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Journal of Experimental Botany, Vol. 65, No. 10, pp. 2703–2714, 2014
doi:10.1093/jxb/ert354 Advance Access publication 11 November, 2013
Review paper
Interplay between cell growth and cell cycle in plants
Robert Sablowski1,* and Marcelo Carnier Dornelas2
1 2 Cell and Developmental Biology Department, John Innes Centre, Norwich Research Park, Norwich, NR4 7UH, UK
Instituto de Biologia, Departamento de Biologia Vegetal, Universidade Estadual de Campinas, Campinas, SP, CEP 13083–862, Brazil
* To whom correspondence should be addressed. E-mail: [email protected]
Received 28 June 2013; Revised 27 September 2013; Accepted 27 September 2013
Abstract
The growth of organs and whole plants depends on both cell growth and cell-cycle progression, but the interaction
between both processes is poorly understood. In plants, the balance between growth and cell-cycle progression
requires coordinated regulation of four different processes: macromolecular synthesis (cytoplasmic growth), turgordriven cell-wall extension, mitotic cycle, and endocycle. Potential feedbacks between these processes include a cellsize checkpoint operating before DNA synthesis and a link between DNA contents and maximum cell size. In addition,
key intercellular signals and growth regulatory genes appear to target at the same time cell-cycle and cell-growth
functions. For example, auxin, gibberellin, and brassinosteroid all have parallel links to cell-cycle progression (through
S-phase Cyclin D–CDK and the anaphase-promoting complex) and cell-wall functions (through cell-wall extensibility
or microtubule dynamics). Another intercellular signal mediated by microtubule dynamics is the mechanical stress
caused by growth of interconnected cells. Superimposed on developmental controls, sugar signalling through the
TOR pathway has recently emerged as a central control point linking cytoplasmic growth, cell-cycle and cell-wall
functions. Recent progress in quantitative imaging and computational modelling will facilitate analysis of the multiple
interconnections between plant cell growth and cell cycle and ultimately will be required for the predictive manipulation of plant growth.
Key words: Arabidopsis, cell cycle, cell size, cell wall, endocycle, TOR.
Introduction
Plant cell sizes vary enormously, both between and within species (Sugimoto-Shirasu and Roberts, 2003). The typical cell
sizes seen in a given species and tissue is under genetic control
and results from the coordinated control of cell growth and
cell division. Size regulation is expected to affect cell function
in multiple ways. For example, altered surface to volume ratio
are expected to change nutrient and ion movement, the size
and geometry of cells may influence intercellular signalling,
and recent studies in yeast revealed that transcription changes
specifically in response to cell dimensions (Wu et al., 2010).
How cell growth and cell cycle are coordinated is one of the
remaining big questions in cell biology. Most of the progress
in this area has been made in fission and budding yeast, where
feedback between cell growth, cell geometry, and cell cycle has
been revealed (Jorgensen and Tyers, 2004; Talia et al., 2007).
In multicellular organisms, the question is further complicated by developmental controls that may override or modify
cell autonomous mechanisms to maintain cell-size homeostasis (Jorgensen and Tyers, 2004). In addition, cells growing within a tissue are also influenced and constrained by
the behaviour of their neighbours; this is especially true of
plant cells, which are locked together by symplastic growth
(Uyttewaal et al., 2012).
Unravelling the coordination between cell cycle and cell
growth will be essential for rational manipulation of plant
growth and shape. The potentially non-intuitive effects of the
feedbacks between growth and division within each cell and
especially in populations of communicating cells also pose
fascinating questions for the fast-developing area of computational modelling of plant development (Roeder et al., 2011;
© The Author 2013. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved.
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2704 | Sablowski and Dornelas
Roeder, 2012). Here are reviewed the interactions between
these processes, how these connections are modulated during
plant development, and how they relate to the genetic control
of plant growth. Where appropriate, plant processes will be
discussed in the wider context of cell cycle and growth regulation across eukaryotes.
Cellular processes that underpin
plant growth
The growth of plant tissues and organs results from a combination of cellular processes. The volume of individual cells
increases through cytoplasmic growth and turgor-driven
cell-wall extension, while the number of cells in the tissue is
increased by mitotic cycles. In addition, a variant form of the
cell cycle, the endocycle, is used to increase DNA contents
without cell division and is often associated with cell enlargement. Before discussing how these cellular processes are coordinated, each is briefly described below.
Cytoplasmic growth
Growth ultimately results from macromolecular synthesis.
The net accumulation of macromolecules and cellular components is sometimes referred to as cytoplasmic growth,
while cell expansion refers to increased cell volume caused
by enlargement of the vacuole. While vacuolar expansion is
considered an energetically cheap way to increase cell size,
cytoplasmic growth is closely linked to energy and nutrient
availability.
A central regulator of cytoplasmic growth is TOR, which
belongs to a family of phosphoinositide 3-kinase-related protein kinases that collectively signal metabolic and genomic
stress. In unicellular organisms, TOR links cellular growth to
nutrient availability, whereas in multicellular organisms, TOR
also has developmental roles as a growth coordinator across
tissues and organs (Zoncu et al., 2011). A key target of the
TOR pathway is ribosome biogenesis, which is highly energyintensive and therefore linked to the energy status of the cell,
in addition to being a prerequisite for much of the rest of biosynthetic activities. Accordingly, substrates of TOR include
the S6 kinase and eIF4E-binding protein 1 (E-BP1), which
regulate mRNA translation (Zoncu et al., 2011). In mammals, however, E-BP1 links the TOR pathway to the translational control of cell-cycle progression, rather than general
cell growth (Dowling et al., 2010). Another cellular process
controlled by TOR is autophagy, which is used to recycle cellular components when nutrients are limiting. In yeast and
mammals, TOR also regulates the actin cytoskeleton, thereby
controlling macromolecular traffic and producing polarized
cell growth (Wullschleger et al., 2006).
In plants, TOR and some TOR pathway components
are conserved (Menand et al., 2002; Berkowitz et al., 2008;
Brioudes et al., 2010). Lower and higher levels of TOR
expression cause the formation of smaller and larger organs,
respectively, suggesting that TOR signalling is an overall limiting factor for plant organ growth (DeProst et al., 2007).
The connection between plant TOR and nutrient signalling
in plants has been revealed recently. In contrast to animals
and yeast, in which amino acids signal nutrient status to the
TOR pathway, plant TOR was shown to have a central role in
linking the expression of growth-related genes to sucrose and
glucose levels (Xiong et al., 2013). Accordingly, in addition
to activating genes implicated in ribosome biogenesis and
repressing autophagy genes, plant TOR also activates metabolic responses to increased sugar levels, including glycolysis,
the TCA cycle, and starch and lipid storage (Caldana et al.,
2013; Xiong et al., 2013).
Turgor-driven cell expansion
Cytoplasmic growth and the cell cycle provide the building
blocks required for plant growth, but the space that can be
occupied by these building blocks is constrained by the cell
walls. Cell enlargement requires that the walls yield to the
cell’s turgor pressure and cell-wall relaxation appears to be
the major control point in this process (Wolf et al., 2012),
although organ primordium emergence can also be influenced by regulated expression of water channels and consequently water movement within the tissues (Peret et al.,
2012). In meristems and organ primordia, the increased
cell volume is occupied mostly by cytoplasmic growth and
increased nuclear volume associated with DNA replication, whereas in more differentiated tissues the increased
volume can result mostly from water uptake and vacuole
enlargement.
Controlled cell-wall relaxation is made possible by the
structure of cell walls, which contain cellulose and hemicellulose fibrils embedded in a matrix of pectin and proteins (Wolf
et al., 2012). These fibrils provide tensile strength and at the
same time can slip past each other to allow cell-wall extension. The orientation of the fibrils also influences the direction of preferential cell expansion and consequently helps to
determine cell and organ shape. Cell expansion is believed to
be a cyclic process with four steps: hydration, extension driven
by turgor pressure, mechanosensing followed by crosslinking
and dehydration, and finally secretion of new cell-wall components (Wolf et al., 2012) (Fig. 1).
In the first step, acidification of the apoplast caused by
P-ATPases triggers hydration and cell-wall loosening, a step
that is facilitated by expansin proteins, which function optimally at low pH and break hydrogen bonds between wall
polymers. This allows slippage of cellulose microfibrils and
consequently cell-wall extension. The extensibility of the cell
wall is also modulated by modification of the pectin matrix
in which the cellulose fibrils are embedded (Wolf et al., 2012).
One of the modifications is methylesterification, which can
alter Ca2+-mediated pectin crosslinking and accessibility to
enzymes involved in pectin turnover. Demethylesterification
is catalysed by pectin methylesterases (PMEs), whose activity
in turn is regulated by PME inhibitory proteins (PMIs). The
importance of pectin demethylesterification in the regulation
of tissue growth has been elegantly demonstrated during the
initiation of organ primordia in Arabidopsis shoot meristem
(Peaucelle et al., 2008, 2011).
Cell cycle and cell growth | 2705
Fig. 1. Overview of the cellular mechanisms that drive plant tissue growth. Cell growth (purple) is driven by cytoplasmic growth and
turgor-driven cell expansion, whereas cell cycle (orange) comprises mitotic cycles and endocycles. Key steps and regulatory points for
each process are explained in the main text. Potential cell-autonomous mechanisms that couple cell size and cell cycle (in green) include
a cell-size checkpoint (purple arrow) and the re-setting of the upper limit for cell growth by DNA contents, resulting in a correlation
between ploidy and cell size (karyo-cytoplasmic ratio; orange arrow).
In the third step of the cell-wall cycle, deformation of the
cell walls has been proposed to open stretch-activated Ca2+
channels (Wolf et al., 2012). The increased cytoplasmic Ca2+
inhibits the P-ATPases and activates plasma membrane
NADPH-oxidase, triggering an increase in reactive oxygen
species in the apoplast. The consequent alkalynization and
cross-linking of cell-wall components by reactive oxygen species stiffen the wall to stop extension.
In the final step, wall thickness is restored by secretion of
more cell-wall material. This includes deposition of new cellulose microfibrils by the cellulose synthase complex, which
migrates on the plasma membrane (Paredez et al., 2006). The
cortical microtubule array targets the insertion of cellulose
synthase complexes into the plasma membrane and influence their trajectories. In this way, the microtubule arrays
are believed to influence the orientation of cell-wall microfibrils and consequently the direction in which the cell wall
can be stretched more easily during the next cycle of cell-wall
extension.
Mitotic cycle
As in any multicellular organism, cell proliferation is a key
component of plant growth. The core feature of cell-cycle
control in eukaryotes is the oscillating activity of the cyclin-dependent kinase (CDK) (Coudreuse and Nurse, 2010),
around which additional mechanisms have evolved to confer robustness and conditional control over cell-cycle progression. Many of these additional features are conserved in
plants, such as specialized cyclins that control CDK activity at different cell-cycle phases (De Veylder et al., 2007),
the function of E2F genes in promoting DNA replication
(DeWitte and Murray, 2003) and the role of RBR genes in
controlling E2F function to balance cell proliferation and cell
differentiation (Doonan and Sablowski, 2010; Gutzat et al.,
2012). Plants also use CDK inhibitors of two different families: KIP-related proteins (KRPs) (Verkest et al., 2005b; Zhao
et al., 2012) and the plant-specific SIAMESE (SIM)-related
proteins (Churchman et al., 2006; Roeder et al., 2010) bind to
S-phase cyclin–CDKs to provide external input into cell-cycle
progression.
However, there are also differences in the plant cell-cycle
control. One of them is in the complexity of the cyclin family (De Veylder et al., 2007). The best-studied plant cyclins
have functions comparable to their counterparts in yeast
or metazoans: Cyclin D homologues control the transition
from G1 to S-phase, while Cyclin B homologues control the
G2–M transition. Plant Cyclin D associates with CDKA,
which is related to mammalian Cdk1, whereas Cyclin B functions in association with CDKB, which is specific to plants.
Furthermore, plants have unusually large numbers of cyclin
isoforms (at least 49 in Arabidopsis), the function of many
of which remains unknown. At the same time, plants do not
appear to have Cyclin E, which in animals is a major target
for organ growth regulators such as Hippo and dMyc (NetoSilva et al., 2009; Halder and Johnson, 2011)
Another difference is the role of Wee1 kinase and Cdc25
phosphatase in controlling the length of G2. In fission yeast
and animals, the Wee1 kinase phosphorylates and inactivates
CDK during G2. This builds up a pool of inactive CDK that
can be rapidly activated at the end of G2 through CDC25catalysed dephosphorylation. This sudden increase in CDK
activity allows a sharp and irreversible transition to mitosis
(O’Farrell, 2001). This was initially expected to be a core
feature of eukaryotic cell-cycle progression, but it turned
out that plants do not have it: mutation of the corresponding phosphorylation sites in plant CDKA have no effect on
cell-cycle progression (Dissmeyer et al., 2009) and functional
2706 | Sablowski and Dornelas
analysis of plant WEE1 revealed a role in the DNA damage checkpoint, but not in normal cell-cycle progression
(De Schutter et al., 2007). It remains unclear whether this or
other cell-cycle differences relate to plant-specific aspects of
development, such as the tight integration with environmental conditions and the lack of programmed cell death or cell
movement as important instruments for tissue growth.
Endocycle
In addition to the mitotic cell cycle, the endocycle plays
a prominent role in plant growth. This short-circuited version of the cell cycle, which allows DNA replication without
chromosome segregation and leads to increased cell ploidy, is
deployed widely in both animal and plant development (Lee
et al., 2009). During the endocycle, S-phase CDK activity
oscillates, while M-phase CDK activity is kept low. In mammals and insects, the S-phase CDK complex Cyclin E–Cdk2
plays a central role in the endocycle. As mentioned above,
plants do not appear to have Cyclin E, so they rely instead
on Cyclin D–CDK complexes to progress through S-phase
during the endocycle (DeWitte and Murray, 2003). However,
a subgroup of Cyclin D genes (CYCD3:1–3) specifically promotes the mitotic cycle instead of endocycles (DeWitte et al.,
2007). In both plants and animals, inhibition of M-phase
during the endocycle is caused in part by premature activation of the Fzr (fizzy-related) protein, which directs the anaphase-promoting complex (APC) to degrade mitotic cyclins
and promotes exit from M-phase during the mitotic cycle
(Lee et al., 2009). In plants, the shift from mitotic to endocycles is induced by both KRP and SIM CDK inhibitors
(Verkest et al., 2005a; Churchman et al., 2006); in addition,
an atypical E2F family member (E2Fe; DEL1: DP-E2F-like
1) promotes mitotic cycles by repressing aFZR homologue
called CCS52A2 (Vlieghe et al., 2005; Lammens et al., 2008).
Higher ploidy correlates with increased cell size both at the
whole-organism level and at the level of individual cells within
a tissue (Sugimoto-Shirasu and Roberts, 2003). The exact
mechanism behind this correlation remains unclear, although
it seems likely that increased ploidy releases a constraint on
maximum cell size because more gene copies are required to
service a larger volume of cytoplasm (as will be discussed in
more detail). Another specific feature in relation to growth is
that the endocycle bypasses the need to protect the cell from
the consequences of defective chromosome segregation and
therefore allows tissue growth to continue in conditions of
genomic stress that induce mitotic arrest (Lazzerini Denchi
et al., 2006; Adachi et al., 2011).
Coordination between cellular growth
processes
Coordination within each cell
Although plant cell sizes vary enormously between species and
tissue types, cell sizes are predictable for a given species and
tissue, implying that the balance between cell growth and cell
division is genetically controlled. This could be achieved by
several means: cell division could depend on cell size, growth
could be subordinate to cell-cycle progression, the regulatory
networks controlling growth and cell cycle could have shared
components, or both processes might be completely interdependent (Jorgensen and Tyers, 2004). Across eukaryotes, the
most common scenario is that cell cycle depends on growth (i.e.
cell growth continues when cell cycle is blocked), but the cell
cycle stalls when cellular growth is inhibited (Su and O’Farrell,
1998). In plants, growth can proceed normally when mitotic
rates are altered (see discussion on ‘compensation’), indicating
that, as in yeast and fly, mitotic rates do not drive cell growth.
In both fission and budding yeast, cell cycle is coupled to
cell growth by cell-size checkpoints that operate at the G1–S
and G2–M transitions (Jorgensen and Tyers, 2004; Talia
et al., 2007). In metazoans, the requirement for cell-size
checkpoints has been controversial, apparently being absent
in some cell types (Conlon and Raff, 2003) but present in
others (Tzur et al., 2009). Recent work indicates that plant
may also have cell-size checkpoint operating at the G1–S
transition (Schiessl et al., 2012) (Fig. 1). It is not known
whether this potential size checkpoint would respond to a
parameter related to cytoplasmic growth, such as ribosome
biogenesis (Jorgensen and Tyers, 2004), or whether it could
respond to actual cell dimensions, as shown recently for fission yeast (Moseley et al., 2009). The tight coordination
between cell size and DNA synthesis seen in the meristem is
relaxed during floral organ initiation (Schiessl et al., 2012),
but the functional significance of this developmental regulation is also unknown. One possibility is that cell geometry is expected to influence auxin transport through tissues
(Laskowski et al., 2008), so cell-size uniformity might be
required for auxin-dependent patterning in the meristem. In
contrast, increased variability in cell size is part of the normal programme of cell differentiation in organ primordia
(Roeder et al., 2010).
Like the mitotic cycle, the endocycle can also be subordinate to cellular growth. For example, reduced growth caused
by starvation or by inhibition of the insulin pathway suppresses endoreplication in insects, and Drosophila dMyc
mutants have reduced endocycles, which may be a secondary
consequence of growth arrest (Maines et al., 2004; Qi and
John, 2007). In Arabidopsis, overexpression of CYCD2;1
reduced cell sizes and inhibited endocycles (Qi and John,
2007). This has been interpreted as evidence that cell enlargement is required for entry into the endocycle; however, it cannot be excluded that CYCD2;1 might have a more direct role
in the choice between mitosis and endocycle, as proposed for
CYCD3;1 (DeWitte et al., 2007).
Although the aforementioned size checkpoints involve input
from cell growth into cell-cycle progression, the interaction
between both processes is not unidirectional (Goranov and
Amon, 2010). In fission yeast, for example, cellular growth
accelerates in G2 and this transition requires completion of
DNA synthesis (Baumgartner and Tolic-Norrelykke, 2009). In
plants, inhibition of DNA synthesis stopped growth in the meristem (Grandjean et al., 2004), although it remains to be seen
whether this was due to a simple metabolic limitation or due to
active signalling between replication stress and cellular growth.
Cell cycle and cell growth | 2707
These observations may relate to the classic ‘karyoplasmic ratio’
hypothesis, which implies that larger amounts of DNA can sustain a larger amount of cytoplasm and therefore ploidy levels
set the upper limit of cytoplasmic growth (Sugimoto-Shirasu
and Roberts, 2003; Jorgensen and Tyers, 2004) (Fig. 1). The
exact molecular mechanism that links DNA content to cell size
remains unknown. In line with the central role of ribosome biogenesis in cellular growth, it has been speculated that ribosome
synthesis could be the limiting factor related to the amount of
DNA available (Sugimoto-Shirasu and Roberts, 2003).
Cytoplasmic increase linked to protein synthesis may not
be the only aspect of cellular growth that is connected to the
cell cycle. As discussed above, cell-wall relaxation is a prerequisite for cell growth by either cytoplasmic increase or vacuolization. Regulation of cell-wall extensibility, for example by
expression of expansin or pectin methylesterase, can control
organ initiation with its associated cytoplasmic growth and
cell division (Pien et al., 2001; Peaucelle et al., 2011). As in the
case of the DNA synthesis inhibition already mentioned, it is
not known whether there is active signalling between cell-wall
functions and cytoplasmic-growth or cell division. In budding yeast, the cell-wall integrity pathway is linked to protein
synthesis (Goranov and Amon, 2010); it would be interesting
to study whether the plant cell-wall integrity pathway (Wolf
et al., 2012) could also feedback to cellular growth through
links to protein synthesis.
In summary, all cellular activities that sustain plant growth
must be interconnected, but it remains unclear how this is
achieved and how complex the coordination is. A plausible candidate for global coordinator is the TOR pathway.
Inhibition of TOR function in Arabidopsis not only confirmed the expected links between plant TOR and translation
processes but revealed novel direct links to cell cycle through
direct E2Fa phosphorylation and activation of DNA synthesis genes (Xiong et al., 2013). In addition, transcriptional
changes in response to TOR inhibition included multiple
genes linked to cell-wall modification, including expansins,
arabinolactan proteins, and lignin synthesis genes (Caldana
et al., 2013; Xiong et al., 2013).
Growth coordination across cells
Superimposed on to the question of how growth-related
mechanisms are coordinated within each cell is the question
of how cells coordinate their growth with each other to produce tissues and organs with genetically programmed shapes
and sizes.
Prominent in discussions of the relation between cell and
organ growth is the phenomenon of ‘compensation’, in which
cell number and cell size are balanced to maintain overall
organ size. This phenomenon has been well studied, for example, in Drosophila imaginal discs. Manipulating the levels of
cell-cycle regulators such as E2F and CDC25 results in imaginal discs of normal size containing fewer, large cells, when
cell cycle is inhibited, or more numerous, smaller cells, when
cell division is stimulated (Su and O’Farrell, 1998). Overall,
these results are consistent with the idea discussed above that
cell proliferation tends to be subordinate to cellular growth.
Plants also show numerous examples where cell proliferation and cell growth are balanced to maintain overall organ
size (Horiguchi and Tsukaya, 2011). For example, cell-cycle
progression has been inhibited by expression of a dominant
inhibitory CDK (Hemerly et al., 1995) or stimulated by
increased expression of S-phase cyclins (DeWitte et al., 2003;
Qi and John, 2007) but in both cases overall leaf growth was
maintained by compensatory changes in cell size. The maintenance of consistent organ sizes in spite of variations in cell
number and cell size have prompted the suggestion that plants
have mechanisms to monitor whole-organ size. A related, but
more extreme idea is the ‘organismal’ hypothesis of plant
growth, which considers that growth is controlled at the organ
level and that cell division simply partitions space to provide
enough DNA to service the growing tissue (John and Qi,
2008). This assumption would predict that growth parameters
vary smoothly across the tissues and do not correlate with
individual cells. Live imaging and quantitative image analysis, however, have recently revealed a surprising heterogeneity in growth rates and growth isotropy between neighbouring
cells (Uyttewaal et al., 2012). This indicates that plant cells
are actually discrete units for growth control, even though
intercellular signals must be superimposed on the cell-autonomous controls to coordinate cell behaviour across the tissue.
Another study showed that growth can be heterogeneous even
within each cell and that the unit for growth control might be
individual facets between cells, rather than whole cells (Elsner
et al., 2012)—in other words, rather than supracellular, the
units for growth control might actually be subcellular.
If plant cells are the units for growth control, then the mechanism that balances cell size with cell division must ultimately
operate within each cell, although cell communication would be
expected to coordinate cell-autonomous decisions. Accordingly,
examples of compensation have been described that are either
cell autonomous or whose effect propagates across the tissues
(Kawade et al., 2010). Mutations that disrupt compensation
could potentially give insight into the coordination between cell
cycle and cell growth; however, multiple layers of complexity
have hampered the genetic analysis. First, the phenomenon is
context-dependent and is not believed to occur in organs with
indeterminate growth (as will be discussed in details). Second,
the details of compensation are variable: sometimes it involves
endoreduplication, sometimes not (Ferjani et al., 2007); as
already mentioned, sometimes it operates cell autonomously,
sometimes not (Kawade et al., 2010). Third, mutants implicated in compensation have defects in diverse processes ranging from ribosomal biogenesis (Fujikura et al., 2009) to the
hydrolysis of cytosolic pyrophosphate (Ferjani et al., 2011) to
chromatin defects leading to DNA damage (Hisanaga et al.,
2013). Although each of these processes has connections to cell
growth or cell cycle, so far the identified genes do not suggest a
central coordinating mechanism.
Intercellular signals that coordinate cell growth and
division
Well-known intercellular signals that pattern tissues can also
participate in growth control. An example from animals is
2708 | Sablowski and Dornelas
the role of the Dpp (Decapentaplegic) diffusible signal in the
fly imaginal discs. In regulating tissue growth, Dpp is unlikely
to target primarily cell division; instead, it appears to control
cellular growth, with cell cycle coupled to it, or it may control
cell growth and cell cycle in parallel (Wartlick and GonzálezGaitán, 2011). Accordingly, Dpp is connected to pathways
such as Hippo and dMyc, which coordinately regulate animal
cell growth and cell cycle (Neto-Silva et al., 2009).
The best-studied plant developmental signal is auxin, which
activates its own intercellular transport to produce dynamic
patterns of distribution across the tissues (Vanneste and Friml,
2009). Auxin function is linked not only to cell differentiation, but also to the cell cycle and cell-wall relaxation (PerrotRechenmann, 2010), so like Dpp, auxin is a good candidate to
integrate tissue patterning and growth. Details of how polarized auxin transport is linked to tissue growth, however, are
still unclear. Local accumulation of auxin in the apoplast is
believed to activate the ABP1 receptor, leading to activation
of proton pumps and consequently to apoplast acidification, which as mentioned above, facilitates cell-wall extension
(Vanneste and Friml, 2009) (Fig. 2). This mechanism would fit
the observation above that individual cell facets are units for
growth control; however, it is not known how far the acidification and wall relaxation extend or how cell-wall extensibility
correlates with local levels of apoplastic auxin.
Gibberellin (GA) is another plant hormone with established links to both cell cycle and cell growth (Fig. 2). GA
has been proposed to promote mitotic cycles by two mechanisms: lowering expression of both KRP and SIM CDK
inhibitors (Achard et al., 2009) and promoting expression
of E2Fe (Claeys et al., 2012), which as mentioned above
represses FZR to prevent premature destruction of mitotic
Cyclin CDK. The molecular basis for the long-known role of
GA in regulating cell expansion remains less clear, but recent
work revealed that DELLA proteins, which are destabilized
by GA, interfere with the function of tubulin chaperones
that are required for proper microtubule dynamics and consequently for cell expansion (Locascio et al., 2013). Unlike
auxin, movement of GA between cells is poorly understood,
although there is recent evidence that GA is actively transported across root tissues to accumulate in endodermal cells
(Shani et al., 2013). Another signal that controls both cell
growth and cell cycle is brassinosteroid, which not only controls cell expansion by targeting wall acidification via BRI1
signalling to P-ATPases (Caesar et al., 2011) and by promoting cytoskeletal changes (Clouse, 2011), but also controls the
balance between cell proliferation and differentiation in the
meristem (Gonzalez-Garcia et al., 2011). Cytokinin also has
direct links to the cell cycle through CYCD expression (RiouKhamlichi et al., 1999); however, so far there are no direct
links to cellular growth.
Apart from sending hormone signals, cells can influence
their neighbours through mechanical effects (Lecuit and Le
Goff, 2007). As mentioned above, mechanical constraints are
especially important in plant tissues, where cells are immobilized and attached to each other through their cell walls.
Accordingly, there is increasing evidence that plant cells
respond to mechanical signals to adjust their growth and
that microtubule arrays play an important role in this process (Uyttewaal et al., 2010) (Fig. 2). Surprisingly, rather
than smoothing growth differences between neighbouring
cells, the cellular response to mechanical signals can function to enhance growth heterogeneity: if the ability to adjust
microtubule arrays is impaired, cellular growth rates become
more uniform and isotropic, but this results in abnormal tissue growth (Uyttewaal et al., 2012). Considering that one
of the functions of auxin is to regulate cell-wall extensibility, it might be expected that mechanical sensing would be
Fig. 2. Overview of intercellular signals that coordinate cell growth and cell cycle in plants. Cellular activities that drive tissue growth are
summarized in the purple and orange boxes as in Fig. 1. Arrows show activation and blunted lines show repression of key regulatory
nodes. In addition to intercellular signals (gibberellin, sucrose, brassinolide, auxin, mechanical stress), TOR is highlighted as a potential
integrator of cytoplasmic growth, cell expansion, and cell cycle.
Cell cycle and cell growth | 2709
integrated with hormonal signalling. The evidence so far,
however, is that mechanical and hormonal signals can be
uncoupled, for example, auxin can induce anisotropic growth
in the absence of microtubule arrays (Hamant et al., 2008).
In addition to diffusible developmental signals and
mechanical interactions, collective cell behaviour can also be
influenced by competition for limiting resources such as nutrients. Sucrose, for example, promotes cell-cycle progression by
activating expression of CYCD genes (Riou-Khamlichi et al.,
2000; Sanz et al., 2011). As explained above, sugar also functions as a systemic signal that regulates organ growth through
the TOR pathway. However, sugar signalling through TOR
appears to impose an overall limit on growth without changing the hormonal and gene regulatory pathways that pattern
the tissues and shape organs: the levels and spatial distribution of auxin responses and of patterning genes remained the
same in root meristems whose growth was arrested by TOR
signalling during starvation (Xiong et al., 2013).
Developmental regulation of cell cycle and
cell growth
Transitions in growth regime
For simplicity, the sections above described the known and
potential links between processes that drive cell growth and
the cell cycle without considering developmental context.
However, it is clear that not all regulatory interactions are relevant in the same cells, at the same time. For example, tissue
growth in meristems and organ primordia combines cytoplasmic growth, turgor-driven wall extension, and mitotic cycles,
whereas in late stages of leaf development, tissue growth is
based on turgor-driven extension with increase in vacuolar
size, combined with endocycles.
Thus regulatory genes that control organ growth need to
target different cellular processes at different developmental stages and often control the transition between different
cellular growth regimes (Breuninger and Lenhard, 2010).
A well-studied example of transition in growth modes occurs
in leaf development. Because of its characteristic reiterative and open-ended development, spatial patterns in plants
are often a record of a temporal sequence of events. In the
case of leaves, as primordia are produced on the flanks of
the meristem, the most distal part of the primordium (the
‘tip’) contains the first cells committed to become an organ
and, thus, also the most mature ones. Accordingly, multiple events in leaf development occur in ‘waves’, including a
wave of mitotic arrest described from the tip to the bottom
of the developing leaf (Donnelly et al., 1999). Quantitative
analysis of cell division and cell shapes revealed that this
wave of mitotic arrest is surprisingly rapid (Kazama et al.,
2010; Andriankaja et al., 2012). This work also showed that
the transition to cell expansion includes activation of photosynthesis-related genes, and acquisition of photosynthetic
capacity is required for the shift to cell expansion. Additional
changes in gene expression include expression of different
isoforms of both cell-wall-related genes and of ribosomal
synthesis and activation of a SIM-like gene, which is a likely
candidate for the mitotic arrest (Andriankaja et al., 2012).
Thus the transition between growth regimes in leaf development requires coordinated changes in metabolism, cell-wall
functions, and cell cycle.
Another example of changes in growth-related cell behaviour occurs during the initiation of organ primordia on the
flanks of the floral meristem. Although growth of the meristem and primordia is generally considered to be ‘proliferative’, recent work showed clear changes in growth regime
in the primordium. This included a shift form isotropic to
anisotropic tissue growth, increased cell proliferation rates,
increased cell volumes, and, as mentioned above, loss of coordination between cell size and DNA synthesis (Schiessl et al.,
2012). All these changes were genetically separable from
primordium initiation and were promoted by the JAGGED
(JAG) gene, revealing a clear link between a regulatory gene
and a transition in tissue growth.
Links between cell cycle, cell growth, and
regulatory genes
Although developmental regulatory genes such as JAG have
clear effects on cell behaviour, the molecular basis for these
effects remains poorly understood. Another example is the
AP2-type transcription factor AINTEGUMENTA (ANT),
which controls growth during the proliferative phase of organ
development. Loss of ANT function causes small organs,
while overexpression increases organ size; the larger organs
had either more cells or larger cells, depending on organ type
(Krizek, 1999; Mizukami and Fischer, 2000). A molecular
link to cell-cycle regulation was suggested by the observation that ectopic ANT activates CYCD3 (Mizukami and
Fischer, 2000). However, activation of CYCD3;1 is not sufficient to produce the growth effects of ANT (DeWitte et al.,
2003), and combined mutation of CYCD3 family members
does not abolish organ growth (DeWitte et al., 2007). This
has been pointed out to suggest that ANT is likely to regulate cellular growth in parallel to cell cycle (Breuninger and
Lenhard, 2010), although the exact links to cellular growth
are unknown.
Genes of the TCP (Teosinte branched 1, CYCLOIDEA,
and PROLIFERATING CELL FACTORS 1 and 2) family
have also been implicated in the control of cell proliferation
in meristems and organ primordia (Martín-Trillo and Cubas,
2010). Accordingly, mutation of genes in the class II TCP
subfamily show organ growth defects combined with changes
in CYCD3 expression (Gaudin et al., 2000; Nath et al., 2003).
Recent data, however, reveal an indirect link between class II
TCPs and the cell cycle: TCP4 activates miRNA 396, which
represses a set of GRF transcription factors, whose expression is in turn associated with mitotic cyclin expression
(Rodriguez et al., 2010). In addition, the interaction with cell
cycle is likely to be just one aspect of TCP function: class II
TCPs have been proposed to control the progression through
multiple maturation stages of the leaf, with cell division being
just one feature of early stages (Efroni et al., 2008). One of
the additional functions of TCP4 is to regulate jasmonic acid
2710 | Sablowski and Dornelas
synthesis (Schommer et al., 2008). Recently it has been demonstrated that this regulation is antagonistically counteracted
by class I TCP 20, probably in cooperation with class I TCP9
(Danisman et al., 2012). Global analysis of genes affected by
the class I TCP20 also revealed enrichment for genes involved
in cell-wall function, including genes encoding several cellulose synthase subunits and several xyloglucanendotransglucosylase hydrolases (XTHs) (Herve et al., 2009; Danisman
et al., 2012).
One difficulty with these analyses is that it remains difficult to extricate direct regulatory interactions from indirect, downstream consequences. The genes affecting plant
organ growth with the most comprehensive analysis of
direct regulatory interactions are the flower organ identity
genes (Kaufmann et al., 2009; Wuest et al., 2012). In this
case, direct interactions are apparent with cell-cycle genes
(CDKA, KRP, and SIM CDK inhibitors, E2Fc), cell-wallrelated genes (encoding cellulose synthase subunits, XTHs,
PME, and PMI), numerous genes related to ribosomal biogenesis, and several of the growth regulatory genes already
mentioned, including JAG and a large fraction of the class II
TCP genes. Additionally, recent data supports the participation of both MADS and TCP transcription factors in
higher-order protein complexes that regulate development
(Dornelas et al., 2011; Guo et al., 2013), potentially adding
another layer of complexity in the developmental regulation
of the interplay between cell cycle and cell growth.
Conclusions
Interdependence and coordinated regulation of growthrelated pathways appear to be the rule. The majority of
molecular signals and developmental regulatory genes target multiple aspects of growth, such as macromolecular
synthesis, cell-wall remodelling, cytoskeleton rearrangements, and cell-cycle progression. This may reflect contextdependent functions of the regulatory genes and signals:
in different cell types and developmental stages, different
cellular processes might become bottlenecks for overall
growth and therefore critical targets for developmental
regulation. Alternatively, parallel regulation of multiple
processes may be required within each cell. Targeting individual growth processes may not be effective in modulating growth because the processes are interdependent or
because they are connected by homeostatic regulatory
loops, exemplified by the potential compensation mechanism discussed above.
Because shape and size are emergent properties of systems that involve small modules (cells) to form larger units
(tissues and organs), unravelling the feedbacks involving
the mechanisms controlling cell growth, cell-cycle progression, and cell-wall modifications will require extensive modelling. Recent examples of how computer simulations can
capture non-intuitive behaviour of cells and tissues include
the analysis of cellular responses to mechanical signals
(Uyttewaal et al., 2012), and modelling of how JAG may
interact with tissue polarity signals such as auxin to control
petal growth (Sauret-Güeto et al., 2013). At the same time,
the development and testing of realistic models will require
data with much finer cellular and temporal resolution.
This will be facilitated by the development of quantitative
imaging analysis methods, both in live tissues (Fernandez
et al., 2010) and with the ability to combined information
on cell geometry and cell cycle (Andriankaja et al., 2012;
Schiessl et al., 2012; for an example of combined imaging
of cell volume and cell-cycle progression, see Fig. 3). In the
long term, understanding the interplay between regulatory
signals and cellular processes during growth will be essential to reap the fruits of predictive manipulation of plant
growth and shape.
Acknowledgements
Fig. 3. Example of quantitative imaging of cell geometry
combined with cell cycle. This 3-D image of an Arabidopsis floral
meristem was reconstructed from confocal microscopy images,
with cells coloured according to their volume (colour bar below
the image) and green nuclei showing incorporation of a nucleotide
analogue to mark cells in S-phase. The methods used are
described in Schiessl et al. (2012).
The authors thank Pauline Haleux and Richard Immink
for critical reading. Work in the Sablowski and Dornelas
laboratories is supported by a joint grant from the BBSRC
(BB J007056 1) and FAPESP (2011 51412-1). RS is additionally supported by the BBSRC (BB F005571 1) and the
John Innes Foundation. MCD also acknowledges funding from Coordenação de Aperfeiçoamento de Pessoal de
Nivel Superior (CAPES, Brazil) and Conselho Nacional de
Desenvolvimento Científico e Tecnológico (CNPq, Brazil).
Cell cycle and cell growth | 2711
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