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Handedness in plant cell expansion: a mutant
perspective on helical growth
Author for correspondence:
Henrik Buschmann
Tel: +49 541 9692248
Email: henrik.buschmann@biologie.
Received: 19 March 2019
Accepted: 4 June 2019
Henrik Buschmann and Agnes Borchers
Botanical Institute, Biology and Chemistry Department, University of Osnabruck, 49076, Osnabruck, Germany
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
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.
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. 1ch). 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).
(e) (g) (i)(h)
(b) (c)
(c) (d)
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
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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
CesA rosette
Cellulose microfibril
(right-handed twist)
Plasma membrane
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
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
(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; Lowe 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.,
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. Krussmann (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 Burgerpark (Osnabruck,
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
V. The mechanics behind twisting: from cells to
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
Unknown R First internode Mediated by
gibberellic acid
Chen et al. (2003)
kegne a-tubulin isotype Eragrostis tef Microtubule R Leaves, coleoptiles tua
Jost et al. (2015)
‘R’ a-tubulin isotype Lolium rigidum Microtubule R Leaves tua
Chu et al. (2018)
Rasen Unknown Cryptomeria
Unknown L/R alt. Needles, branches This paper
SUN IQ67 domain family Solanum
Microtubule L? Leaves oe Wu et al. (2011)
tid1 a-tubulin isotype Oryza sativa Microtubule R Leaves, stem tua
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
Endo-membrane suppr. Roots Propyzamide
Pandey et al. (2008)
ark2 Kinesin A. thaliana MAP R Roots Sakai et al. (2008)
aux1 Transporter, APC group A. thaliana Plasma and endo-
L Roots May be R in
Mirza (1987), Okada &
Shimura (1990)
cmu (1,2) Kinesin-light-chain
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
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
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,
Chevalier et al. (2005)
tch2 Calmodulin-like 24 A. thaliana Microtubule L Roots Missense allele
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
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,
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
Thitamadee et al. (2002)
tua a-tubulin isotype A. thaliana Microtubule R Roots e.g. tua6
Ishida et al. (2007a)
tub b-tubulin isotype A. thaliana Microtubule L Roots e.g. tub4
Ishida et al. (2007a,b)
tub b-tubulin isotype A. thaliana Microtubule R Roots e.g. tub3
Ishida et al. (2007a,b)
twd1/ucu2 FKBP-type
A. thaliana Plasma membrane L/R alt. Roots, hypocotyls, stems, carpels Defective auxin
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
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)
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.,
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
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
cell wall
oriented and dynamic Microtubules
oriented and twisted Cellulose Microfibrils
gibberellic acid
(RHM1; MUR8)
and Ca2+ influx
(salt stre ss)
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
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
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
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 Schaffner in 1993 (Burger, 1971;
Kranz & Kirchheim, 1987; Mainke & Koornneef, 1997). Even
though the mutant was used in subsequent investigations (Fabri &
Schaffner, 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.
We are grateful to four anonymous reviewers for their productive
comments. Sabine Zachgo is thanked for helpful discussions and
hosting this research in Osnabruck. We are also grateful to Lothar
Krienitz, who was involved in the sampling and classification of
Nitellopsis obtusa. The Botanical Garden in Osnabruck is thanked
for providing plant material. We further acknowledge Toni
Schaffner 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.
Henrik Buschmann
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Geometry and growth and division direction of individual cells are major contributors to plant organ shape and these processes are dependent on dynamics of microtubules (MT). Different MT structures, like the cortical microtubules, preprophase band and mitotic spindle, are characterized by diverse architectural dynamics (Hashimoto, 2015). While several MT binding proteins have been identified that have various effects on MT stability and architecture, they do not discriminate between the different MT structures. It is therefore likely that specific MT binding proteins exist that differentiate between MT structures in order to allow for the differences in architectural dynamics. Although evidence for the effect of specific cues, such as light and auxin, on MT dynamics has been shown in recent years (Lindeboom et al. , 2013; Chen et al. , 2014), it remains unknown how such cues are integrated and lead to specific effects. Here we provide evidence for how auxin and calcium signaling can be integrated to modulate MT dynamics, by means of IQD proteins. We show that the Arabidopsis IQD15-18 subclade of this family is regulated by auxin signaling, can bind calmodulins in a calcium-dependent manner and are evolutionarily conserved. Furthermore, AtIQD15-18 directly bind SPIRAL2 protein in vitro and in vivo and modulate its function, likely in a calmodulin-dependent way, thereby providing a missing link between two important regulatory pathways of MT dynamics. One sentence summary IQD proteins integrate auxin and calcium signaling, two major signaling pathways, to control the cytoskeleton dynamics and cell shape of Arabidopsis .
Full-text available
The sections in this article are Introduction Division Planes and the Establishment of Axiality Setting up for Axial Growth: Distinguishing Lateral and End Walls Establishing Axial Growth Polar Auxin Transport and its Regulation by the Actin Cytoskeleton Bending and Twisting – The Consequences of Differential Growth Conclusions and Future Perspectives Acknowledgements
Full-text available
Many plants grow organs and tissues with twisted shapes. Arabidopsis mutants with impaired microtubule dynamics exhibit such a phenotype constitutively. Although the activity of the corresponding microtubule regulators is better understood at the molecular level, how large-scale twisting can emerge in the mutants remains largely unknown. Classically, oblique cortical microtubules would constrain the deposition of cellulose microfibrils in cells, and such conflicts at the cell level would be relaxed at the tissue scale by supracellular torsion. This model implicitly assumes that cell-cell adhesion is a key step to transpose local mechanical conflicts into a macroscopic twisting phenotype. Here we tested this prediction using the quasimodo1 mutant, which displays cell-cell adhesion defects. Using the spriral2/tortifolia1 mutant with hypocotyl helical growth, we found that qua1-induced cell-cell adhesion defects restore straight growth in qua1-1 spr2-2. Detached cells in qua1-1 spr2-2 displayed helical growth, confirming that straight growth results from the lack of mechanical coupling between cells rather than a restoration of SPR2 activity in the qua1 mutant. Because adhesion defects in qua1 depend on tension in the outer wall, we also showed that hypocotyl twisting in qua1-1 spr2-2 could be restored when decreasing the matrix potential of the growth medium, i.e., by reducing the magnitude of the pulling force between adjacent cells, in the double mutant. Interestingly, the induction of straight growth in qua1-1 spr2-2 could be achieved beyond hypocotyls, as leaves also displayed a flat phenotype in the double mutant. Altogether, these results provide formal experimental support for a scenario in which twisted growth in spr2 mutant would result from the relaxation of local mechanical conflicts between adjacent cells via global organ torsion.
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Microtubules are filamentous structures necessary for cell division, motility and morphology. Microtubule dynamics are critically regulated by microtubule-associated proteins (MAPs). We outline the molecular mechanism by which the MAP, COMPANION OF CELLULOSE SYNTHASE1 (CC1), controls microtubule-bundling and dynamics in plants under salt stress conditions. CC1 contains an intrinsically disordered N-terminus that joins microtubules through conserved hydrophobic regions at evenly distributed foci. Structural data on the microtubule-bound CC1 N-terminus and mutation studies revealed the regions, and specific amino acids, that contribute to microtubule-binding. The importance of these regions and amino acids was confirmed through in vivo live cell imaging, which also explains how CC1 maintains cellulose synthesis during salt exposure. Surprisingly, the microtubule-binding mechanism of CC1 is remarkably similar to that of the prominent neuropathology-related protein Tau. Hence, we outline how MAP functions have converged during evolution across animal and plant cells.
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Plant organ shape is determined by the spatial-temporal expression of genes that control the direction and rate of cell division and expansion, as well as the mechanical constraints provided by the rigid cell walls and surrounding cells. Despite the importance of organ morphology during the plant life cycle, the interplay of patterning genes with these mechanical constraints and the cytoskeleton is poorly understood. Shapes of harvestable plant organs such as fruits, leaves, seeds and tubers vary dramatically among, and within crop plants. Years of selection have led to the accumulation of mutations in genes regulating organ shapes, allowing us to identify new genetic and molecular components controlling morphology as well as the interactions among the proteins. Using tomato as a model, we discuss the interaction of Ovate Family Proteins (OFPs) with a subset of TONNEAU1-recruiting motif family of proteins (TRMs) as a part of the protein network that appears to be required for interactions with the microtubules leading to coordinated multicellular growth in plants. In addition, SUN and other members of the IQD family also exert their effects on organ shape by interacting with microtubules. In this review, we aim to illuminate the probable mechanistic aspects of organ growth mediated by OFP-TRM and SUN/IQD via their interactions with the cytoskeleton.
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Tortuous-stem plants have extremely high ornamental value due to the zigzag shape or natural twisting of the branches. At present, the research about tortuous-stem plants focuses mainly on the morphological characteristics, anatomic structure and genetic characteristics, but few studies have been conducted on the genetic mechanism of tortuous stem traits. In recent years, numerous tortuous-stem mutants have been screened from Arabidopsis thaliana, Zea mays, Glycine max, Lycopersicon esculentum, Prunus and Populus indicating that tortuous traits may be closely related to the abnormal geotropic growth, uneven distribution of hormones and asymmetric development of vascular bundles. In this paper, advances in morphological characteristics, environmental regulation, genetic patterns, molecular mechanism and application prospects of tortuous-stem plants were summarized, aiming at providing the basis for revealing the molecular mechanism of tortuous stem traits.
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Cellulose, the most abundant biopolymer on the planet, is synthesized at the plasma membrane of plant cells by the cellulose synthase complex (CSC). Cellulose is the primary load-bearing polysaccharide of plant cell walls and enables cell walls to maintain cellular shape and rigidity. The CSC is comprised of functionally distinct cellulose synthase A (CESA) proteins, which are responsible for synthesizing cellulose, and additional accessory proteins. Moreover, CESA-like (CSL) proteins are proposed to synthesize other essential non-cellulosic polysaccharides that comprise plant cell walls. The deposition of cell-wall polysaccharides is dynamically regulated in response to a variety of developmental and environmental stimuli, and post-translational phosphorylation has been proposed as one mechanism to mediate this dynamic regulation. In this review, we discuss CSC composition, the dynamics of CSCs in vivo, critical studies that highlight the post-translational control of CESAs and CSLs, and the receptor kinases implicated in plant cell-wall biosynthesis. Furthermore, we highlight the emerging importance of post-translational phosphorylation-based regulation of CSCs on the basis of current knowledge in the field.
Plant growth and development are a genetically predetermined series of events but can change dramatically in response to environmental stimuli, involving perpetual pattern formation and reprogramming of development. The rate of growth is determined by cell division and subsequent cell expansion, which are restricted and controlled by the cell wall-plasma membrane-cytoskeleton continuum, and are coordinated by intricate networks that facilitate intra- and intercellular communication. An essential role in cellular signaling is played by calcium ions, which act as universal second messengers that transduce, integrate, and multiply incoming signals during numerous plant growth processes, in part by regulation of the microtubule cytoskeleton. In this review, we highlight recent advances in the understanding of calcium-mediated regulation of microtubule-associated proteins, their function at the microtubule cytoskeleton, and their potential role as hubs in crosstalk with other signaling pathways.
This article briefly reviews recent advances in nano-scale and micro-scale assessments of primary cell wall structure, mechanical behaviors and expansive growth. Cellulose microfibrils have hydrophobic and hydrophilic faces which may selectively bind different matrix polysaccharides and adjacent microfibrils. These distinctive binding interactions may guide partially aligned cellulose microfibrils in primary cell walls to form a planar, load-bearing network within each lamella of polylamellate walls. Consideration of expansive growth of cross-lamellate walls leads to a surprising inference: side-by-side sliding of microfibrils may be a key rate-limiting physical step, potentially targeted by specific wall loosening agents. Atomic force microscopy shows different patterns of microfibril movement during force-driven extension versus enzymatic loosening. Consequently, simulations of cell growth as elastic deformation of isotropic cell walls may need to be augmented to incorporate the distinctive behavior of growing cell walls.
Ectopic expression of the rice WINDING 1 (WIN1) gene leads to a spiral phenotype only in shoots but not in roots. Rice WIN1 belongs to a specific class of proteins in cereal plants containing a Bric-a-Brac/Tramtrack/Broad (BTB) complex, a non-phototropic hypocotyl 3 (NPH3) domain and a coiled-coil motif. The WIN1 protein is predominantly localized to the plasma membrane, but is also co-localized to plasmodesmata, where it exhibits a punctate pattern. It is observed that WIN1 is normally expressed in roots and the shoot-root junction, but not in the rest of shoots. In roots, WIN1 is largely localized to the apical and basal sides of cells. However, upon ectopic expression, WIN1 appears on the longitudinal sides of leaf sheath cells, correlated with the appearance of a spiral phenotype in shoots. Despite the spiral phenotype, WIN1-overexpressing plants exhibit a normal phototropic response. Although treatments with exogenous auxins or a polar auxin transport inhibitor do not alter the spiral phenotype, the excurvature side has a higher auxin concentration than the incurvature side. Furthermore, actin filaments are more prominent in the excurvature side than in the incurvature side, which correlates with cell size differences between these two sides. Interestingly, ectopic expression of WIN1 does not cause either unequal auxin distribution or actin filament differences in roots, so a spiral phenotype is not observed in roots. The action of WIN1 appears to be different from that of other proteins causing a spiral phenotype, and it is likely that WIN1 is involved in 1-N-naphthylphthalamic acid-insensitive plasmodesmata-mediated auxin transport.