Science 6: 561–565.
Mirza J. 1987. Spiral growth of hypocotyls in mutant aux-1 of Arabidopsis thaliana.
Arabidopsis Information Service 23:62–63
Monshausen GB, Haswell ES. 2013. A force of nature: molecular mechanisms of
mechanoperception in plants. Journal of Experimental Botany 64: 4663–4680.
Ó2019 The Authors
New Phytologist Ó2019 New Phytologist Trust
New Phytologist (2019)
www.newphytologist.com
New
Phytologist Tansley review Review 15

Morrison EE, Wardleworth BN, Askham JM, Markham AF, Meredith DM. 1998.
EB1, a protein which interacts with the APC tumour suppressor, is associated with
the microtubule cytoskeleton throughout the cell cycle. Oncogene 17: 3471–3477.
Nakajima K, Furutani I, Tachimoto H, Matsubara H, Hashimoto T. 2004.
SPIRAL1 encodes a plant-specific microtubule-localized protein required for
directional control of rapidly expanding Arabidopsis cells. Plant Cell 16: 1178–
1190.
Nakamura M, Lindeboom JJ, Saltini M, Mulder BM, Ehrhardt DW. 2018. SPR2
protects minus ends to promote severing and reorientation of plant cortical
microtubule arrays. Journal of Cell Biology 217: 915–927.
Ngo QA, Vogler H, Lituiev DS, Nestorova A, Grossniklaus U. 2014. A calcium
dialog mediated by the FERONIA signal transduction pathway controls plant
sperm delivery. Developmental Cell 29: 491–500.
Nogales E, Whittaker M, Milligan RA, Downing KH. 1999. High-resolution
model of the microtubule. Cell 96:79–88.
Okada K, Shimura Y. 1990. Reversible root tip rotation in Arabidopsis seedlings
induced by obstacle-touching stimulus. Science 250: 274–276.
Oliva M, Dunand C. 2007. Waving and skewing: how gravity and the surface of
growth media affect root development in Arabidopsis.New Phytologist 176:37–43.
Pampaloni F, Florin EL. 2008. Microtubule architecture: inspiration for novel
carbon nanotube-based biomimetic materials. Trends in Biotechnology 26: 302–
310.
Pandey S, Monshausen GB, Ding L, Assmann SM. 2008. Regulation of root-wave
response by extra large and conventional G proteins in Arabidopsis thaliana.The
Plant Journal 55: 311–322.
Paredez AR, Somerville CR, Ehrhardt DW. 2006. Visualization of cellulose
synthase demonstrates functional association with microtubules. Science 312:
1491–1495.
Peaucelle A, Wightman R, Hofte H. 2015. The control of growth symmetry
breaking in the Arabidopsis hypocotyl. Current Biology 25: 1746–1752.
Perez-Perez JM, Ponce MR, Micol JL. 2001. Mutations in the ULTRACURVATA2
gene of Arabidopsis thaliana, which encodes a FKBP-like protein, cause dwarfism,
leaf epinasty and helical rotation of several organs. International Journal of
Developmental Biology 45: S49–S50.
Pozo-Guisado E, Casas-Rua V, Tomas-Martin P, Lopez-Guerrero AM, Alvarez-
Barrientos A, Martin-Romero FJ. 2013. Phosphorylation of STIM1 at ERK1/2
target sites regulates interaction with the microtubule plus-end binding protein
EB1. Journal of Cell Science 126: 3170–3180.
Reinholz E. 1947a. Ausl€osung von R€ontgen-Mutationen bei Arabidopsis thaliana
(L.) HEYNH. und ihre Bedeutung f€ur die Pflanzenz€uchtung und
Evolutionstheorie. FIAT Report 1006:1–70.
Reinholz E. 1947b. R€ontgenmutationen bei Arabidopsis thaliana (L.) Heynh.
Naturwissenschaften 34:26–28.
Robinson DG, Preston RD, White RK. 1972. Fine-structure of swarmers of
Cladophora and Chaetomorpha. 3. Wall synthesis and development. Planta 107:
131–144.
Roelofsen PA. 1965. Ultrastructure of the wall in growing cells and its relation to the
direction of growth. Advances in Botanical Research 2:69–149.
Roy R, Bassham DC. 2014. Root growth movements: waving and skewing. Plant
Science 221–222:42–47.
Ruben GC, Bokelman GH, Krakow W. 1989. Triple-stranded left-hand helical
cellulose microfibril in Acetobacter xylinum and in tobacco primary cell wall. In:
Paice NGLAMG, ed. Plant cell wall polymers. Washington, DC, USA: ACS
Symposium, 278–298.
Rutherford R, Gallois P, Masson PH. 1998. Mutations in Arabidopsis thaliana
genes involved in the tryptophan biosynthesis pathway affect root waving on tilted
agar surfaces. The Plant Journal 16: 145–154.
Saffer AM, Carpita NC, Irish VF. 2017. Rhamnose-containing cell wall polymers
suppress helical plant growth independently of microtubule orientation. Current
Biology 27: 2248–2259.
Sakai T, Honing H, Nishioka M, Uehara Y, Takahashi M, Fujisawa N, Saji K, Seki
M, Shinozaki K, Jones MA et al. 2008. Armadillo repeat-containing kinesins and
a NIMA-related kinase are required for epidermal-cell morphogenesis in
Arabidopsis.The Plant Journal 53: 157–171.
Sambade A, Findlay K, Schaffner AR, Lloyd CW, Buschmann H. 2014. Actin-
dependent and -independent functions of cortical microtubules in the
differentiation of Arabidopsis leaf trichomes. Plant Cell 26: 1629–1644.
Sambade A, Pratap A, Buschmann H, Morris RJ, Lloyd C. 2012. The influence of
light on microtubule dynamics and alignment in the Arabidopsis hypocotyl. Plant
Cell 24: 192–201.
Schindelman G, Morikami A, Jung J, Baskin TI, Carpita NC, Derbyshire P,
McCann MC, Benfey PN. 2001. COBRA encodes a putative GPI-anchored
protein, which is polarly localized and necessary for oriented cell expansion in
Arabidopsis.Genes & Development 15: 1115–1127.
Schulgasser K, Witztum A. 2004. The hierarchy of chirality. Journal of Theoretical
Biology 230: 281–288.
Sedbrook JC, Carroll KL, Hung KF, Masson PH, Somerville CR. 2002. The
Arabidopsis SKU5 gene encodes an extracellular glycosyl phosphatidylinositol-
anchored glycoprotein involved in directional root growth. Plant Cell 14: 1635–
1648.
Serrano-Cartagena J, Robles P, Ponce MR, Micol JL. 1999. Genetic analysis of leaf
form mutants from the Arabidopsis Information Service collection. Molecular and
General Genetics 261: 725–739.
Shih HW, Miller ND, Dai C, Spalding EP, Monshausen GB. 2014. The receptor-
like kinase FERONIA is required for mechanical signal transduction in
Arabidopsis seedlings. Current Biology 24: 1887–1892.
Shoji T, Narita NN, Hayashi K, Asada J, Hamada T, Sonobe S, Nakajima K,
Hashimoto T. 2004. Plant-specific microtubule-associated protein SPIRAL2 is
required for anisotropic growth in Arabidopsis.Plant Physiology 136: 3933–3944.
Shoji T, Suzuki K, Abe T, Kaneko Y, Shi H, Zhu JK, Rus A, Hasegawa PM,
Hashimoto T. 2006. Salt stress affects cortical microtubule organization and
helical growth in Arabidopsis.Plant and Cell Physiology 47: 1158–1168.
Simmons C, S€oll D, Miglicaccio F. 1995. Circumnutation and gravitropism cause
root waving in Arabidopsis thaliana.Journal of Experimental Botany 46: 143–150.
Sivaguru M, Pike S, Gassmann W, Baskin TI. 2003. Aluminum rapidly
depolymerizes cortical microtubules and depolarizes the plasma membrane:
evidence that these responses are mediated by a glutamate receptor. Plant and Cell
Physiology 44: 667–675.
Smyth DR. 2016. Helical growth in plant organs: mechanisms and significance.
Development 143: 3272–3282.
Snow R. 1950. On the interpretation of geostrophic and auxin torsions. New
Phytologist 49: 145–154.
Snow R. 1962. Geostrophism. In: Ruhland W, ed. Physiology of movements,
encyclopedia of plant physiology. Berlin, Germany: Springer, 378–389.
Somerville C. 2006. Cellulose synthesis in higher plants. Annual Review of Cell and
Developmental Biology 22:53–78.
Speicher TL, Li PZ, Wallace IS. 2018. Phosphoregulation of the plant cellulose
synthase complex and cellulose synthase-like proteins. Plants 7: e52.
Sugimoto K, Williamson RE, Wasteneys GO. 2000. New techniques enable
comparative analysis of microtubule orientation, wall texture, and growth rate in
intact roots of Arabidopsis.Plant Physiology 124: 1493–1506.
Sunohara H, Kawai T, Shimizu-Sato S, Sato Y, Sato K, Kitano H. 2009. A
dominant mutation of TWISTED DWARF 1 encoding an alpha-tubulin protein
causes severe dwarfism and right helical growth in rice. Genes & Genetic Systems 84:
209–218.
Szymanski DB, Cosgrove DJ. 2009. Dynamic coordination of cytoskeletal and cell
wall systems during plant cell morphogenesis. Current Biology 19: 800–811.
Tangchun Z, Li L, Zhang Q. 2018. Advances in research on tortuous traits of plants.
Euphytica 214:1–15.
Thion L, Mazars C, Nacry P, Bouchez D, Moreau M, Ranjeva R, Thuleau P. 1998.
Plasma membrane depolarization-activated calcium channels, stimulated by
microtubule-depolymerizing drugs in wild-type Arabidopsis thaliana protoplasts,
display constitutively large activities and a longer half-life in ton 2 mutant cells
affected in the organization of cortical microtubules. The Plant Journal 13: 603–
610.
Thion L, Mazars C, Thuleau P, Graziana A, Rossignol M, Moreau M, Ranjeva R.
1996. Activation of plasma membrane voltage-dependent calcium-permeable
channels by disruption of microtubules in carrot cells. FEBS Letters 393:13–18.
Thitamadee S, Tuchihara K, Hashimoto T. 2002. Microtubule basis for left-
handed helical growth in Arabidopsis.Nature 417: 193–196.
Uyttewaal M, Burian A, Alim K, Landrein B, Borowska-Wykret D, Dedieu A,
Peaucelle A, Ludynia M, Traas J, Boudaoud A et al. 2012. Mechanical stress acts
via katanin to amplify differences in growth rate between adjacent cells in
Arabidopsis.Cell 149: 439–451.
New Phytologist (2019) Ó2019 The Authors
New Phytologist Ó2019 New Phytologist Trust
www.newphytologist.com
Review Tansley review
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Phytologist
16

Vaddepalli P, Fulton L, Batoux M, Yadav RK, Schneitz K. 2011. Structure-
function analysis of STRUBBELIG, an Arabidopsis atypical receptor-like kinase
involved in tissue morphogenesis. PLoS ONE 6: e19730.
Vain T, Crowell EF, Timpano H, Biot E, Desprez T, Mansoori N, Trindade LM,
Pagant S, Robert S, Hofte H et al. 2014. The cellulase KORRIGAN is part of the
cellulose synthase complex. Plant Physiology 165: 1521–1532.
Van der Does D,Boutrot F, Engelsdorf T, Rhodes J, McKenna JF, Vernhettes S,
KoevoetsI, Tintor N, VeerabaguM, Miedes E et al.2017.The Ar abidopsis leucine-
richrepeatreceptorkinaseMIK2/LRR-KISSconnectscellwallintegritysensing,root
growth and response to abiotic andbiotic stresses. PLoS Genetics 13:e1006832.
Vaughn LM, Baldwin KL, Jia G, Verdonk JC, Strohm AK, Masson PH. 2011.
Cytoskeleton and root growth behavior. In: Liu B, ed. The plant cytoskeleton.
Advances in plant biology, vol. 2. New York, NY, USA: Springer, 307–326.
Verger S, Liu M, Hamant O. 2019. Mechanical conflicts in twisting growth revealed
by cell-cell adhesion defects. Frontiers in Plant Science 10:1–9.
Voxeur A, Hofte H. 2016. Cell wall integrity signaling in plants: “To grow or not to
grow that’s the question”. Glycobiology 26: 950–960.
Wada H. 2012. Hierarchical helical order in the twisted growth of plant organs.
Physical Review Letters 109: 128104.
Wang C, Li J, Yuan M. 2007. Salt tolerance requires cortical microtubule
reorganization in Arabidopsis.Plant and Cell Physiology 48: 1534–1547.
Wang L, Nick P. 2017. Cold sensing in grapevine-Which signals are upstream of the
microtubular “thermometer”. Plant, Cell & Environment 40: 2844–2857.
Wang Y, Wang B, Gilroy S, Wassim Chehab E, Braam J. 2011. CML24 is involved
in root mechanoresponses and cortical microtubule orientation in Arabidopsis.
Journal of Plant Growth Regulation 30: 467–479.
Wang X, Zhu L, Liu B, Wang C, Jin L, Zhao Q, Yuan M. 2007. Arabidopsis
MICROTUBULE-ASSOCIATED PROTEIN18 functions in directional cell
growth by destabilizing cortical microtubules. Plant Cell 19: 877–889.
Wasteneys GO, Collings DA. 2004. Expanding beyond the great divide: the
cytoskeleton and axial growth. In: Hussey PJ, ed. The plant cytoskeleton in cell
differentiation and development. Oxford, UK: Blackwell Publishing, CRC Press,
83–115.
Wendrich JR, Yang B-J, Mijnhout P, Xue H-W, Rybel BD, Weijers D. 2018. IQD
proteins integrate auxin and calcium signaling to regulate microtubule dynamics
during Arabidopsis development. bioRxiv doi: 10.1101/275560.
Whittington AT, Vugrek O, Wei KJ, Hasenbein NG, Sugimoto K, Rashbrooke
MC, Wasteneys GO. 2001. MOR1 is essential for organizing cortical
microtubules in plants. Nature 411: 610–613.
Wightman R, Chomicki G, Kumar M, Carr P, Turner SR. 2013. SPIRAL2
determines plant microtubule organization by modulating microtubule severing.
Current Biology 23: 1902–1907.
Wu S, Xiao H, Cabrera A, Meulia T, van der Knaap E. 2011. SUN regulates
vegetative and reproductive organ shape by changing cell division patterns. Plant
Physiology 157: 1175–1186.
Yao M, Wakamatsu Y, Itoh TJ, Shoji T, Hashimoto T. 2008. Arabidopsis SPIRAL2
promotes uninterrupted microtubule growth by suppressing the pause state of
microtubule dynamics. Journal of Cell Science 121: 2372–2381.
Yuen CY, Pearlman RS, Silo-Suh L, Hilson P, Carroll KL, Masson PH. 2003.
WVD2 and WDL1 modulate helical organ growth and anisotropic cell expansion
in Arabidopsis.Plant Physiology 131: 493–506.
Yuen CY, Sedbrook JC, Perrin RM, Carroll KL, Masson PH. 2005. Loss-of-
function mutations of ROOT HAIR DEFECTIVE3 suppress root waving,
skewing, and epidermal cell file rotation in Arabidopsis.Plant Physiology 138: 701–
714.
Zhang Q, Lin F, Mao T, Nie J, Yan M, Yuan M, Zhang W. 2012. Phosphatidic acid
regulates microtubule organization by interacting with MAP65-1 in response to
salt stress in Arabidopsis.Plant Cell 24: 4555–4576.
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