Analysis of PIN2 polarity regulation and Mob1 function
in Arabidopsis root development
Inaugural-Dissertation zur Erlangung der Doktorwürde
der biologischen Fakultät
der Albert-Ludwigs-Universität Freiburg im Breisgau
Vorgelegt von
Francesco Pinosa
Februar 2010
Dekan: Prof. Dr. Ad Aertsen
Vorsitzender des Promotionsausschußes: Prof. Dr. Eberhard Schäfer
Betreuer der Doktorarbeit: Prof. Dr. Klaus Palme
Referent: Dr. Roman Ulm
Koreferent: PD Dr. Eva Decker
Tag der Verkündigung des Prüfungsergebnisses: 13.04.2010
Table of contents
Page Nr.
Summary
1
Aims of the thesis
3
Conclusions
3
Chapter I
General introduction
7
PIN polarity and cell cycle regulation in root
development
Chapter II
Discrete distribution of AtPIN2 in plasma membrane
35
domains
Chapter III
Effects of sphingolipid biosynthesis inhibition on PIN2
57
polarity and root development in Arabidopsis
Chapter IV
Analysis of AtMOB1 function in plant development
Acknowledgements
81
109
Summary
1
Summary
The plant hormone auxin plays an essential role in root growth and development. PIN
proteins, auxin efflux facilitators, are polarly localised at the plasma membrane (PM)
and direct auxin flow. Local auxin levels are thus established and coordinate cell
division and differentiation. Through the regulatory activity of cell-cycle components,
such as MOB proteins, a highly organised root structure is finally generated.
In the first part of this study, the mechanisms regulating the establishment and
maintenance of AtPIN2 polarity in root cells were investigated. It was shown for the
first time that PIN2 clustered in discrete PM domains which were stable over time as
well as upon perturbations of the cytoskeleton or the PM lipid composition. PIN2
domains remained associated with the cell wall (CW) after plasmolysis of epidermal
cells. These observations suggested a possible PIN2-CW interaction which might
prevent the free lateral diffusion of the protein and assist the mechanism of PIN2
polarity maintenance. Chemical inhibition of sphingolipid biosynthesis impaired the
correct establishment of PIN2 polarity after cytokinesis by reducing the rate of
endocytosis. The use of auxin reporters revealed that the defects in PIN2 polarity
affected root growth and gravity response by altering the local auxin levels both prior
to and after gravistimulation. Thus, the sphingolipid composition of the PM proved to
be essential in the establishment of PIN polarity and root development.
In the second part of this thesis, the role of AtMOB1 genes in root and plant
development was studied. MOB1 proteins are cell-cycle regulators conserved among
eukaryotes and acting at mitotic exit and cytokinesis. Arabidopsis MOB1 proteins
exhibited a nuclear localisation regulated throughout the progression of mitosis and
cytokinesis, supporting the hypothesis that they are involved in cell division control. A
loss-of-function mutant for MOB1A displayed defects in root growth and root
meristem organisation, in particular for the cell files surrounding the quiescent centre.
In addition, normal plant growth and reproduction were impaired. The majority of
these defects were confirmed by the analysis of independently generated RNAi lines.
Summary
2
The root phenotype of MOB1A knock-out and RNAi transformants underlined the
importance of cell division regulation for proper tissue patterning.
The results reported in this thesis provide new insights into the mechanism of PIN
polarity regulation and the function of MOB1 genes and highlight the relevance of
both factors for root development.
Aims and conclusions
3
Aims of the thesis
The work presented here has been aimed at the analysis of root development
regulation. At the beginning of this study, the following goals were set:
I. to identify novel factors that contribute to the polarity maintenance of PIN auxin
carriers. For this purpose, AtPIN2 was chosen as model case;
II. to investigate the role of membrane lipid composition in the regulation of PIN
polarity. In particular, the function of sphingolipids in the establishment of
AtPIN2 polarity was addressed;
III. to clarify the function of the cell division regulator AtMOB1 in root meristem
organisation.
Conclusions
The distribution of PIN2 in the plasma membrane (PM) of Arabidopsis root cells was
investigated by live-imaging of a functional PIN2-GFP fusion protein and
immunocytochemical approaches (Chapter II). PIN2 was clustered in discrete PM
domains of approximately 400 nm size. PIN2 domains exhibited a low motility within
the PM as demonstrated by FRAP measurements of their lateral diffusion. In addition,
PIN2 domains were maintained upon perturbations of the cytoskeleton as well as
changes in the PM lipid composition. After plasmolysis of epidermal cells PIN2
domains remained associated to the cell wall. The current hypothesis predicts that
PIN2 polarity is maintained by slow lateral diffusion and constitutive endocytosis.
However, the factor responsible for PIN2 low lateral motility has so far remained
elusive. The results obtained suggest that the association of PIN2 domains with the cell
wall prevents the free lateral diffusion of PIN2.
The role of sphingolipids in the establishment of PIN2 polarity was demonstrated by
chemical inhibition of sphingolipid biosynthesis (Chapter III). The primary effect of
this treatment was a general reduction in the rate of endocytosis. This hindered the
establishment of PIN2 polarity after cytokinesis, in turn affecting auxin distribution at
the root tip both prior to and after gravistimulation. Consequently, root growth and
Aims and conclusions
4
gravity response were impaired. Similar effects on PIN2 polarity establishment were
recently reported for alterations in the sterol composition of the PM (Men et al., 2008).
Although sphingolipids and sterols are known to be the major components of lipid
microdomains (reviewed by(Zappel and Panstruga, 2008), a direct link between PIN2
polarity regulation and these microdomains cannot be deduced. Indeed, alterations in
the membrane levels of sterols or sphingolipids result in a rather unspecific effect on
endocytosis. In addition, PIN2 does not co-purify with sterol- and sphingolipidenriched membrane fractions (unpublished data from Cho, Y.J., Teale, W., Palme,
K.;(Titapiwatanakun et al., 2009). Independent variations in sterol and sphingolipid
contents, whether or not they affect lipid microdomain clustering, could lead to a
comparable loss of membrane integrity and therefore to the general defects in
endocytic trafficking.
The function of two Arabidopsis MOB1-like genes in root and plant development was
investigated through loss-of-function and RNAi lines (Chapter IV). While MOB1B
knock-out did not exhibit any visible defect, the expression of MOB1A was required
for proper plant growth and reproduction. In particular, the roots of mob1A null
mutants exhibited a reduced size and a shorter meristem as well as defects in the spatial
arrangement of cells surrounding the quiescent centre. This phenotype was also
common to independently generated RNAi lines for MOB1A, which displayed
alterations in the expression pattern of different identity markers for the root tip.
MOB1 proteins showed a nuclear localisation regulated throughout the progression of
mitosis and cytokinesis, hence strongly indicating their involvement in cell division,
similarly to their eukaryotic orthologs. Originally identified in yeast, MOB1 is a
component of a signalling network mediating exit from mitosis and cytokinesis (Luca
and Winey, 1998; Luca et al., 2001). In multicellular organisms, MOB1 proteins play a
role in fundamental processes like cell proliferation and apoptosis, thus controlling
appropriate cell number and organ size (reviewed by(Vitulo et al., 2007). The
functional characterisation of plant MOB1 genes reported here highlights the
Aims and conclusions
5
importance of cell division regulation for the proper patterning and development of the
root.
References
Luca FC, Mody M, Kurischko C, Roof DM, Giddings TH, Winey M (2001) Saccharomyces cerevisiae
Mob1p is required for cytokinesis and mitotic exit. Mol Cell Biol 21: 6972-6983
Luca FC, Winey M (1998) MOB1, an essential yeast gene required for completion of mitosis and
maintenance of ploidy. Mol Biol Cell 9: 29-46
Men S, Boutte Y, Ikeda Y, Li X, Palme K, Stierhof YD, Hartmann MA, Moritz T, Grebe M (2008) Steroldependent endocytosis mediates post-cytokinetic acquisition of PIN2 auxin efflux carrier
polarity. Nat Cell Biol 10: 237-244
Titapiwatanakun B, Blakeslee JJ, Bandyopadhyay A, Yang H, Mravec J, Sauer M, Cheng Y, Adamec J,
Nagashima A, Geisler M, Sakai T, Friml J, Peer WA, Murphy AS (2009) ABCB19/PGP19
stabilises PIN1 in membrane microdomains in Arabidopsis. Plant J 57: 27-44
Vitulo N, Vezzi A, Galla G, Citterio S, Marino G, Ruperti B, Zermiani M, Albertini E, Valle G, Barcaccia
G (2007) Characterization and evolution of the cell cycle-associated mob domain-containing
proteins in eukaryotes. Evol Bioinform Online 3: 121-158
Zappel NF, Panstruga R (2008) Heterogeneity and lateral compartmentalization of plant plasma
membranes. Curr Opin Plant Biol 11: 632-640
- Chapter I -
PIN polarity and cell-cycle regulation in root
development
Chapter I
8
Abstract
The growth of the root is sustained by the proliferative activity of a meristematic pool
of cells located at the tip. Meristem organisation and function depend on the ability of
individual cells to perceive the positional signals received from the neighbouring cells
and to consequently coordinate their cell-cycle activity. A primary cue of spatial
information is provided by the concentration of the phytohormone auxin. Local
differences in auxin levels are generated by the concerted activity of several PIN
proteins, which mediate auxin efflux from cells thereby promoting its transport along a
tissue. The directionality of auxin transport is determined by the polar plasma
membrane localisation of PIN proteins to one side of the cell. The establishment and
maintenance of PIN polarity are highly dynamic processes, which involve many
cellular players. The local auxin levels resulting from PIN activity are subsequently
translated into coordinated events of cell division and cell differentiation. Specific
cell-cycle regulators have been shown to play a relevant function into these processes
and are thus involved in the correct tissue patterning of the meristem. In this review,
the role of auxin transport in root development and the mechanisms regulating PIN
polarity will be first summarised. Finally, the importance of cell-cycle control in root
meristem development will be discussed by presenting two examples of cell-cycle
components.
1. Introduction (Raven, 1999)
Plant roots accomplish a crucial function for plant growth providing anchorage to the
ground and absorption of nutrients and water and serve sometimes as storage organs.
The root system originates from a primary root that develops during embryogenesis.
Lateral roots are produced post-embryonically from the primary root and share with it
an essentially identical structure.
The root of the model plant Arabidopsis thaliana represents a very useful system for
investigating the basis of root development. The Arabidopsis root displays a simple
structure and its mature part is composed of radially organised cell layers which from
inside to outside form vasculature, pericycle, endodermis, cortex and epidermis (Dolan
Chapter I
9
et al., 1993). Each layer consists of cell files that originate from “initial cells” located at
their apical end in the root meristem (Fig. 1). The initials continuously generate new
cells which in turn undergo a highly stereotyped sequence of divisions along the
meristematic zone. Cells reaching the distal border of the root meristem enter the
elongation zone and are subsequently differentiated according to their cell types. At
the extreme apex, columella and lateral root cap (LRC) cells are also formed from their
initials with similar series of cell divisions. Initial cells surround a group of few
mitotically inactive cells in the middle of the root meristem that constitutes a
“quiescent centre” (QC). The QC function is to maintain the stem-cell identity of the
initial cells by inhibiting their differentiation (van den Berg et al., 1997).
The highly organised structure of the root apical meristem is fundamental to the proper
growth of the root and to its correct response to multiple environmental stimuli.
Several factors mediate the appropriate patterning of the root meristem. Among these,
the distribution of the plant growth regulator auxin provides positional cues required
for meristem organisation (Sabatini et al., 1999; Blilou et al., 2005). In addition, the
patterning process depends on coordinated events of cell division and cell
Chapter I
10
differentiation, which are controlled by the activity of specific cell-cycle regulators
(Blilou et al., 2002; Wildwater et al., 2005).
In the first part of this review, recent advances in the understanding of auxin
distribution will be summarised, focusing in particular on the process of polar auxin
transport and on the mechanisms regulating the localisation of auxin carrier proteins.
In the second part, specific examples of cell-cycle regulators that control
postembryonic development of the root meristem will be presented and discussed.
(Cambridge, 1996)
2. The role of auxin and polar auxin transport in root development
Auxin is implicated in many
physiological processes in plants, coordinating
development at the cellular and organ level and ultimately up to the whole plant
(reviewed by(Bhalerao and Bennett, 2003; Fleming, 2006; Tanaka et al., 2006; Teale et
al., 2006; Benjamins and Scheres, 2008; Mockaitis and Estelle, 2008). Auxin displays the
characteristics of a typical hormone-like substance, since it is synthesised in one
location and then transported to a site of action where it is perceived by a receptor.
Furthermore, it has the ability to regulate development in a dose-dependent manner
via concentration gradients (reviewed by(Benjamins and Scheres, 2008; Vanneste and
Friml, 2009).
In roots, free auxin accumulates in the extreme apex with a maximum centred over the
QC and the columella initials (Sabatini et al., 1999; Friml et al., 2002; Petersson et al.,
2009). Auxin accumulation overlaps with the expression domain of PLETHORA (PLT)
genes, encoding AP2-domain transcription factors (Aida et al., 2004). PLT proteins act
in a dose-dependent manner to regulate root development and their concentration
maximum in the root stem cell niche is instructive to stem cell fate (Galinha et al.,
2007). The control of QC identity and the consequent stem cell maintenance depends
on the joint action of PLT proteins with SHORTROOT (SHR) and SCARECROW
(SCR), two GRAS family transcription factors that participate also in the radial
patterning of the root (Di Laurenzio et al., 1996; Helariutta et al., 2000; Sabatini et al.,
2003; Aida et al., 2004). The expression of PLT genes responds to auxin accumulation
and depends on auxin response transcription factors (Aida et al., 2004). Thus, auxin
Chapter I
11
accumulation acts as the master organiser of the pattern of root stem cell niche,
providing the spatial input for PLT gene expression. This hypothesis was demonstrated
for both the specification of the stem cell niche during embryo development and for its
maintenance during post-embryonic root growth (Aida et al., 2004; Blilou et al., 2005;
Xu et al., 2006; Galinha et al., 2007).
Intercellular auxin transport and local auxin biosynthesis in the root apex are both
required to maintain the correct distribution of auxin (Sabatini et al., 1999; Friml et al.,
2002; Blilou et al., 2005; Stepanova et al., 2008). Defects in these two processes
independently cause abnormalities in the pattern of the stem cell niche and in root
development. In particular, the intercellular transport of auxin is fundamental to
stabilise the auxin maximum at the level of QC and columella initials, therefore
focusing the expression of PLT genes in these cells (Sabatini et al., 1999; Blilou et al.,
2005). The ability of auxin to move between cells in a directional manner makes it
unique among plant signalling molecules.
2.1.
The polar transport of auxin and its molecular components
The distribution of auxin along the plant body involves two distinct major pathways.
The first serves as a rapid, long-distance source-to-sink conveyance and occurs by
loading auxin into the phloem in young shoot tissues, which are biosynthetically
highly active, and transporting it towards sink tissues like the root (Cambridge and
Morris, 1996;(Swarup et al., 2001; Marchant et al., 2002). The second type of transport
operates over shorter distances in a slow, carrier-mediated, cell-to-cell manner. Its
main feature is to be predominantly unidirectional and is thus referred to as “polar
auxin transport” (PAT).
In an attempt to explain the cellular mechanism of PAT, the chemiosmotic hypothesis
was formulated in the Seventies based on the biochemical and physiological data
available at the time (Rubery and Sheldrake, 1974; Raven, 1975). The model proposed
that the pH difference between apoplast and cytoplasm causes the retention of auxin
inside the cell. As auxin exhibits the characteristics of a weak acid (pKa = 4.75), a small
portion (approximately 16%) is protonated in the acidic environment of the apoplast
Chapter I
12
(pH 5.5), thus becoming lipophilic and able to diffuse passively through the plasma
membrane (PM). In the more basic cytoplasm (pH 7.0), the deprotonation of auxin
prevents it from permeating through the PM and traps it inside the cell. Thus, the exit
of auxin from the cell requires the existence of specific efflux carriers. The
directionality of auxin flow was predicted to rely on the asymmetric localisation of the
efflux carriers on one side of the cell within a field of cells. Besides the passive diffusion
of auxin inside the cell, additional influx carriers promote active auxin uptake. The
components of auxin influx correspond to the members of the AUX1/LIKE AUX1
(AUX1/LAX) family, which encode transmembrane proteins sharing significant
similarity with plant amino acid permeases (Bennett et al., 1996; Swarup et al., 2004;
Yang et al., 2006; Swarup et al., 2008). These proteins play important roles in processes
such as phyllotaxis, gravitropism, lateral root spacing, lateral root emergence and root
hair development (Bennett et al., 1996; Bainbridge et al., 2008; Swarup et al., 2008);
Jones et al., 2009). Auxin efflux is promoted by the cooperative action of PIN proteins
and P-glycoproteins of the ABCB transporter family (ABCB/PGP) (Blakeslee et al.,
2007; Mravec et al., 2008). The latter were identified as binding-proteins of the auxin
transport inhibitor N-1-naphthylphthalamic acid (NPA) (Murphy et al., 2002) and
mediate auxin efflux both in plant and non-plant systems (Geisler et al., 2005; Terasaka
et al., 2005; Cho et al., 2007).
PIN proteins are plant-specific transmembrane proteins, which fulfil the characteristics
of efflux carriers predicted by the chemiosmotic hypothesis. Indeed, they display polar
subcellular localisations that correlate with the directions of auxin flow (Wisniewska et
al., 2006) and have been shown to directly transport auxin (Petrasek et al., 2006;
Mravec et al., 2009; Titapiwatanakun et al., 2009). Single and multiple pin mutants
show typical defects in auxin-related processes, such as tropisms, embryo development,
root meristem patterning, organogenesis and vascular tissue differentiation (Galweiler
et al., 1998; Muller et al., 1998; Friml et al., 2002; Friml et al., 2003; Reinhardt et al.,
2003; Blilou et al., 2005; Scarpella et al., 2006). The Arabidopsis genome contains eight
PIN family members, which exhibit specific expression domains and distinct
localisations of the corresponding proteins (reviewed by(Paponov et al., 2005). In the
Chapter I
13
root meristem, PIN proteins orchestrate a complex loop of auxin flow that stabilises the
accumulation pattern required for proper root development (Blilou et al., 2005) (Fig. 2).
PIN2
fluxes in Arabidopsis root tip. The different
directions of auxin transport are established
by the distinct polar localisation of PIN
proteins in each root layer.
PIN2
PIN1
PIN4
PIN2
PI N2
PIN2
PIN2
PIN2
PIN2
PIN2
PIN1
PIN3
PIN7
PIN2
Figure 2. Schematic representation of auxin
PIN3
PIN7
The localisation of PIN1 to the lower side (basally) of stele cells promotes the acropetal
flow of auxin towards the stem cell niche. PIN4 further supports the accumulation of
auxin in QC and columella initials by localising basally in provascular cells and in a
non-polar manner in QC and cells surrounding it. The acropetal transport of auxin is
aided by the basal localisation of PIN3 and PIN7 in vascular cells of the elongation
zone. PIN3 and PIN7 are also expressed without pronounced polarity in tiers two and
three of columella cells. In epidermis and lateral root cap, PIN2 apical localisation
promotes the basipetal flow of auxin towards the elongation zone, away from its
maximum. However, from the epidermis auxin can re-enter the acropetal transport
route through the localisation of PIN2 to the lower side of young cortical cells (Fig. 2).
The distinct pathways of PIN-mediated auxin transport are assumed to be responsible
for different developmental and morphogenic processes. The acropetal transport
Chapter I
14
promoted by PIN1, PIN3, PIN4 and PIN7 in the stele is required to stabilise the auxin
maximum at the root apex, in turn maintaining the PLT-dependent stem cell domain
(Blilou et al., 2005). The basipetal transport to meristematic cells, mediated by PIN2,
plays a crucial role in the regulation of meristem size (Blilou et al., 2005). Furthermore,
the basipetal route is involved in the root response to changes in the gravity vector
(Rashotte et al., 2000). Upon gravistimulation, PIN3 is re-localised to the new lower
side of columella cells and drives the redistribution of auxin towards epidermis and
LRC cells at the bottom side of the root (Friml et al., 2002; Ottenschlager et al., 2003).
On the same side, PIN2 successively promotes the accumulation of auxin at the
elongation zone, which in turn causes the inhibition of cell expansion and leads to root
bending (Young et al., 1990; Luschnig et al., 1998; Muller et al., 1998; Ottenschlager et
al., 2003).
In conclusion, the polarity of PIN proteins in the different cell files of the root is
important for the control of the local auxin levels, which in turn regulate proper root
growth.
2.2.
Regulation of PIN polar targeting
PIN proteins represent prominent markers of plant cell polarity. Their polar targeting
is determined by a number of cellular factors that collectively contribute to the control
of auxin efflux (Fig. 3). In the next paragraphs, the main players involved in the
modulation of PIN polarity will be presented and their physiological importance
discussed.
2.2.1. Phosphorylation
Different PIN proteins display opposite polar localisations in the same cell, as in the
case of PIN1 and PIN2 in mature cortical cells or in epidermal cells with ectopically
expressed PIN1 (Wisniewska et al., 2006). This differential targeting requires a cellular
mechanism able to identify some polarity signals embedded in the protein sequence.
Furthermore, the insertion of a GFP tag at a specific position within the hydrophilic
loop of PIN1 reverses its polar localisation in epidermal cells from basal to apical
Chapter I
15
(Wisniewska et al., 2006). Sequence-based signals for decisions on PIN targeting to a
specific PM domain are related to phosphorylation sites present in PIN protein
sequences. The Ser/Thr protein kinase PINOID (PID) and the protein phosphatase 2A
(PP2A) influence the phosphorylation status of PIN proteins, dictating their apical-tobasal distribution (Friml et al., 2004; Michniewicz et al., 2007). In plants overexpressing
PID or with PP2A loss-of-function, PIN1, PIN2 and PIN4 are highly phosphorylated
and their polarity is shifted from basal to apical. In this situation, the root meristem is
depleted of auxin and root development is affected (Christensen et al., 2000; Benjamins
et al., 2001; Friml et al., 2004; Michniewicz et al., 2007). In contrast, inhibition of PID
function results in low phosphorylation levels of PIN1 and in its preferential basal
targeting, which in turn causes defects in embryo and shoot organogenesis (Christensen
et al., 2000; Benjamins et al., 2001; Friml et al., 2004). Noteworthily, PID directly
phosphorylates the central hydrophilic loop of PIN proteins and PP2A counteracts this
action (Michniewicz et al., 2007).
2.2.2. Sterols
Plant sterols play a significant role in the regulation of PAT and in auxin-related
developmental processes. Genetic studies carried out on two enzymes acting at
different steps of sterol biosynthesis have revealed the importance of PM sterol
composition for correct PIN polar targeting. In the sterol-deficient sterol
methyltransferase 1 (smt1) mutant, the polarity of PIN1, PIN3 and AUX1 is disturbed
and auxin distribution in the root tip as well as meristem patterning is affected
(Willemsen et al., 2003). Root gravitropism defects in the cyclopropylsterol isomerase 1
(cpi1) mutant are brought about by a lack in the establishment of PIN2 polarity after
cytokinesis of meristematic cells (Men et al., 2008). PIN2 is normally delivered to both
sides of the cell plate during cell division and after cytokinesis is retrieved from one
side in order to maintain the polarity of the mother cell in both daughter cells. In the
cpi1 mutant, the endocytosis of PM proteins is impaired and PIN2 remains localised at
both apical and basal sides of post-cytokinetic cells. These results suggest that the
removal of PIN2 from one side of the newly formed cell wall after cell division requires
Chapter I
16
a sterol-dependent endocytosis step (Men et al., 2008). Furthermore, sterols and PIN2
have been shown to share the same trafficking pathway (Grebe et al., 2003). The
relation between sterols and PIN proteins is highlighted also by the observation that
ABCB19/PGP19 stabilises PIN1 in detergent-resistant membrane fractions enriched in
sterols and sphingolipids (Titapiwatanakun et al., 2009). These biochemical
preparations
are
assumed
to
correspond
to
membrane
microdomains
that
compartmentalise cellular processes and function as signalling platforms (reviewed
by(Zappel
and
Panstruga,
2008).
However,
the
functional
relevance
of
ABCB19/PGP19-PIN1 association in membrane microdomains remains to be
established.
2.2.3. Endocytosis and polar recycling (Mayer, 1991)
Genetic and pharmacological studies have suggested that the polar localisation of PIN
proteins is regulated by their constitutive cycling between the PM and endosomal
compartments. The first evidence for PIN cycling was obtained using the fungal toxin
brefeldin A (BFA), which interferes with vesicle trafficking (Steinmann et al., 1999;
Geldner et al., 2001). In the presence of BFA and the protein biosynthesis inhibitor
cycloheximide, PIN1 internalises from the PM and accumulates into so-called BFA
compartments. Upon BFA removal, PIN1 is relocated back to the PM. These results
indicate that PIN1 undergoes continuous rounds of endocytosis from and recycling to
the PM. Moreover, the use of actin depolymerising drugs in combination with BFA
treatments demonstrated that PIN cycling depends on an intact actin cytoskeleton
(Geldner et al., 2001). The prominent target of BFA action in Arabidopsis is GNOM, a
guanine-nucleotide exchange factor for ADP-ribosylation factor GTPases (ARF GEF)
that regulates vesicle budding at endosomes (Shevell et al., 1994; Geldner et al., 2003).
When GNOM activity is inhibited, the formation of exocytic vesicles carrying PIN1
and other cargos to the PM is prevented. In the gnom mutant, the polar localisation of
PIN1 is thus impaired, causing severe defects in embryo development (Mayer et al.,
1991;(Steinmann et al., 1999). Besides BFA treatment of Arabidopsis roots, the
internalisation of PIN proteins and their subsequent recycling to the PM were also
Chapter I
17
directly documented by means of a photoconvertible fluorescent version of PIN2
(Dhonukshe et al., 2007).
Endocytosis and recycling to the proper PM polar domain appear fundamental to the
mechanism of PIN polarity establishment. A recent study has demonstrated that newly
synthesised PIN proteins are originally delivered in a non-polar fashion to the PM and
Chapter I
18
their asymmetric distribution is then achieved through internalisation and polar
recycling (Dhonukshe et al., 2008). Interferences in PIN endocytosis prevent the
establishment of PIN polarity and result in severe developmental defects (Dhonukshe
et al., 2008).
The recycling pathways of PIN proteins differentiate according to their polar
destination. While PIN basal targeting requires GNOM activity and is severely
impaired after BFA treatment or in the gnom mutant, PIN apical targeting is only
partially affected by these conditions (Kleine-Vehn et al., 2008). This different
behaviour was demonstrated by monitoring the fate of PIN2 and ectopically expressed
PIN1, apically and basally localised in the same epidermal cells, respectively (KleineVehn et al., 2008). Upon BFA treatment, basal PIN1 was completely retrieved from the
PM and accumulated in BFA-compartments, while PIN2 largely maintained its PM
localisation. Thus, apical cargos employ an alternative recycling pathway that might
involve different, BFA-insensitive, ARF GEFs. However, it was proposed that basal and
apical targeting pathways are interconnected by a transcytosis mechanism, defined as
the trafficking of polar cargos from one side of the cell to the other. This hypothesis is
based on the observation that upon prolonged BFA treatment PIN1 and PIN2
translocate from the basal to the apical side of stele and cortical cells, respectively
(Kleine-Vehn et al., 2008). If correct, the transcytosis mechanism might regulate the
rapid changes in PIN polarity observed during important developmental processes such
as tropisms and morphogenesis (Friml et al., 2002; Benkova et al., 2003; Friml et al.,
2003; Kleine-Vehn et al., 2008).
Additional players, which are specifically involved in PIN2 subcellular trafficking, have
been identified. PIN2 internalisation from the PM requires the activity of the GNOMLIKE1 (GNL1) ARF GEF (Teh and Moore, 2007). Furthermore, PIN2 cycling depends
on endosomal compartments containing SORTING NEXIN1 (SNX1), which are distinct
from GNOM endosomes (Jaillais et al., 2006).
In summary, the polar targeting of PIN proteins is achieved and maintained through a
dynamic mechanism of vesicular trafficking that involves different subcellular
compartments.
Chapter I
19
2.2.4. Auxin
The establishment of tissue pattern and polarity requires the ability of individual cells
to perceive their relative position among the neighbouring cells. With the formulation
of the “canalisation hypothesis” decades ago, it was proposed that auxin might provide
such a positional cue (Sachs, 1981). This process would involve a positive feedback
mechanism between auxin levels and the capacity and directionality of auxin flow.
Concordantly, auxin has been shown to control PIN abundance and polarity at
multiple levels. On the transcriptional level, auxin regulates not only PIN expression
but also that of PID, which in turn determines PIN polar destination (Benjamins et al.,
2001; Peer et al., 2004; Vieten et al., 2005). PIN protein abundance can be modulated
by auxin also at the post-translational level. Low concentrations of auxin, which
normally occur at the upper side of gravistimulated roots, promote PIN2 internalisation
and vacuolar targeting for proteolysis (Abas et al., 2006; Kleine-Vehn et al., 2008).
Prolonged high auxin levels induce PIN2 proteolysis as well, but through a different
mechanism, which involves PIN2 ubiquitination, internalisation and degradation by
the 26S proteasome (Abas et al., 2006). Another auxin-dependent feedback seems to
play a role at the level of PIN subcellular trafficking. Synthetic auxin analogues such as
naphthalene-1-acetic acid (1-NAA) and 2,4-dichlorophenoxyacetic acid (2,4-D) have
been shown to inhibit endocytosis, including the internalisation of PIN proteins
(Paciorek et al., 2005). This effect results in the accumulation of PIN at the PM and at
least in tobacco suspension cells it leads to an increase in auxin efflux. In the case of
PIN2, the positive effect of auxin on its PM levels depends on auxin signalling and
correct sterol composition of the PM (Pan et al., 2009).
Local auxin accumulation induces rearrangements in PIN polarity in different
developmental processes such as phyllotaxis and vascular tissue formation or
regeneration (Heisler et al., 2005; Sauer et al., 2006; Scarpella et al., 2006). Additionally,
exogenous auxin application is sufficient to bring about changes in PIN polar
localisation (Sauer et al., 2006). The underlying mechanism remains unknown but
seems to involve auxin-dependent derepression of transcription factors of the Auxin
Response Factor (ARF) class (reviewed by(Quint and Gray, 2006).
Chapter I
20
In conclusion, PIN polarity and abundance appear to be modulated by auxin through
different cellular processes, constituting a feedback mechanism able to fine-tune auxin
gradients.
2.2.5. Modulation of PIN polarity by developmental and environmental stimuli
The polarity of PIN proteins within specific tissues can be modulated by developmental
programs and environmental signals. Changes in PIN localisation lead to a redirection
of the auxin flow and to subsequent modifications of auxin distribution, which in turn
trigger different morphogenic responses.
Changes in PIN polarity occur already during embryogenesis and are required for the
specification of the root pole and the formation of the root meristem (Friml et al., 2003;
Weijers et al., 2005). During embryo development in Arabidopsis, the localisation of
PIN7 in the suspensor changes from apical (towards the apical cell) at early stages to
basal at later stages, while PIN1, which is non-polar in the proembryo, becomes basally
localised in cells adjacent to the future root pole (Friml et al., 2003). These
modifications of PIN polarity drive auxin accumulation in the region of root meristem
specification. Post-embryonic organogenesis is similarly promoted by rearrangements
in PIN polarity. The resulting changes in auxin distribution are important for defining
the position of new leaves and flowers emerging from the shoot apical meristem
(Reinhardt et al., 2003; Heisler et al., 2005), and the new growth axis of lateral roots
(Benkova et al., 2003). Additional examples of PIN polarity alterations during
developmental programs are provided by leaf vasculature formation (Scarpella et al.,
2006) and vasculature regeneration after wounding (Sauer et al., 2006).
Modifications in the PM distribution of PIN proteins can also result from
environmental stimuli, such as changes in the gravity vector. Upon gravistimulation of
Arabidopsis roots, the statoliths present in columella cells sediment to the bottom side
and trigger the relocation of PIN3 to the same side from its originally uniform
distribution. Auxin flow is thus redirected to the lower side of the root, causing in turn
root bending (Friml et al., 2002).
Chapter I
21
3. Cell-cycle regulation in root development
Proper development of plant organs requires an intensive coordination between cellcycle control and differentiation processes. This cross-talk appears particularly crucial
in meristematic regions, where cell division and differentiation have to occur in a
balanced manner to maintain the stem-cell pool and eventually organ shape and size
(reviewed by(Jakoby and Schnittger, 2004; de Jager et al., 2005; Gutierrez, 2005;
Ingram and Waites, 2006; Maughan et al., 2006; Bogre et al., 2008). Cell-cycle
regulators represent major candidates to perform the choice between cell proliferation
and cell differentiation programs on the base of different internal signals. The main
components of the eukaryotic cell-cycle machinery are conserved also in plants
(Vandepoele et al., 2002), but the mechanism by which cell expansion and division
rates are controlled across plant tissues remains largely unknown. Due to the essential
function of several cell-cycle regulators, defects in their activity cause lethal
phenotypes already in the haploid gametophytic phase (Capron et al., 2003; Kwee and
Sundaresan, 2003; Ebel et al., 2004; Iwakawa et al., 2006; Liu et al., 2008). Thus,
studying the role played by these proteins in the context of post-embryonic
development has been difficult.
Few mutants with a defective activity of proteins operating at crucial steps of the
cell-cycle have been useful tools to demonstrate the importance of cell-cycle regulation
for root meristem patterning and development. HOBBIT and RETINOBALSTOMARELATED proteins represent two examples of components acting in this regulatory
process and their roles will be here presented and discussed.
3.1.
HOBBIT
The HOBBIT (HBT) gene encodes a protein homologous to CDC27/Nuc2, a component
of the anaphase-promoting complex/cyclosome (APC/C) conserved among eukaryots
(Blilou et al., 2002). The APC/C is a multi-subunit ubiquitin ligase triggering
proteolytic degradation of several cell-cycle regulators, including mitotic cyclins
(reviewed by(Peters, 2006; Li and Zhang, 2009). All APC/C subunits have counterparts
in plants (Capron et al., 2003; Fulop et al., 2005; Eloy et al., 2006) and they have been
Chapter I
22
implicated in regulating both mitotic cycles and endocycles (Cebolla et al., 1999;
Serralbo et al., 2006; Lammens et al., 2008; Perez-Perez et al., 2008). Since ploidy levels
in plants correlate with cell size (Sugimoto-Shirasu and Roberts, 2003), the APC/C can
influence both cell division and cell expansion (Cebolla et al., 1999; Capron et al., 2003;
Kwee and Sundaresan, 2003; Serralbo et al., 2006; Perez-Perez et al., 2008).
In contrast to other genes encoding core APC/C subunits, HBT is not essential for
gametophytic development in Arabidopsis (Capron et al., 2003; Kwee and Sundaresan,
2003; Perez-Perez et al., 2008). In fact, this function is accomplished by the redundant
activity of HBT and its homolog CDC27a (Perez-Perez et al., 2008). Tissue patterning
during embryo development does not require HBT function, but defects caused by
mutations in the HBT gene appear first during post-embryonic growth (Blilou et al.,
2002). In mature embryos of homozygous hbt mutants of strong loss-of-function
alleles, the differentiation of root meristematic cell types, such as QC, columella root
cap and lateral root cap, is impaired and the maintenance of tissue-specific gene
expression patterns is affected (Willemsen et al., 1998; Blilou et al., 2002). As a
consequence, root growth is arrested.
The primary effects brought about by the elimination of HBT activity during root
development have been investigated with a system for the induction of loss-of-function
clones (Serralbo et al., 2006). Impaired cell division and reduced cell expansion, caused
by alterations in mitotic progression and endoreduplication, respectively, represent the
primary consequences of a reduced expression of HBT in roots (Serralbo et al., 2006).
This result was confirmed by the analysis of homozygous plants carrying hypomorphic
mutations (Perez-Perez et al., 2008). Thus, the effects on cell division planes and the
failed maintenance of cell identity observed in null HBT mutants are only secondary
effects of cell division and cell expansion defects. This implies that interference with
cell-cycle regulation can affect cell fate determination during root growth.
3.2.
RETINOBLASTOMA-RELATED
The RETINOBLASTOMA-RELATED (RBR) protein is the Arabidopsis ortholog of the
mammalian retinoblastoma (RB), a cell-cycle regulator that acts at the transition from
Chapter I
23
G1 to S phase (Weinberg, 1995). RB prevents cell cycle progression and hence cell
division by inhibiting E2F transcription factors. The activity of RB is inhibited through
phosphorylation by cyclin-dependent kinases (CDKs), which are in turn regulated by
CDK inhibitors. Plant genomes contain orthologs for all the components of this G1-S
regulatory pathway (Inze, 2005). RBR is a single-copy gene in Arabidopsis and its
function is already required during gametophytic development (Ebel et al., 2004). In
post-embryonic roots, RBR is expressed in the meristem with the highest transcript
levels in cells that have completed cell division (Wildwater et al., 2005).
The role of RBR in root development has been investigated by gene silencing with a
RNA-interference construct (rRBr) specifically expressed in the root after early
embryogenesis (Wildwater et al., 2005). rRBr root apices develop additional columella
stem cell layers and display excessive LRC/epidermis and ground tissue stem cells. The
ectopic columella stem cell layers derive from a prolonged maintenance of stem cell
identity in the daughter cells of the original columella initials, resulting in turn in cell
proliferation and delayed differentiation (Wildwater et al., 2005). Interestingly, rRBr
roots do not display any defect in meristem size, meristem cell number and size of
differentiated cells, suggesting that stem cells are particularly sensitive to alterations in
RBR levels (Wildwater et al., 2005).
The SCARECROW (SCR) transcription factor seems to act in the pathway upstream of
RBR, as demonstrated by introducing rRBR in a scr loss-of-function mutant (scr-4).
scr-4,rRBr roots exhibit a further increase in stem cell proliferation and rRBR restores
the QC function lost in scr-4 (Wildwater et al., 2005). These results indicate that the
scr defects in QC specification are caused by uncontrolled RBR activity, which leads to
premature differentiation. Thus, suppressing the accumulation of RBR transcripts in scr
can rescue the QC defects. Additional support to this model comes from the
observation that induced overexpression of RBR drives the rapid differentiation of the
stem cell pool (Wildwater et al., 2005). Based on these results, Wildwater et al. (2005)
proposed that RBR action might occur through SCR-dependent inhibition of RBR in
the QC itself, which in turn would lead to a cell-non-autonomous effect on stem cells.
Moreover, interference with the transcript levels of the other upstream and
Chapter I
24
downstream components of the RBR pathway similarly affect the number of columella
stem cells (Wildwater et al., 2005). Overexpression of factors promoting cell division,
such as CycD and E2Fa, leads to an accumulation of columella stem cells. In contrast,
the constitutive expression of the cyclin-dependent-kinase inhibitor KRP2, which
prevents cell-cycle progression, results in the loss of columella stem cells.
In conclusion, the RBR pathway seems to directly influence the differentiation of stem
cells and their proximal daughters and it thus plays an important role in root meristem
patterning and development.
4. Conclusions
Proper root development depends on the activity of the root apical meristem, In this
tissue, different internal and external signals are integrated in the regulation of cell
division and cell differentiation. An auxin gradient at the root tip provides positional
cues for the specification and the maintenance of the stem cell niche, which represents
a constant source of new cells recruited for root growth. The fine-tuning of auxin
distribution is controlled by the polar PM localisation of PIN proteins, which mediate
the cell-to-cell movement of auxin along precise directions in the different cell layers
of the root. Hence, root development hinges ultimately on the correct establishment
and maintenance of PIN polarity. Several factors like vesicle trafficking, protein
phosphorylation and the sterol composition of the PM have been implicated in the
coordination of PIN polarity. However, additional players might exist and certain
regulatory processes, such as the mechanism that maintains PIN polarity counteracting
lateral diffusion along the PM, remain to be elucidated.
The developmental program of the root requires the positional input provided by auxin
to be elaborated and translated into coordinated events of cell division and cell
differentiation. Certain cell-cycle regulators, such as HBT and RBR, influence the
correct proceeding of these two processes acting at different stages of the cell-cycle.
The function of both HBT and RBR is crucial for the mechanism of cell fate
determination in the root meristem. Thus, cell-cycle regulators play a significant role
in the tissue patterning of the root and ultimately in its correct development. Other
Chapter I
25
components of the cell-cycle machinery might be potentially involved in the proper
organisation of the root meristem and thus await identification.
Acknowledgements
I am grateful to R. Simon and to the editors of The International Journal of Developmental Biology for
the permission to reproduce the image of Figure 1, to F. Ditengou for art work and to F. Santos Schröter
and C. Becker for critical reading of the manuscript and helpful comments.
References
Abas L, Benjamins R, Malenica N, Paciorek T, Wisniewska J, Moulinier-Anzola JC, Sieberer T, Friml J,
Luschnig C (2006) Intracellular trafficking and proteolysis of the Arabidopsis auxin-efflux
facilitator PIN2 are involved in root gravitropism. Nat Cell Biol 8: 249-256
Aida M, Beis D, Heidstra R, Willemsen V, Blilou I, Galinha C, Nussaume L, Noh YS, Amasino R, Scheres
B (2004) The PLETHORA genes mediate patterning of the Arabidopsis root stem cell niche. Cell
119: 109-120
Bainbridge K, Guyomarc'h S, Bayer E, Swarup R, Bennett M, Mandel T, Kuhlemeier C (2008) Auxin
influx carriers stabilize phyllotactic patterning. Genes Dev 22: 810-823
Benjamins R, Quint A, Weijers D, Hooykaas P, Offringa R (2001) The PINOID protein kinase regulates
organ development in Arabidopsis by enhancing polar auxin transport. Development 128: 40574067
Benjamins R, Scheres B (2008) Auxin: the looping star in plant development. Annu Rev Plant Biol 59:
443-465
Benkova E, Michniewicz M, Sauer M, Teichmann T, Seifertova D, Jurgens G, Friml J (2003) Local,
efflux-dependent auxin gradients as a common module for plant organ formation. Cell 115: 591602
Bennett MJ, Marchant A, Green HG, May ST, Ward SP, Millner PA, Walker AR, Schulz B, Feldmann
KA (1996) Arabidopsis AUX1 gene: a permease-like regulator of root gravitropism. Science 273:
948-950
Bhalerao RP, Bennett MJ (2003) The case for morphogens in plants. Nat Cell Biol 5: 939-943
Blakeslee JJ, Bandyopadhyay A, Lee OR, Mravec J, Titapiwatanakun B, Sauer M, Makam SN, Cheng Y,
Bouchard R, Adamec J, Geisler M, Nagashima A, Sakai T, Martinoia E, Friml J, Peer WA,
Chapter I
26
Murphy AS (2007) Interactions among PIN-FORMED and P-glycoprotein auxin transporters in
Arabidopsis. Plant Cell 19: 131-147
Blilou I, Frugier F, Folmer S, Serralbo O, Willemsen V, Wolkenfelt H, Eloy NB, Ferreira PC, Weisbeek
P, Scheres B (2002) The Arabidopsis HOBBIT gene encodes a CDC27 homolog that links the
plant cell cycle to progression of cell differentiation. Genes Dev 16: 2566-2575
Blilou I, Xu J, Wildwater M, Willemsen V, Paponov I, Friml J, Heidstra R, Aida M, Palme K, Scheres B
(2005) The PIN auxin efflux facilitator network controls growth and patterning in Arabidopsis
roots. Nature 433: 39-44
Bogre L, Magyar Z, Lopez-Juez E (2008) New clues to organ size control in plants. Genome Biol 9: 226
Cambridge AP, and Morris, D.A. (1996) Transfer of exogenous auxin from the phloem to the polar auxin
transport pathway in pea (Pisum sativum L.) Planta 199: 583-588
Capron A, Okresz L, Genschik P (2003) First glance at the plant APC/C, a highly conserved ubiquitinprotein ligase. Trends Plant Sci 8: 83-89
Capron A, Serralbo O, Fulop K, Frugier F, Parmentier Y, Dong A, Lecureuil A, Guerche P, Kondorosi E,
Scheres B, Genschik P (2003) The Arabidopsis anaphase-promoting complex or cyclosome:
molecular and genetic characterization of the APC2 subunit. Plant Cell 15: 2370-2382
Cebolla A, Vinardell JM, Kiss E, Olah B, Roudier F, Kondorosi A, Kondorosi E (1999) The mitotic
inhibitor ccs52 is required for endoreduplication and ploidy-dependent cell enlargement in
plants. Embo J 18: 4476-4484
Cho M, Lee SH, Cho HT (2007) P-glycoprotein4 displays auxin efflux transporter-like action in
Arabidopsis root hair cells and tobacco cells. Plant Cell 19: 3930-3943
Christensen SK, Dagenais N, Chory J, Weigel D (2000) Regulation of auxin response by the protein
kinase PINOID. Cell 100: 469-478
de Jager SM, Maughan S, Dewitte W, Scofield S, Murray JA (2005) The developmental context of cellcycle control in plants. Semin Cell Dev Biol 16: 385-396
Dhonukshe P, Aniento F, Hwang I, Robinson DG, Mravec J, Stierhof YD, Friml J (2007) Clathrinmediated constitutive endocytosis of PIN auxin efflux carriers in Arabidopsis. Curr Biol 17: 520527
Dhonukshe P, Tanaka H, Goh T, Ebine K, Mahonen AP, Prasad K, Blilou I, Geldner N, Xu J, Uemura T,
Chory J, Ueda T, Nakano A, Scheres B, Friml J (2008) Generation of cell polarity in plants links
endocytosis, auxin distribution and cell fate decisions. Nature 456: 962-966
Di Laurenzio L, Wysocka-Diller J, Malamy JE, Pysh L, Helariutta Y, Freshour G, Hahn MG, Feldmann
KA, Benfey PN (1996) The SCARECROW gene regulates an asymmetric cell division that is
essential for generating the radial organization of the Arabidopsis root. Cell 86: 423-433
Dolan L, Janmaat K, Willemsen V, Linstead P, Poethig S, Roberts K, Scheres B (1993) Cellular
organisation of the Arabidopsis thaliana root. Development 119: 71-84
Chapter I
27
Ebel C, Mariconti L, Gruissem W (2004) Plant retinoblastoma homologues control nuclear proliferation
in the female gametophyte. Nature 429: 776-780
Eloy NB, Coppens F, Beemster GT, Hemerly AS, Ferreira PC (2006) The Arabidopsis anaphase promoting
complex (APC): regulation through subunit availability in plant tissues. Cell Cycle 5: 1957-1965
Fleming AJ (2006) Plant signalling: the inexorable rise of auxin. Trends Cell Biol 16: 397-402
Friml J, Benkova E, Blilou I, Wisniewska J, Hamann T, Ljung K, Woody S, Sandberg G, Scheres B,
Jurgens G, Palme K (2002) AtPIN4 mediates sink-driven auxin gradients and root patterning in
Arabidopsis. Cell 108: 661-673
Friml J, Vieten A, Sauer M, Weijers D, Schwarz H, Hamann T, Offringa R, Jurgens G (2003) Effluxdependent auxin gradients establish the apical-basal axis of Arabidopsis. Nature 426: 147-153
Friml J, Wisniewska J, Benkova E, Mendgen K, Palme K (2002) Lateral relocation of auxin efflux
regulator PIN3 mediates tropism in Arabidopsis. Nature 415: 806-809
Friml J, Yang X, Michniewicz M, Weijers D, Quint A, Tietz O, Benjamins R, Ouwerkerk PB, Ljung K,
Sandberg G, Hooykaas PJ, Palme K, Offringa R (2004) A PINOID-dependent binary switch in
apical-basal PIN polar targeting directs auxin efflux. Science 306: 862-865
Fulop K, Tarayre S, Kelemen Z, Horvath G, Kevei Z, Nikovics K, Bako L, Brown S, Kondorosi A,
Kondorosi E (2005) Arabidopsis anaphase-promoting complexes: multiple activators and wide
range of substrates might keep APC perpetually busy. Cell Cycle 4: 1084-1092
Galinha C, Hofhuis H, Luijten M, Willemsen V, Blilou I, Heidstra R, Scheres B (2007) PLETHORA
proteins as dose-dependent master regulators of Arabidopsis root development. Nature 449:
1053-1057
Galweiler L, Guan C, Muller A, Wisman E, Mendgen K, Yephremov A, Palme K (1998) Regulation of
polar auxin transport by AtPIN1 in Arabidopsis vascular tissue. Science 282: 2226-2230
Geisler M, Blakeslee JJ, Bouchard R, Lee OR, Vincenzetti V, Bandyopadhyay A, Titapiwatanakun B, Peer
WA, Bailly A, Richards EL, Ejendal KF, Smith AP, Baroux C, Grossniklaus U, Muller A,
Hrycyna CA, Dudler R, Murphy AS, Martinoia E (2005) Cellular efflux of auxin catalyzed by the
Arabidopsis MDR/PGP transporter AtPGP1. Plant J 44: 179-194
Geldner N, Anders N, Wolters H, Keicher J, Kornberger W, Muller P, Delbarre A, Ueda T, Nakano A,
Jurgens G (2003) The Arabidopsis GNOM ARF-GEF mediates endosomal recycling, auxin
transport, and auxin-dependent plant growth. Cell 112: 219-230
Geldner N, Friml J, Stierhof YD, Jurgens G, Palme K (2001) Auxin transport inhibitors block PIN1
cycling and vesicle trafficking. Nature 413: 425-428
Grebe M, Xu J, Mobius W, Ueda T, Nakano A, Geuze HJ, Rook MB, Scheres B (2003) Arabidopsis sterol
endocytosis involves actin-mediated trafficking via ARA6-positive early endosomes. Curr Biol
13: 1378-1387
Gutierrez C (2005) Coupling cell proliferation and development in plants. Nat Cell Biol 7: 535-541
Chapter I
28
Heisler MG, Ohno C, Das P, Sieber P, Reddy GV, Long JA, Meyerowitz EM (2005) Patterns of auxin
transport and gene expression during primordium development revealed by live imaging of the
Arabidopsis inflorescence meristem. Curr Biol 15: 1899-1911
Helariutta Y, Fukaki H, Wysocka-Diller J, Nakajima K, Jung J, Sena G, Hauser MT, Benfey PN (2000)
The SHORT-ROOT gene controls radial patterning of the Arabidopsis root through radial
signaling. Cell 101: 555-567
Ingram GC, Waites R (2006) Keeping it together: co-ordinating plant growth. Curr Opin Plant Biol 9: 1220
Inze D (2005) Green light for the cell cycle. Embo J 24: 657-662
Iwakawa H, Shinmyo A, Sekine M (2006) Arabidopsis CDKA;1, a cdc2 homologue, controls proliferation
of generative cells in male gametogenesis. Plant J 45: 819-831
Jaillais Y, Fobis-Loisy I, Miege C, Rollin C, Gaude T (2006) AtSNX1 defines an endosome for auxincarrier trafficking in Arabidopsis. Nature 443: 106-109
Jakoby M, Schnittger A (2004) Cell cycle and differentiation. Curr Opin Plant Biol 7: 661-669
Kleine-Vehn J, Dhonukshe P, Sauer M, Brewer PB, Wisniewska J, Paciorek T, Benkova E, Friml J (2008)
ARF GEF-dependent transcytosis and polar delivery of PIN auxin carriers in Arabidopsis. Curr
Biol 18: 526-531
Kleine-Vehn J, Leitner J, Zwiewka M, Sauer M, Abas L, Luschnig C, Friml J (2008) Differential
degradation of PIN2 auxin efflux carrier by retromer-dependent vacuolar targeting. Proc Natl
Acad Sci U S A 105: 17812-17817
Kwee HS, Sundaresan V (2003) The NOMEGA gene required for female gametophyte development
encodes the putative APC6/CDC16 component of the Anaphase Promoting Complex in
Arabidopsis. Plant J 36: 853-866
Lammens T, Boudolf V, Kheibarshekan L, Zalmas LP, Gaamouche T, Maes S, Vanstraelen M, Kondorosi
E, La Thangue NB, Govaerts W, Inze D, De Veylder L (2008) Atypical E2F activity restrains
APC/CCCS52A2 function obligatory for endocycle onset. Proc Natl Acad Sci U S A 105: 1472114726
Li M, Zhang P (2009) The function of APC/CCdh1 in cell cycle and beyond. Cell Div 4: 2
Liu J, Zhang Y, Qin G, Tsuge T, Sakaguchi N, Luo G, Sun K, Shi D, Aki S, Zheng N, Aoyama T, Oka A,
Yang W, Umeda M, Xie Q, Gu H, Qu LJ (2008) Targeted degradation of the cyclin-dependent
kinase inhibitor ICK4/KRP6 by RING-type E3 ligases is essential for mitotic cell cycle
progression during Arabidopsis gametogenesis. Plant Cell 20: 1538-1554
Luschnig C, Gaxiola RA, Grisafi P, Fink GR (1998) EIR1, a root-specific protein involved in auxin
transport, is required for gravitropism in Arabidopsis thaliana. Genes Dev 12: 2175-2187
Chapter I
29
Marchant A, Bhalerao R, Casimiro I, Eklof J, Casero PJ, Bennett M, Sandberg G (2002) AUX1 promotes
lateral root formation by facilitating indole-3-acetic acid distribution between sink and source
tissues in the Arabidopsis seedling. Plant Cell 14: 589-597
Maughan SC, Murray JA, Bogre L (2006) A greenprint for growth: signalling the pattern of proliferation.
Curr Opin Plant Biol 9: 490-495
Mayer U, Torres Ruiz, R.A., Berleth, T., Misera, S., Jürgens, G. (1991) Mutations affecting body
organization in the Arabidopsis embryo. Nature 353: 402-407
Men S, Boutte Y, Ikeda Y, Li X, Palme K, Stierhof YD, Hartmann MA, Moritz T, Grebe M (2008) Steroldependent endocytosis mediates post-cytokinetic acquisition of PIN2 auxin efflux carrier
polarity. Nat Cell Biol 10: 237-244
Michniewicz M, Zago MK, Abas L, Weijers D, Schweighofer A, Meskiene I, Heisler MG, Ohno C, Zhang
J, Huang F, Schwab R, Weigel D, Meyerowitz EM, Luschnig C, Offringa R, Friml J (2007)
Antagonistic regulation of PIN phosphorylation by PP2A and PINOID directs auxin flux. Cell
130: 1044-1056
Mockaitis K, Estelle M (2008) Auxin receptors and plant development: a new signaling paradigm. Annu
Rev Cell Dev Biol 24: 55-80
Mravec J, Kubes M, Bielach A, Gaykova V, Petrasek J, Skupa P, Chand S, Benkova E, Zazimalova E,
Friml J (2008) Interaction of PIN and PGP transport mechanisms in auxin distributiondependent development. Development 135: 3345-3354
Mravec J, Skupa P, Bailly A, Hoyerova K, Krecek P, Bielach A, Petrasek J, Zhang J, Gaykova V, Stierhof
YD, Dobrev PI, Schwarzerova K, Rolcik J, Seifertova D, Luschnig C, Benkova E, Zazimalova E,
Geisler M, Friml J (2009) Subcellular homeostasis of phytohormone auxin is mediated by the
ER-localized PIN5 transporter. Nature 459: 1136-1140
Muller A, Guan C, Galweiler L, Tanzler P, Huijser P, Marchant A, Parry G, Bennett M, Wisman E,
Palme K (1998) AtPIN2 defines a locus of Arabidopsis for root gravitropism control. Embo J 17:
6903-6911
Murphy AS, Hoogner KR, Peer WA, Taiz L (2002) Identification, purification, and molecular cloning of
N-1-naphthylphthalmic acid-binding plasma membrane-associated aminopeptidases from
Arabidopsis. Plant Physiol 128: 935-950
Ottenschlager I, Wolff P, Wolverton C, Bhalerao RP, Sandberg G, Ishikawa H, Evans M, Palme K (2003)
Gravity-regulated differential auxin transport from columella to lateral root cap cells. Proc Natl
Acad Sci U S A 100: 2987-2991
Paciorek T, Zazimalova E, Ruthardt N, Petrasek J, Stierhof YD, Kleine-Vehn J, Morris DA, Emans N,
Jurgens G, Geldner N, Friml J (2005) Auxin inhibits endocytosis and promotes its own efflux
from cells. Nature 435: 1251-1256
Chapter I
30
Pan J, Fujioka S, Peng J, Chen J, Li G, Chen R (2009) The E3 ubiquitin ligase SCFTIR1/AFB and
membrane sterols play key roles in auxin regulation of endocytosis, recycling, and plasma
membrane accumulation of the auxin efflux transporter PIN2 in Arabidopsis thaliana. Plant Cell
21: 568-580
Paponov IA, Teale WD, Trebar M, Blilou I, Palme K (2005) The PIN auxin efflux facilitators:
evolutionary and functional perspectives. Trends Plant Sci 10: 170-177
Peer WA, Bandyopadhyay A, Blakeslee JJ, Makam SN, Chen RJ, Masson PH, Murphy AS (2004)
Variation in expression and protein localization of the PIN family of auxin efflux facilitator
proteins in flavonoid mutants with altered auxin transport in Arabidopsis thaliana. Plant Cell
16: 1898-1911
Perez-Perez JM, Serralbo O, Vanstraelen M, Gonzalez C, Criqui MC, Genschik P, Kondorosi E, Scheres B
(2008) Specialization of CDC27 function in the Arabidopsis thaliana anaphase-promoting
complex (APC/C). Plant J 53: 78-89
Peters JM (2006) The anaphase promoting complex/cyclosome: a machine designed to destroy. Nat Rev
Mol Cell Biol 7: 644-656
Petersson SV, Johansson AI, Kowalczyk M, Makoveychuk A, Wang JY, Moritz T, Grebe M, Benfey PN,
Sandberg G, Ljung K (2009) An auxin gradient and maximum in the Arabidopsis root apex
shown by high-resolution cell-specific analysis of IAA distribution and synthesis. Plant Cell 21:
1659-1668
Petrasek J, Mravec J, Bouchard R, Blakeslee JJ, Abas M, Seifertova D, Wisniewska J, Tadele Z, Kubes M,
Covanova M, Dhonukshe P, Skupa P, Benkova E, Perry L, Krecek P, Lee OR, Fink GR, Geisler
M, Murphy AS, Luschnig C, Zazimalova E, Friml J (2006) PIN proteins perform a rate-limiting
function in cellular auxin efflux. Science 312: 914-918
Quint M, Gray WM (2006) Auxin signaling. Curr Opin Plant Biol 9: 448-453
Rashotte AM, Brady SR, Reed RC, Ante SJ, Muday GK (2000) Basipetal auxin transport is required for
gravitropism in roots of Arabidopsis. Plant Physiol 122: 481-490
Raven J (1975) Transport of indolacetic acid in plant cells in relation to pH and electrical potential
gradients, and its ignificaance for polar IAA transport. New Phytologist 74: 163-172
Raven PH, Evert, R.F., and Eichhorn, S.E. (1999) The Root: Structure and Development. In Biology of
Plants, Sixth Edition. (New York: W.H. Freeman): pp. 589-609
Reinhardt D, Pesce ER, Stieger P, Mandel T, Baltensperger K, Bennett M, Traas J, Friml J, Kuhlemeier C
(2003) Regulation of phyllotaxis by polar auxin transport. Nature 426: 255-260
Rubery PH, Sheldrake AR (1974) Carrier-mediated auxin transport. Planta 118: 101-121
Sabatini S, Beis D, Wolkenfelt H, Murfett J, Guilfoyle T, Malamy J, Benfey P, Leyser O, Bechtold N,
Weisbeek P, Scheres B (1999) An auxin-dependent distal organizer of pattern and polarity in the
Arabidopsis root. Cell 99: 463-472
Chapter I
31
Sabatini S, Heidstra R, Wildwater M, Scheres B (2003) SCARECROW is involved in positioning the stem
cell niche in the Arabidopsis root meristem. Genes Dev 17: 354-358
Sachs T (1981) The control of patterned differentiation of vascular tissues. Adv. Bot. Res. 9: 151-262
Sauer M, Balla J, Luschnig C, Wisniewska J, Reinohl V, Friml J, Benkova E (2006) Canalization of auxin
flow by Aux/IAA-ARF-dependent feedback regulation of PIN polarity. Genes Dev 20: 29022911
Scarpella E, Marcos D, Friml J, Berleth T (2006) Control of leaf vascular patterning by polar auxin
transport. Genes Dev 20: 1015-1027
Serralbo O, Perez-Perez JM, Heidstra R, Scheres B (2006) Non-cell-autonomous rescue of anaphasepromoting complex function revealed by mosaic analysis of HOBBIT, an Arabidopsis CDC27
homolog. Proc Natl Acad Sci U S A 103: 13250-13255
Shevell DE, Leu WM, Gillmor CS, Xia G, Feldmann KA, Chua NH (1994) EMB30 is essential for normal
cell division, cell expansion, and cell adhesion in Arabidopsis and encodes a protein that has
similarity to Sec7. Cell 77: 1051-1062
Stahl Y, Simon R (2005) Plant stem cell niches. Int J Dev Biol 49: 479-489
Steinmann T, Geldner N, Grebe M, Mangold S, Jackson CL, Paris S, Galweiler L, Palme K, Jurgens G
(1999) Coordinated polar localization of auxin efflux carrier PIN1 by GNOM ARF GEF. Science
286: 316-318
Stepanova AN, Robertson-Hoyt J, Yun J, Benavente LM, Xie DY, Dolezal K, Schlereth A, Jurgens G,
Alonso JM (2008) TAA1-mediated auxin biosynthesis is essential for hormone crosstalk and
plant development. Cell 133: 177-191
Sugimoto-Shirasu K, Roberts K (2003) "Big it up": endoreduplication and cell-size control in plants. Curr
Opin Plant Biol 6: 544-553
Swarup K, Benkova E, Swarup R, Casimiro I, Peret B, Yang Y, Parry G, Nielsen E, De Smet I, Vanneste S,
Levesque MP, Carrier D, James N, Calvo V, Ljung K, Kramer E, Roberts R, Graham N,
Marillonnet S, Patel K, Jones JD, Taylor CG, Schachtman DP, May S, Sandberg G, Benfey P,
Friml J, Kerr I, Beeckman T, Laplaze L, Bennett MJ (2008) The auxin influx carrier LAX3
promotes lateral root emergence. Nat Cell Biol 10: 946-954
Swarup R, Friml J, Marchant A, Ljung K, Sandberg G, Palme K, Bennett M (2001) Localization of the
auxin permease AUX1 suggests two functionally distinct hormone transport pathways operate in
the Arabidopsis root apex. Genes Dev 15: 2648-2653
Swarup R, Kargul J, Marchant A, Zadik D, Rahman A, Mills R, Yemm A, May S, Williams L, Millner P,
Tsurumi S, Moore I, Napier R, Kerr ID, Bennett MJ (2004) Structure-function analysis of the
presumptive Arabidopsis auxin permease AUX1. Plant Cell 16: 3069-3083
Tanaka H, Dhonukshe P, Brewer PB, Friml J (2006) Spatiotemporal asymmetric auxin distribution: a
means to coordinate plant development. Cell Mol Life Sci 63: 2738-2754
Chapter I
32
Teale WD, Paponov IA, Palme K (2006) Auxin in action: signalling, transport and the control of plant
growth and development. Nat Rev Mol Cell Biol 7: 847-859
Teh OK, Moore I (2007) An ARF-GEF acting at the Golgi and in selective endocytosis in polarized plant
cells. Nature 448: 493-496
Terasaka K, Blakeslee JJ, Titapiwatanakun B, Peer WA, Bandyopadhyay A, Makam SN, Lee OR, Richards
EL, Murphy AS, Sato F, Yazaki K (2005) PGP4, an ATP binding cassette P-glycoprotein,
catalyzes auxin transport in Arabidopsis thaliana roots. Plant Cell 17: 2922-2939
Titapiwatanakun B, Blakeslee JJ, Bandyopadhyay A, Yang H, Mravec J, Sauer M, Cheng Y, Adamec J,
Nagashima A, Geisler M, Sakai T, Friml J, Peer WA, Murphy AS (2009) ABCB19/PGP19
stabilises PIN1 in membrane microdomains in Arabidopsis. Plant J 57: 27-44
van den Berg C, Willemsen V, Hendriks G, Weisbeek P, Scheres B (1997) Short-range control of cell
differentiation in the Arabidopsis root meristem. Nature 390: 287-289
Vandepoele K, Raes J, De Veylder L, Rouze P, Rombauts S, Inze D (2002) Genome-wide analysis of core
cell cycle genes in Arabidopsis. Plant Cell 14: 903-916
Vanneste S, Friml J (2009) Auxin: a trigger for change in plant development. Cell 136: 1005-1016
Vieten A, Vanneste S, Wisniewska J, Benkova E, Benjamins R, Beeckman T, Luschnig C, Friml J (2005)
Functional redundancy of PIN proteins is accompanied by auxin-dependent cross-regulation of
PIN expression. Development 132: 4521-4531
Weijers D, Sauer M, Meurette O, Friml J, Ljung K, Sandberg G, Hooykaas P, Offringa R (2005)
Maintenance of embryonic auxin distribution for apical-basal patterning by PIN-FORMEDdependent auxin transport in Arabidopsis. Plant Cell 17: 2517-2526
Weinberg RA (1995) The retinoblastoma protein and cell cycle control. Cell 81: 323-330
Wildwater M, Campilho A, Perez-Perez JM, Heidstra R, Blilou I, Korthout H, Chatterjee J, Mariconti L,
Gruissem W, Scheres B (2005) The RETINOBLASTOMA-RELATED gene regulates stem cell
maintenance in Arabidopsis roots. Cell 123: 1337-1349
Willemsen V, Friml J, Grebe M, van den Toorn A, Palme K, Scheres B (2003) Cell polarity and PIN
protein positioning in Arabidopsis require STEROL METHYLTRANSFERASE1 function. Plant
Cell 15: 612-625
Willemsen V, Wolkenfelt H, de Vrieze G, Weisbeek P, Scheres B (1998) The HOBBIT gene is required
for formation of the root meristem in the Arabidopsis embryo. Development 125: 521-531
Wisniewska J, Xu J, Seifertova D, Brewer PB, Ruzicka K, Blilou I, Rouquie D, Benkova E, Scheres B,
Friml J (2006) Polar PIN localization directs auxin flow in plants. Science 312: 883
Xu J, Hofhuis H, Heidstra R, Sauer M, Friml J, Scheres B (2006) A molecular framework for plant
regeneration. Science 311: 385-388
Yang Y, Hammes UZ, Taylor CG, Schachtman DP, Nielsen E (2006) High-affinity auxin transport by the
AUX1 influx carrier protein. Curr Biol 16: 1123-1127
Chapter I
33
Young LM, Evans ML, Hertel R (1990) Correlations between gravitropic curvature and auxin movement
across gravistimulated roots of Zea mays. Plant Physiol 92: 792-796
Zappel NF, Panstruga R (2008) Heterogeneity and lateral compartmentalization of plant plasma
membranes. Curr Opin Plant Biol 11: 632-640
- Chapter II -
Discrete distribution of AtPIN2 in
plasma membrane domains
Chapter II
36
Abstract
The phytohormone auxin plays a pivotal role in the development of several plant
tissues and organs. Differences in local auxin levels are generated by the polar transport
of auxin, which in turn depends on the asymmetric distribution of PIN efflux carriers
on one side of the cell. The lateral diffusion of PIN proteins within the plasma
membrane might potentially dissipate their polar localisation and requires therefore an
antagonising mechanism. However, such a mechanism has so far remained elusive. In
this study, we investigated how the maintenance of PIN2 polarity in Arabidopsis root
cells is achieved. We show that PIN2 localises to discrete domains within the plasma
membrane measuring approximately 400 nm in diameter. The clustered distribution of
PIN2 was not altered by cytoskeleton depolymerisation or by changes in the membrane
lipid composition. FRAP (Fluorescence Recovery After Photobleaching) experiments
and time-series observations documented the very low lateral motility of these clusters,
indicating that they occupy stable plasma membrane positions. PIN2 domains displayed
an association with the cell wall upon plasmolysis of epidermal cells. These PIN2-cell
wall association sites did not correspond to plasmodesmata positions, as demonstrated
by the absence of colocalisation between PIN2 domains and a plasmodesmata marker.
Our data provide new insights into the mechanism contributing to the retention of
PIN2 asymmetric distribution and suggest that the cell wall might play a key role in it.
Introduction
Auxin is a major plant hormone coordinating fundamental processes in plant
development,
like
embryogenesis,
organogenesis
and
growth
responses
to
environmental stimuli (Reinhardt et al., 2000; Bhalerao et al., 2002; Friml et al., 2002;
Esmon et al., 2006; Weijers et al., 2006). Besides its long-distance bidirectional
transport through the phloem, free auxin can be transported from cell-to-cell in a
strictly unidirectional manner (reviewed by(Kramer and Bennett, 2006; Robert and
Friml, 2009). This polar transport provides positional cues required for the specification
of tissue patterns, thus revealing the morphogenic function of auxin (Sabatini et al.,
1999; Benkova et al., 2003; Friml et al., 2003; Heisler et al., 2005). Members of the PIN
Chapter II
37
protein family, differentially localised on one side of the cell, act as auxin efflux carriers
and determine the direction of auxin flow (Galweiler et al., 1998; Muller et al., 1998).
Together with PINs, the AUX family of auxin influx carriers and the ABCB-type
multidrug resistance P-glycoproteins (MDR/PGP) also play an important role in auxin
cell-to-cell movement (Bennett et al., 1996; Geisler et al., 2005).
In Arabidopsis thaliana roots, PIN2 is localised to the apical side (towards the roothypocotyl junction) of epidermal and mature cortical cells and to the basal side
(towards the root apex) of young cortical cells in the meristematic zone (Muller et al.,
1998; Blilou et al., 2005). Upon gravistimulation, PIN2 mediates the asymmetric flow of
auxin from the root tip towards the elongation zone, promoting its accumulation on
the lower side of the root. This in turn enables the differential growth of the root and
the realignment to the gravity vector (Chen et al., 1998; Ottenschlager et al., 2003).
The polar distribution of PIN proteins seems to be regulated by continuous rounds of
endocytosis and vesicle recycling to the proper side of the cell (Steinmann et al., 1999;
Geldner et al., 2001; Geldner et al., 2003). This hypothesis is mainly based on the
effects of the fungal toxin brefeldin A (BFA), which inhibits the activity of certain
ARF-GEFs (guanine-nucleotide exchange factors for ADP-ribosylation factor GTPases).
BFA treatment thus affects the recycling of vesicles from endosomes to the plasma
membrane (PM) and results in the accumulation of internalised PINs in intracellular
compartments, the so-called BFA bodies. As apically localised PIN2 in epidermal cells
appears partially resistant to BFA treatments (Geldner et al., 2003; Kleine-Vehn et al.,
2008), its recycling might be mediated by BFA-insensitive ARF-GEFs. In addition, a
functional actin cytoskeleton is required for the intracellular dynamics of auxin carriers
and for the efficient transport of auxin from cell-to-cell (Dhonukshe et al., 2008).
Nevertheless, PIN polar localisation is independent of actin (Rahman et al., 2007) and
microtubules (Geldner et al., 2001).
During cell division PIN2 is targeted to the cell plate. At the end of cytokinesis it gets
removed from one daughter membrane in an endocytic process requiring a correct
sterol composition of cell membranes (Men et al., 2008). This observation indicates a
role for sterols in PIN2 polarity establishment after cytokinesis. Animal cell membranes
Chapter II
38
present a lateral organisation in dynamic lipid microdomains, called rafts, which are
enriched in sterols and sphingolipids (Simons and Ikonen, 1997). Lipid rafts have been
proposed to cluster specific proteins involved in various cellular processes, thus
providing a compartmentalisation to their activity. In the last years, the existence of
analogous lipid microdomains has been proven also for plant membranes and several
proteins were shown to co-purify with membrane fractions enriched in sterols and
sphingolipids (Mongrand et al., 2004; Borner et al., 2005). For few of these proteins the
biochemical association with lipid microdomains was supported by the localisation in
discrete PM domains (Grossmann et al., 2006; Raffaele et al., 2009). So far however, a
localisation of PIN2 in distinct PM domains has not been shown.
Here, we describe the specific targeting of PIN2 to discrete domains along the PM of
Arabidopsis epidermal and cortical root cells. We investigate the role of the
cytoskeleton and of the membrane lipid composition in the definition of this
heterogeneous distribution, showing that PIN2 domains are stable under different
conditions and display slow dynamics. Although PIN2 domains do not colabel with a
plasmodesmata marker, our results indicate a possible interaction between PIN2 and
the cell wall. This might in turn provide an anchoring mechanism for PIN2 at fixed
positions along the PM.
Material and methods
Plant material and growth conditions
Arabidopsis thaliana (L.) Heynh. (Col-0) was used as wild type. pPIN2:PIN2-GFP was
generated by insertion of CATGFP into PIN2 coding sequence at position 1436 from
ATG and its expression was driven by a 1.3 kb promoter region upstream of the PIN2
gene (Wolff, P.; unpublished data). The functionality of the fusion protein was verified
by complementation of the eir1-1 (pin2) agravitropic phenotype (Muller et al., 1998).
All the other fluorescent and mutant lines were kindly donated: pAUX1::AUX1-YFP
by M.J. Bennett, 35S::TMK1-GFP by A.B. Bleecker, 35S::GFP by O. Voinnet,
pAHA2::AHA2-GFP by A.T. Fuglsang, pSTM::P30-GFP(1x) by P.C. Zambryski, smt1orc
by V. Willemsen and cpi1-1 by M. Grebe.
Chapter II
39
Seeds were surface sterilised for 15 min with a solution of 5% w/v calcium hypochlorite
and 0.02% Triton X-100. After 3 washes in sterile water, they were left to dry under
sterile conditions. Seeds were sown on plates containing 1% w/v sucrose, half-strength
MS salts (Duchefa) and 12 g/l agar-agar (Roth) (pH 5.8). After two days of vernalisation
at 4°C in darkness, plates were transferred to a growth chamber (16h light/8h darkness,
22°C) for seed germination and were maintained in a vertical position.
Immunocytochemistry
For whole-mount immunolocalisations of PIN2 in non plasmolysed root cells, four days
old seedlings were fixed with 3% (w/v) paraformaldehyde and 0.02% Triton X-100 in
MTSB (pH 7.0) for 45 min and washed three times with dH2O. The subsequent steps
were performed in an InsituPro VS robot (Intavis). Briefly, tissue permeabilisation was
achieved by 30 min incubation in 0.15% (w/v) driselase (Sigma) and 0.15% (w/v)
macerozyme (Sigma) in 10 mM MES (pH 5.3) at 37°C, followed by four washes in
MTSB and two subsequent treatments of 20 min each with 10% (v/v) DMSO, 3% (v/v)
Nonidet P40 (Fluka) in MTSB. After five washes in MTSB, blocking was performed
with 3% BSA (Carl Roth, Germany) in MTSB for one hour. Guinea pig anti-PIN2
(Ditengou et al., 2008) primary antibody (1:1000) in 3% BSA in MTSB was applied for 4
hours at RT, followed by seven washes in MTSB. Goat anti-guinea pig A555-conjugated
(1:600) secondary antibody (Invitrogen) was applied for 3 hours at RT, followed by ten
washes in MTSB. Samples were mounted in Prolong Gold antifade reagent (Molecular
Probes).
Fixation protocol was modified for plasmolysed samples. Seedlings were incubated for
30 min in half strength MS liquid medium before they were transferred to 0.5 M
sorbitol (Sigma) for 20 min of plasmolysis. Subsequently, a 15 min incubation in 50%
ethanol and 0.5 M sorbitol was followed by 30 min treatment in pure ethanol at -20°C.
Ethanol was then replaced with 3% paraformaldehyde (in MTSB) in a stepwise manner
(85%, 70%, 50%, 30% and 15% EtOH), changing solution every 5 min on ice. Samples
were finally incubated for additional 20 min in 3% paraformaldehyde and washed
three times with dH2O. Subsequent steps were performed as described above.
Chapter II
40
Chemical treatments
For latrunculin B, oryzalin and MβCD treatments, 4 days old seedlings were incubated
for the indicated time in half-strength MS, 1% (w/v) sucrose medium buffered at pH
5.8 with 50 mM MES (Sigma), containing the respective chemical. Latrunculin B
(Biomol) and oryzalin (Duchefa) were diluted from respectively 1.26 mM and 50 mM
DMSO stocks. For control experiments an equivalent amount of DMSO was added to
the liquid medium. Myriocin (Sigma) was dissolved initially in methanol (1 mg/ml) and
diluted into molten agar just prior to gelling. Control media received the same amount
of methanol.
Plasmolysis of root epidermal cells was achieved by pre-incubating the seedlings for 30
min in MS liquid medium and subsequently transferring them to the same medium
supplemented with 0.5 M sorbitol.
Confocal microscopy
Images were acquired using a Zeiss LSM 5 DUO scanning microscope. GFP was excited
using a 488 nm argon laser line in conjunction with a 500-550 band-pass filter. A555
fluorochrome was detected in multitracking mode using a 561 nm laser line and a 575
long pass filter. Images were analyzed with the LSM image browser (Carl Zeiss
MicroImaging) and the Imaris software (Bitplane) was used for 3D reconstructions.
FRAP analyses
FRAP experiments were adapted after Men et al. (2008). Five days old seedlings were
pre-incubated for 30 min in liquid half strength MS medium, 1% sucrose, buffered to
pH 5.8 with 50 mM MES and containing 50 μM cycloheximide (Duchefa; 50 mM stock
in ethanol), 0.02% sodium azide (Roth; 2% stock in water) and 50 mM 2-deoxy-Dglucose (Sigma; 1M stock in water). Seedlings were subsequently mounted on a slide,
where a chamber filled with the same medium described above was created using a
frame of electrical tape and finally a coverslip was fixed on top. FRAP measurements
were performed using a Zeiss LSM 510 NLO inverted microscope, employing a watercorrected 63x objective NA = 1.2 and a 488 nM argon laser excitation. GFP emission
Chapter II
41
was detected with a 500-550 band pass filter. Using the Time Series mode of the Zeiss
LSM510 software, one pre-bleach image and one bleach scan were acquired, followed
by up to 25 iterations of post-bleach images. Per root, 3 μM square regions of interest
(ROIs) were selected for bleaching membranes of four epidermal cells. Within these
ROIs, photobleaching was achieved with 40 iterations of the 488 nm laser scanning at
100% main laser power and 40% transmittance. Pre- and post-bleach scans were
acquired using four frame averages in frame scanning mode, at 1024 x 1024 pixel
format and zoom factor three. Iterations of postbleach images were taken in intervals of
30 s for PIN2-GFP, but in 15 s intervals for TMK1-GFP, as this showed a faster
fluorescence recovery. Image analysis was performed using the FRAP Profiler plugin of
the Image J software (http://rsbweb.nih.gov/ij/). Fluorescence intensities of the
bleached ROIs and of the whole image were determined for pre- and post-bleach time
points. The values of the bleached ROIs measured for the bleach scan were used to
correct for background fluorescence. Mean values from the whole image were instead
employed to normalise for loss of fluorescence within the bleach and FRAP periods
caused by the initial photobleaching and excitation during post-bleach image
acquisition.
Results
PIN2 displays a discontinuous distribution along the PM
PIN2 has been shown to localise apically (towards the root-shoot junction) in
epidermal root cells and basally (towards the root tip) in young cortical cells. Careful
observations of epidermis and cortex cells of pPIN2::PIN2-GFP transgenic plants using
high resolution confocal microscopy revealed a not uniform PM labelling of the fusion
protein but rather a patchy distribution (Fig. 1A-B). Size measurements of 89 different
fluorescent membrane clusters from several images indicated that their diameter was
approximately 400 ± 60 nm. It should be noted that this size is likely overestimated due
to fluorescence diffusion, as reviewed by Hanson and Kohler (2001). In order to
exclude the possibility that the patchy arrangement of PIN2-GFP fusion protein could
be due to some properties of the GFP moiety, we performed immunolocalisations of the
Chapter II
42
Figure 1. PIN2 displays a patchy distribution along the PM. (A) PIN2-GFP polar localisation in epidermal
root cells. (B) Detail from (A). PIN2-GFP distribution along the PM exhibits clusters of fluorescent signal
(indicated by arrowheads). (C) Immunolocalisation of PIN2 in wild-type epidermal root cells. Note the
patchy pattern of the labelling (arrowheads). (D-E) Three-dimensional reconstruction of PIN2-GFP (left)
and AUX1-YFP (right) localisation in epidermal cells. AUX1-YFP shows a more homogenous
distribution along the PM in comparison with PIN2-GFP clustered fluorescence. Scale bars are 5 μm (A)
and 2 μm (B-C).
endogenous PIN2 protein in wild-type seedlings. A discontinuous distribution in
discrete PM domains was observed also for the labelled endogenous PIN2, confirming
the result obtained with PIN2-GFP (Fig. 1C). By contrast, the fluorescence of the auxin
influx carrier AUX1-YFP (Swarup et al., 2004) along the PM of epidermal root cells
appeared homogenously distributed and did not exhibit any clustering (Fig. 1E).
These observations indicate that PIN2 resides in PM subdomains of unknown nature.
Chapter II
43
PIN2 distribution in discrete PM domains is conserved under different conditions
Several factors might be responsible for PIN2 clustering along the PM. We first
addressed the question whether this pattern could be influenced by the organisation of
the cytoskeleton. Indeed, a cortical array of actin filaments and microtubules runs
along the cell periphery and clear evidence has been provided for the association of the
cytoskeleton with the PM (Vesk et al., 1996; Collings et al., 1998). Therefore, four days
old seedlings were treated either with latrunculin B (1 μM), an inhibitor of actin
polymerisation, or with oryzalin (1 μM), a compound that causes microtubules
depolymerisation. After 120 min of treatment, no effect was visible on the patchy
localisation of PIN2-GFP (Fig. 2A-C).
Figure 2. Cytoskeleton perturbations or changes in the membrane lipid composition do not affect PIN2
clustering in PM domains. (A-C) PIN2-GFP seedlings treated for two hours either with 1 μM latrunculin
B (B) or with 1 μM oryzalin (C) do not show any difference in PIN2 patchy labelling in comparison with
control seedlings treated for the same time with the equivalent volume of DMSO (A). (D-F) PIN2
immunolocalisation in wild-type (D), smt1orc (E) and cpi1-1 (F) seedlings. No difference in PIN2
clustering is visible between wt and the two mutants with an altered content of membrane sterols. (G-I)
PIN2-GFP domains are preserved in seedlings treated for one hour with 20 mM Methyl-β-Cyclodextrin
(MβCD) (H), which removes sterols from cell membranes, as well as in seedlings grown on 10 nm
Myriocin (I), an inhibitor of sphingolipid biosynthesis. As control for the Myriocin treatment, seedlings
were grown on medium containing an equivalent amount of methanol (G). No control is presented for
the MβCD treatment, as the chemical was dissolved in water. Scale bars are 2 μm (A-I).
Chapter II
44
A second possible explanation for the observed distribution of PIN2 could be the
presence of discrete lipid microdomains along the PM. These are small, highly
dynamic, sterol- and sphingolipid- enriched domains (Pike, 2006), whose existence has
been reported also in plants (reviewed by(Zappel and Panstruga, 2008). In order to test
this hypothesis, we used genetic and pharmacological approaches to interfere with the
composition of lipid microdomains. Immunodetection of PIN2 in smt1orc (Willemsen et
al., 2003) and cpi1-1 (Men et al., 2008), two mutants with impaired sterol biosynthesis
and consequently with altered membrane sterol levels, showed a non uniform labelling
of the PM, similarly to what observed in wild-type roots (Fig. 2D-F). When PIN2-GFP
seedlings were either grown on 10 nM myriocin, an inhibitor of sphingolipid
biosynthesis (Spassieva et al., 2002), or treated with methyl-β-cyclodextrin (20 mM), a
drug causing a reduction in membrane sterol contents (Roche et al., 2008), fluorescence
was still distributed in discrete domains (Fig. 2G-I). Thus, the sterol and sphingolipid
composition of the PM, which defines its organisation in lipid microdomains, does not
play a role in PIN2 clustering.
These results indicate that PIN2 localisation in PM domains is stable upon different
perturbations of the lipid membrane composition and the cytoskeleton.
PIN2-GFP is prevented from free movement along the PM
The high stability observed for PIN2 domains prompted us to investigate their
dynamics along the membrane. To this end, we first performed a time series
examination of PIN2 domain movements along one axis of the PM. Fig. 3A shows that
PIN2-GFP clusters retained their relative positions over a 2 min observation. Moreover,
none of the drug treatments mentioned above, perturbing the cytoskeleton or the lipid
membrane composition, increased the motility of PIN2 domains (data not shown).
Subsequently, we monitored the lateral diffusion of PIN2-GFP at the PM of root
epidermal cells in fluorescence recovery after photobleaching (FRAP) experiments. The
measurements were performed in the presence of protein biosynthesis and energy
inhibitors as described in Men et al. (2008), in order to monitor the lateral motility of
PIN2-GFP rather than biosynthetic or endocytic trafficking.
Chapter II
45
Figure 3. PIN2 clusters occupy stable positions along the PM and the free lateral motility of PIN2 is
prevented. (A) Time-series observation of PIN2-GFP domains. Images are presented in the “Rainbow”
mode of the LSM image browser (Carl Zeiss MicroImaging) to highlight PIN2-GFP clusters. The position
of each domain (indicated by arrowheads) is maintained over a 2 min observation. (B) FRAP analysis
after bleaching of 3 μm square ROIs (Regions Of Interest) at the PM of PIN2-GFP (top panel series) and
TMK1-GFP (bottom panel series) epidermal root cells. Images show root areas before photobleaching
(pre), immediately after photobleaching of selected ROIs (yellow squares) (0’’) and at different time
points during fluorescence recovery indicated in seconds (‘’). PIN2-GFP displays a slower fluorescence
recovery in comparison with TMK1-GFP. (C) Quantitative analysis of FRAP experiments corrected for
background fluorescence and loss-of-fluorescence intensity due to excitation during image acquisition.
The y-axes reports values of relative fluorescence calculated setting pre-bleach intensities to 1 and
immediate post-bleach intensities to 0. Data points and error bars represent averages ± SE from three
independent experiments involving a total of 12 ROIs. Note the reduced and slow fluorescence recovery
of PIN2-GFP when compared with that of TMK1-GFP. Scale bars are 2 μm (A-B).
The bleached area of the PM recovered only a small fraction of its initial fluorescence
(~0.2 of relative fluorescence), indicating that the PIN2-GFP molecules outside this area
Chapter II
46
remained preferentially anchored in their positions (Fig. 3B-C). Additionally, we
compared the behaviour of PIN2-GFP with that of TMK1-GFP, a fluorescent fusion of
TRANSMEMBRANE KINASE 1, which localises in a non-polar manner at the PM.
Although TMK1-GFP protein size is bigger than PIN2-GFP (150 kDa vs 96 kDa), its
fluorescence recovery was significantly faster (Fig. 3B-C), thus suggesting that PIN2 is
prevented from freely moving along the PM.
PIN2 patchy distribution might reflect interactions with the cell wall
The stability of PIN2 membrane domain distribution together with the observation
that its lateral motility is hindered might indicate the presence of some elements
anchoring PIN2 in its PM positions. A suitable candidate to accomplish this function is
the cell wall (CW), given its interactions with the PM (reviewed by(Oparka, 1994). In
order to investigate this hypothesis, PIN2-GFP root epidermal cells were plasmolysed
using 0.5 M sorbitol as an osmotic compound. PIN2-GFP labelled not only portions of
the retracted PM but also Hechtian strands and distinct punctuate structures along the
CW (Fig. 4A). To demonstrate the specificity of PIN2-GFP localisation after
plasmolysis, we employed as controls one line constitutively expressing the GFP
protein alone (35S::GFP;(Dalmay et al., 2000) and one line carrying a GFP fusion of the
PM-localised H(+)-ATPase 2 (pAHA2::AHA2-GFP). Plasmolysed epidermal cells of
these lines showed no fluorescence signal retained at the CW, indicating that the
characteristic behaviour observed for PIN2-GFP is not caused by the GFP moiety and is
not common to other PM proteins (Fig. 4B-C). Additionally, we developed a new
fixation protocol that permits to preserve the structure of plasmolysed cells during
immunocytochemistry experiments. This protocol is based on the incubation of
samples in ethanol and progressive substitution with paraformaldehyde. We could
confirm the localisation at the CW of plasmolysed epidermal cells also for the
immunolabelled endogenous PIN2 protein in wild-type roots (Fig. 4D). These results
suggest the existence of interactions between PIN2 and the CW.
Chapter II
47
Figure 4. PIN2 displays interactions with the CW upon plasmolysis. (A-D) Fluorescence (left panel) and
correspondent DIC (right panel) images of plasmolysed epidermal root cells. (A-C) PIN2-GFP (A),
AHA2-GFP (B) and 35S::GFP (C) seedlings pre-incubated for 30 min in 10 mM MES (pH 5.8) and
successively plasmolysed for 20 min in 0.5 M sorbitol. PIN2-GFP labels Hechtian strands and remains
partially localised at the CW (arrowheads) in plasmolysed cells. In contrast, the PM localised AHA2-GFP
and the cytoplasmic soluble GFP completely retract together with the protoplast upon plasmolysis and
no fluorescent labelling of the CW is visible (arrowheads). (D) Immunolocalisation of PIN2 in wild-type
roots indicates its persistence at the CW upon plasmolysis. Scale bars are 10 μm (A-D).
PIN2 domains and a plasmodesmata marker exhibit a different PM distribution pattern
Proteins associated with plasmodesmata (PD) exhibit a punctuate PM distribution and
are retained at the CW after plasmolysis (Baluska et al., 1999; Sagi et al., 2005; Raffaele
et al., 2009; Simpson et al., 2009). Given the similar localisation pattern and behaviour
of PIN2, we asked whether PIN2 domains correspond to PD sites.
Chapter II
48
Figure 5. PIN2 domains do not colocalise with plasmodesmata (PD). (A-F) Immunolocalisation of PIN2
(red) in unplasmolysed (A-C) and plasmolysed (D-F) epidermal cells of a pSTM::P30-GFP line presenting
fluorescently labelled (green) root plasmodesmata (PD). Fig. D, E and F display PIN2 and PD fluorescent
marker remaining at the cell wall upon plasmolysis with 0.5 M sorbitol. (A and D) Single plane images
show PIN2 localisation (left panel), GFP-labelled PD (middle panel) and merged channels (right panel).
(B and E) Three-dimensional reconstruction of a transversal side between two cells. (C and F)
Colocalisation analysis between PIN2 and GFP-labelled PD. Lower panels report the fluorescence
intensity profiles measured in the directions indicated by the arrows in the upper panels. Note the
absence of a clear correlation between PIN2 and GFP-labelled PD fluorescence patterns. Scale bars are
2 μm (A, C, D and F).
We immunolocalised PIN2 in a pSTM::P30-GFP line, in which root cells
plasmodesmata are fluorescently labelled by expression of the Tobacco mosaic virus
P30 movement protein translationally fused to GFP under the control of the SHOOT
MERISTEMLESS (STM) promoter (Kim et al., 2005). PIN2 membrane domains and the
PD marker did not colocalise (Fig. 5A-C). This was supported by the intensity profile
analysis of the two fluorescent signals, which did not display any kind of correlation
(Fig. 5C). Even in plasmolysed cells, where part of PIN2 domains and the PD marker
Chapter II
49
remained at the CW, no colocalisation between the two fluorescent signals was
observed (Fig. 5D-F). We therefore concluded that PIN2 occupies specific PM domains
distinct from plasmodesmata and that its interactions with the CW must rely on a
different mechanism.
Discussion
The polarity of auxin transport depends on the asymmetric distribution of PIN proteins
at the PM (Wisniewska et al., 2006). Although a number of factors involved in polar
PIN localisation have been identified (reviewed by(Feraru and Friml, 2008), the
mechanisms responsible for polarity establishment, maintenance and regulation remain
to be clarified. An important question is how polarity dissipation by protein lateral
diffusion along the PM is prevented. Cells of vertebrate animals possess anchored
protein complexes at the borders of PM polar domains, called tight junctions, which
prevent the lateral diffusion of proteins between different cell surfaces (Aijaz et al.,
2006). However, no analogous structures have been found so far in plants.
Here we report that PIN2 distribution along the PM, in contrast to the auxin influx
carrier AUX1, is not homogenous but clustered in domains of approximately 400 nm
size. Recently, a similar patchy distribution was shown for a fluorescent fusion of the
plant specific protein remorin (REM), which is associated with PM lipid microdomains
and localises also in plasmodesmata (Raffaele et al., 2009). The size of GFP-REM
clusters measured around 600 nm, a value comparable to PIN2 domains diameter,
although these sizes are likely to be overestimated due to fluorescence diffusion
(Hanson and Kohler, 2001). Similarly to PIN2 and REM in Arabidopsis, the hexoseproton symporter HUP1 shows a spotty distribution in the PM of the green alga
Chlorella kessleri (Grossmann et al., 2006). Furthermore, when HUP1-GFP fusion was
heterologously expressed in Saccaromyces cerevisiae, it distributed non-homogenously
and colocalised with proteins residing in specific PM subdomains collectively termed
“Membrane Compartment occupied by Can1” (MMC) (Grossmann et al., 2006). The
HUP1 protein extracted from Chlorella and the GFP fusion protein extracted from S.
cerevisiae purify with the fraction of detergent resistant membranes (DRMs), which
Chapter II
50
are considered the biochemical counterpart of lipid microdomains (Grossmann et al.,
2006). Interestingly, the 300 nm-large yeast MMC domains are preserved upon
perturbation of actin and microtubule cytoskeleton and display a very high stability
over time (Malinska et al., 2003, 2004). MMC properties are similar to what we show
here for PIN2 domains: they can not be disrupted by depolymerisation of the
cytoskeleton and their PM positions are retained over time. Thus, a parallel between
the similar distribution and properties of REM and HUP1 on one side and PIN2 on the
other might point towards a hypothetical localisation of PIN2 in lipid microdomains.
However, our results demonstrate that interfering with the membrane lipid
composition, either genetically or pharmacologically, does not lead to any change in
the stability of PIN2 domains. Moreover, although some studies have already addressed
this question (Titapiwatanakun et al., 2009), there is no evidence so far for the
co-purification of PIN2 with DRMs. Further experiments are therefore required to
decipher the nature of PIN2 domains.
In yeast, the restricted membrane localisation of some polar proteins to one side of the
cell can be maintained by endocytosis if their lateral diffusion is slow (Valdez-Taubas
and Pelham, 2003). PIN2 endocytosis is well documented (Dhonukshe et al., 2007) and
our FRAP experiments confirmed the results of Men et al. (2008), demonstrating a slow
motility for PIN2 along the PM. The model of polarity maintenance proposed for yeast
proteins could be therefore hypothesised also for PIN2. However the mechanism
preventing PIN2 free lateral diffusion has not yet been unravelled. While for some
yeast proteins this relies on the correct lateral organisation of the PM in lipid domains,
our data and previous observations (Men et al., 2008) show that changes in the
membrane content of sterols and sphingolipids, constituents of lipid microdomains, do
not affect PIN2 motility.
Alternatively, the CW could provide a suitable anchorage to prevent PIN2 free lateral
movement. Indeed, upon plasmolysis of epidermal root cells, PIN2 remained partially
associated with the CW, suggesting a possible interaction between the two. This
observation prompted us to examine whether PIN2 localises within plasmodesmata,
possibly accounting also for its patchy distribution. Our immunocytochemistry
Chapter II
51
experiments provided no evidence for a colocalisation of PIN2 with a PD marker,
neither in unplasmolysed nor in plasmolysed root cells. It was previously shown that
PM-CW attachment sites, displayed after plasmolysis, include not only PD but also
many other adhesion points (Vesk et al., 1996). It remains to be investigated at the
ultrastructural level whether PIN2 distribution is in any spatial relationship with PD.
These results leave however open the question about the role played by symplastic
routes in auxin transport along a cell file.
On the base of our data, we postulate a scenario where PIN2 might interact with the
CW either directly or indirectly through its association with proteins anchoring the
PM to the CW. This interaction might keep PIN2 domains anchored in their positions,
thus contributing to the maintenance of PIN2 polarity. This hypothesis is not in
contrast with the reported endocytosis of PIN2. Indeed, it has been shown that also cell
surface material, like pectins, is internalised in root cells (Baluska et al., 2002).
In summary, our results indicate that PIN2 resides in highly stable domains that are not
dependent on the lipid composition of the plasma membrane. PIN2 protein clusters
display a very low lateral motility, which might be due to their interactions with the
CW. Additional experiments are needed to clarify the nature of these interactions and
to prove their role in the maintenance of PIN2 polarity.
Acknowledgements
We thank O. Tietz for the introduction to the FRAP technique; X. Li for the production of PIN2
antibody; M.J. Bennett, A.B. Bleecker, O. Voinnet, A.T. Fuglsang, P.C. Zambryski, V. Willemsen and M.
Grebe for providing seeds of marker and mutant lines; R. Nitschke and the Life Imaging Center
(University of Freiburg) for the use of confocal microscopes; F. Santos Schröter, C. Neu, C. Becker and A.
Dovzhenko for critical reading of the manuscript and helpful comments.
References
Aijaz S, Balda MS, Matter K (2006) Tight junctions: molecular architecture and function. Int Rev Cytol
248: 261-298
Chapter II
52
Baluska F, Hlavacka A, Samaj J, Palme K, Robinson DG, Matoh T, McCurdy DW, Menzel D, Volkmann
D (2002) F-actin-dependent endocytosis of cell wall pectins in meristematic root cells. Insights
from brefeldin A-induced compartments. Plant Physiol 130: 422-431
Baluska F, Samaj J, Napier R, Volkmann D (1999) Maize calreticulin localizes preferentially to
plasmodesmata in root apex. Plant J 19: 481-488
Benkova E, Michniewicz M, Sauer M, Teichmann T, Seifertova D, Jurgens G, Friml J (2003) Local,
efflux-dependent auxin gradients as a common module for plant organ formation. Cell 115: 591602
Bennett MJ, Marchant A, Green HG, May ST, Ward SP, Millner PA, Walker AR, Schulz B, Feldmann
KA (1996) Arabidopsis AUX1 gene: a permease-like regulator of root gravitropism. Science 273:
948-950
Bhalerao RP, Eklof J, Ljung K, Marchant A, Bennett M, Sandberg G (2002) Shoot-derived auxin is
essential for early lateral root emergence in Arabidopsis seedlings. Plant J 29: 325-332
Blilou I, Xu J, Wildwater M, Willemsen V, Paponov I, Friml J, Heidstra R, Aida M, Palme K, Scheres B
(2005) The PIN auxin efflux facilitator network controls growth and patterning in Arabidopsis
roots. Nature 433: 39-44
Borner GH, Sherrier DJ, Weimar T, Michaelson LV, Hawkins ND, Macaskill A, Napier JA, Beale MH,
Lilley KS, Dupree P (2005) Analysis of detergent-resistant membranes in Arabidopsis. Evidence
for plasma membrane lipid rafts. Plant Physiol 137: 104-116
Chen R, Hilson P, Sedbrook J, Rosen E, Caspar T, Masson PH (1998) The arabidopsis thaliana
AGRAVITROPIC 1 gene encodes a component of the polar-auxin-transport efflux carrier. Proc
Natl Acad Sci U S A 95: 15112-15117
Collings DA, Asada T, Allen NS, Shibaoka H (1998) Plasma membrane-associated actin in bright yellow
2 tobacco cells. Evidence for interaction with microtubules. Plant Physiol 118: 917-928
Dalmay T, Hamilton A, Rudd S, Angell S, Baulcombe DC (2000) An RNA-dependent RNA polymerase
gene in Arabidopsis is required for posttranscriptional gene silencing mediated by a transgene
but not by a virus. Cell 101: 543-553
Dhonukshe P, Aniento F, Hwang I, Robinson DG, Mravec J, Stierhof YD, Friml J (2007) Clathrinmediated constitutive endocytosis of PIN auxin efflux carriers in Arabidopsis. Curr Biol 17: 520527
Dhonukshe P, Grigoriev I, Fischer R, Tominaga M, Robinson DG, Hasek J, Paciorek T, Petrasek J,
Seifertova D, Tejos R, Meisel LA, Zazimalova E, Gadella TW, Jr., Stierhof YD, Ueda T, Oiwa K,
Akhmanova A, Brock R, Spang A, Friml J (2008) Auxin transport inhibitors impair vesicle
motility and actin cytoskeleton dynamics in diverse eukaryotes. Proc Natl Acad Sci U S A 105:
4489-4494
Chapter II
53
Ditengou FA, Teale WD, Kochersperger P, Flittner KA, Kneuper I, van der Graaff E, Nziengui H, Pinosa
F, Li X, Nitschke R, Laux T, Palme K (2008) Mechanical induction of lateral root initiation in
Arabidopsis thaliana. Proc Natl Acad Sci U S A 105: 18818-18823
Esmon CA, Tinsley AG, Ljung K, Sandberg G, Hearne LB, Liscum E (2006) A gradient of auxin and
auxin-dependent transcription precedes tropic growth responses. Proc Natl Acad Sci U S A 103:
236-241
Feraru E, Friml J (2008) PIN polar targeting. Plant Physiol 147: 1553-1559
Friml J, Vieten A, Sauer M, Weijers D, Schwarz H, Hamann T, Offringa R, Jurgens G (2003) Effluxdependent auxin gradients establish the apical-basal axis of Arabidopsis. Nature 426: 147-153
Friml J, Wisniewska J, Benkova E, Mendgen K, Palme K (2002) Lateral relocation of auxin efflux
regulator PIN3 mediates tropism in Arabidopsis. Nature 415: 806-809
Galweiler L, Guan C, Muller A, Wisman E, Mendgen K, Yephremov A, Palme K (1998) Regulation of
polar auxin transport by AtPIN1 in Arabidopsis vascular tissue. Science 282: 2226-2230
Geisler M, Blakeslee JJ, Bouchard R, Lee OR, Vincenzetti V, Bandyopadhyay A, Titapiwatanakun B, Peer
WA, Bailly A, Richards EL, Ejendal KF, Smith AP, Baroux C, Grossniklaus U, Muller A,
Hrycyna CA, Dudler R, Murphy AS, Martinoia E (2005) Cellular efflux of auxin catalyzed by the
Arabidopsis MDR/PGP transporter AtPGP1. Plant J 44: 179-194
Geldner N, Anders N, Wolters H, Keicher J, Kornberger W, Muller P, Delbarre A, Ueda T, Nakano A,
Jurgens G (2003) The Arabidopsis GNOM ARF-GEF mediates endosomal recycling, auxin
transport, and auxin-dependent plant growth. Cell 112: 219-230
Geldner N, Friml J, Stierhof YD, Jurgens G, Palme K (2001) Auxin transport inhibitors block PIN1
cycling and vesicle trafficking. Nature 413: 425-428
Grossmann G, Opekarova M, Novakova L, Stolz J, Tanner W (2006) Lipid raft-based membrane
compartmentation of a plant transport protein expressed in Saccharomyces cerevisiae. Eukaryot
Cell 5: 945-953
Hanson MR, Kohler RH (2001) GFP imaging: methodology and application to investigate cellular
compartmentation in plants. J Exp Bot 52: 529-539
Heisler MG, Ohno C, Das P, Sieber P, Reddy GV, Long JA, Meyerowitz EM (2005) Patterns of auxin
transport and gene expression during primordium development revealed by live imaging of the
Arabidopsis inflorescence meristem. Curr Biol 15: 1899-1911
Kim I, Kobayashi K, Cho E, Zambryski PC (2005) Subdomains for transport via plasmodesmata
corresponding to the apical-basal axis are established during Arabidopsis embryogenesis. Proc
Natl Acad Sci U S A 102: 11945-11950
Kleine-Vehn J, Dhonukshe P, Sauer M, Brewer PB, Wisniewska J, Paciorek T, Benkova E, Friml J (2008)
ARF GEF-dependent transcytosis and polar delivery of PIN auxin carriers in Arabidopsis. Curr
Biol 18: 526-531
Chapter II
54
Kramer EM, Bennett MJ (2006) Auxin transport: a field in flux. Trends Plant Sci 11: 382-386
Malinska K, Malinsky J, Opekarova M, Tanner W (2003) Visualization of protein compartmentation
within the plasma membrane of living yeast cells. Mol Biol Cell 14: 4427-4436
Malinska K, Malinsky J, Opekarova M, Tanner W (2004) Distribution of Can1p into stable domains
reflects lateral protein segregation within the plasma membrane of living S. cerevisiae cells. J
Cell Sci 117: 6031-6041
Men S, Boutte Y, Ikeda Y, Li X, Palme K, Stierhof YD, Hartmann MA, Moritz T, Grebe M (2008) Steroldependent endocytosis mediates post-cytokinetic acquisition of PIN2 auxin efflux carrier
polarity. Nat Cell Biol 10: 237-244
Mongrand S, Morel J, Laroche J, Claverol S, Carde JP, Hartmann MA, Bonneu M, Simon-Plas F, Lessire
R, Bessoule JJ (2004) Lipid rafts in higher plant cells: purification and characterization of Triton
X-100-insoluble microdomains from tobacco plasma membrane. J Biol Chem 279: 36277-36286
Muller A, Guan C, Galweiler L, Tanzler P, Huijser P, Marchant A, Parry G, Bennett M, Wisman E,
Palme K (1998) AtPIN2 defines a locus of Arabidopsis for root gravitropism control. Embo J 17:
6903-6911
Oparka KJ (1994) Plasmolysis: new insights into an old process. New Phytologist 126: 571-591
Ottenschlager I, Wolff P, Wolverton C, Bhalerao RP, Sandberg G, Ishikawa H, Evans M, Palme K (2003)
Gravity-regulated differential auxin transport from columella to lateral root cap cells. Proc Natl
Acad Sci U S A 100: 2987-2991
Pike LJ (2006) Rafts defined: a report on the Keystone Symposium on Lipid Rafts and Cell Function. J
Lipid Res 47: 1597-1598
Raffaele S, Bayer E, Lafarge D, Cluzet S, Retana SG, Boubekeur T, Leborgne-Castel N, Carde JP,
Lherminier J, Noirot E, Satiat-Jeunemaitre B, Laroche-Traineau J, Moreau P, Ott T, Maule AJ,
Reymond P, Simon-Plas F, Farmer EE, Bessoule JJ, Mongrand S (2009) Remorin, a Solanaceae
Protein Resident in Membrane Rafts and Plasmodesmata, Impairs Potato virus X Movement.
Plant Cell
Rahman A, Bannigan A, Sulaman W, Pechter P, Blancaflor EB, Baskin TI (2007) Auxin, actin and growth
of the Arabidopsis thaliana primary root. Plant J 50: 514-528
Reinhardt D, Mandel T, Kuhlemeier C (2000) Auxin regulates the initiation and radial position of plant
lateral organs. Plant Cell 12: 507-518
Robert HS, Friml J (2009) Auxin and other signals on the move in plants. Nat Chem Biol 5: 325-332
Roche Y, Gerbeau-Pissot P, Buhot B, Thomas D, Bonneau L, Gresti J, Mongrand S, Perrier-Cornet JM,
Simon-Plas F (2008) Depletion of phytosterols from the plant plasma membrane provides
evidence for disruption of lipid rafts. Faseb J 22: 3980-3991
Chapter II
55
Sabatini S, Beis D, Wolkenfelt H, Murfett J, Guilfoyle T, Malamy J, Benfey P, Leyser O, Bechtold N,
Weisbeek P, Scheres B (1999) An auxin-dependent distal organizer of pattern and polarity in the
Arabidopsis root. Cell 99: 463-472
Sagi G, Katz A, Guenoune-Gelbart D, Epel BL (2005) Class 1 reversibly glycosylated polypeptides are
plasmodesmal-associated proteins delivered to plasmodesmata via the golgi apparatus. Plant Cell
17: 1788-1800
Simons K, Ikonen E (1997) Functional rafts in cell membranes. Nature 387: 569-572
Simpson C, Thomas C, Findlay K, Bayer E, Maule AJ (2009) An Arabidopsis GPI-anchor plasmodesmal
neck protein with callose binding activity and potential to regulate cell-to-cell trafficking. Plant
Cell 21: 581-594
Spassieva SD, Markham JE, Hille J (2002) The plant disease resistance gene Asc-1 prevents disruption of
sphingolipid metabolism during AAL-toxin-induced programmed cell death. Plant J 32: 561-572
Steinmann T, Geldner N, Grebe M, Mangold S, Jackson CL, Paris S, Galweiler L, Palme K, Jurgens G
(1999) Coordinated polar localization of auxin efflux carrier PIN1 by GNOM ARF GEF. Science
286: 316-318
Swarup R, Kargul J, Marchant A, Zadik D, Rahman A, Mills R, Yemm A, May S, Williams L, Millner P,
Tsurumi S, Moore I, Napier R, Kerr ID, Bennett MJ (2004) Structure-function analysis of the
presumptive Arabidopsis auxin permease AUX1. Plant Cell 16: 3069-3083
Titapiwatanakun B, Blakeslee JJ, Bandyopadhyay A, Yang H, Mravec J, Sauer M, Cheng Y, Adamec J,
Nagashima A, Geisler M, Sakai T, Friml J, Peer WA, Murphy AS (2009) ABCB19/PGP19
stabilises PIN1 in membrane microdomains in Arabidopsis. Plant J 57: 27-44
Valdez-Taubas J, Pelham HR (2003) Slow diffusion of proteins in the yeast plasma membrane allows
polarity to be maintained by endocytic cycling. Curr Biol 13: 1636-1640
Vesk PA, Vesk M, Gunning BES (1996) Field emission scanning electron microscopy of microtubule
arrays in higher plant cells. Protoplasma 195: 168-182
Weijers D, Schlereth A, Ehrismann JS, Schwank G, Kientz M, Jurgens G (2006) Auxin triggers transient
local signaling for cell specification in Arabidopsis embryogenesis. Dev Cell 10: 265-270
Willemsen V, Friml J, Grebe M, van den Toorn A, Palme K, Scheres B (2003) Cell polarity and PIN
protein positioning in Arabidopsis require STEROL METHYLTRANSFERASE1 function. Plant
Cell 15: 612-625
Wisniewska J, Xu J, Seifertova D, Brewer PB, Ruzicka K, Blilou I, Rouquie D, Benkova E, Scheres B,
Friml J (2006) Polar PIN localization directs auxin flow in plants. Science 312: 883
Zappel NF, Panstruga R (2008) Heterogeneity and lateral compartmentalization of plant plasma
membranes. Curr Opin Plant Biol 11: 632-640
- Chapter III -
Effects of sphingolipid biosynthesis inhibition on
PIN2 polarity and root development in Arabidopsis
Chapter III
58
Abstract
Several cellular processes occurring at the level of biological membranes are essential
for plant development and plant response to environmental signals. The physical
properties of cell membranes and their organisation in distinct domains are defined by
differential membrane lipid compositions. The cell-to-cell movement of auxin relies on
the plasma membrane localisation of efflux carriers belonging to the PIN protein
family. The membrane content of sterols has previously been shown to affect the
asymmetric distribution of some PIN proteins to one side of the cell. In this study, we
investigate the influence of membrane sphingolipid content on PIN polarity in
Arabidopsis roots. Inhibition of sphingolipid biosynthesis by the fungal antibiotic
myriocin caused major root growth defects and impaired the correct root graviresponse
by affecting the basipetal transport of auxin. PIN2 polarity was affected, with the
protein localising to both apical and basal sides of epidermal and cortical cells. This
altered distribution of PIN2 was related to a reduced rate of endocytosis, which
impaired the removal of the protein from one of the two daughter membranes of the
cell plate at the end of cytokinesis. Our data confirm the importance of a correct lipid
composition for the establishment of PIN2 polarity and highlight the role of
sphingolipids in this process.
Introduction
Local auxin maxima and gradients within plant tissues give rise to coordinated cellular
responses that determine plant development, growth and morphogenesis at every stage
of the life cycle (reviewed by(Bhalerao and Bennett, 2003; Tanaka et al., 2006;
Benjamins and Scheres, 2008; Vanneste and Friml, 2009). The differential distribution
of auxin is mainly achieved through its active directional movement from cell-to-cell
over short distances (reviewed by(Robert and Friml, 2009). This polar auxin transport
(PAT) is mediated by plasma membrane-based influx and efflux carriers, which belong
respectively to the AUX1/LIKE AUX1 (AUX1/LAX) family and to the PIN and ABCtype multidrug resistance P-glycoproteins (MDR/PGP) families (Bennett et al., 1996;
Galweiler et al., 1998; Muller et al., 1998; Noh et al., 2001; Swarup et al., 2001; Swarup
Chapter III
59
et al., 2008). The members of the PIN protein family accomplish a relevant function in
PAT, as they determine the direction of auxin flow through their asymmetric
localisation on a specific side of the cell (Wisniewska et al., 2006). In roots for example,
PIN2 localises apically in epidermal cells and mainly basally in cortical cells (Muller et
al., 1998). It thereby regulates the basipetal flux of auxin from the root tip towards the
elongation zone. This route of transport is fundamental to the proper bending of the
root upon gravistimulation. In accordance, mutations in the gene coding for PIN2
affect the correct realignment to the new gravity vector (Chen et al., 1998; Luschnig et
al., 1998; Muller et al., 1998; Utsuno et al., 1998).
The polar distribution of PIN proteins is controlled by their constitutive endocytosis
and recycling back to the cell surface (Geldner et al., 2001; Dhonukshe et al., 2007).
Several factors have been shown to influence the polar targeting of PIN proteins to the
plasma membrane (PM) (reviewed by(Feraru and Friml, 2008). Among these is the
lipid composition of the PM and in particular the correct content of sterols. Analysis of
two sterol-deficient mutants, orc and cyclopropylsterol isomerase1 (cpi1), revealed
major defects in the polar localisation of PIN proteins (Willemsen et al., 2003; Men et
al., 2008). In the case of cpi1, the asymmetric distribution of PIN2 is disrupted and the
protein localises to both the apical and the basal sides of the cell. During cell division
PIN2 is targeted to the cell plate and at the end of cytokinesis it is removed by
endocytosis from one of the two daughter membranes in order to maintain the polarity
of the mother cell in the newly formed cells (Men et al., 2008). The altered membrane
sterol content of cpi1 impairs this endocytosis step and PIN2 remains inserted into both
sides of the two daughter cells.
The importance of sterols for the correct establishment of PIN polarity raises the
question about the relation between PIN membrane localisation and lipid rafts. The last
have been recently defined as “small, highly dynamic, sterol- and sphingolipidenriched domains that compartmentalise cellular processes” (Pike, 2006). Similar lipid
microdomains have been found in plants as well and seem to play a role in an
increasing number of physiological processes (reviewed by(Zappel and Panstruga,
2008). Recently, it was shown that ABCB19 stabilises PIN1 in sterol- and sphingolipid-
Chapter III
60
enriched membrane fractions, so called DRMs for Detergent-Resistant Membranes
(Titapiwatanakun et al., 2009). However, the presence of PIN2 in such fractions, which
are considered the biochemical counterpart of lipid microdomains, has not been
reported so far.
Sphingolipids represent one of the major lipid components of plant PMs and are
concentrated in membrane microdomains (Yoshida and Uemura, 1986; Lynch and
Steponkus, 1987; Borner et al., 2005). In addition, sphingolipid-derived molecules can
act as cellular signals and induce programmed cell death in plants (Liang et al., 2003;
Wang et al., 2008). The fundamental structural unit common to all sphingolipids is
ceramide, which consists of a C18 long-chain base (LCB) linked to a fatty acid through
an amide linkage. This basic ceramide structure can be modified by differences in chain
length, methyl branching, number of hydroxyl groups and degree of unsaturation
(Sperling and Heinz, 2003). Besides their role as signalling molecules, ceramides serve
as precursors for the formation of more complex sphingolipids through the addition of
various glycosyl residues and other polar phosphate-containing headgroups (Liang et
al., 2003; Sperling and Heinz, 2003). The major complex sphingolipids reported in
higher plants include monoglucosylceramides and inositolphosphoceramides, which
contain respectively a glucose and a more polar inositolphosphate head group (Lynch
and Dunn, 2004). Analysis of different mutant lines with an abolished or reduced
activity of genes involved in sphingolipid biosynthesis or modification revealed an
essential role played by this class of membrane lipids in plant development (Zheng et
al., 2005; Chen et al., 2006; Teng et al., 2008; Beaudoin et al., 2009). For example, a
mutation in Arabidopsis thaliana LCB1 gene, coding for a subunit of serine
palmitoyltransferase, the enzyme catalysing the first step of sphingolipid biosynthesis,
was found to cause embryo lethality (Chen et al., 2006).
In this study, we investigate the role of sphingolipids into the process of PIN2 polarity
regulation. We show that inhibition of sphingolipid biosynthesis by myriocin, a fungal
antibiotic, affects root growth and graviresponse and impairs the establishment of PIN2
polarity after cytokinesis. Our data collectively confirm the essential function
accomplished by sphingolipids during plant development and demonstrate that this
Chapter III
61
class of lipids, similarly to sterols, plays an important part in the definition of plant cell
polarity.
Materials and methods
Plant material and growth conditions
Arabidopsis thaliana (L.) Heynh. (Col-0) was used as wild type. pPIN2::PIN2-GFP was
generated by insertion of CATGFP into PIN2 coding sequence at position 1436 from
ATG and its expression was driven by a 1.3 kb promoter region upstream of the PIN2
gene (Wolff, P.; unpublished data). The functionality of the fusion protein was verified
by the complementation of the eir1-1 (pin2) agravitropic phenotype (Muller et al.,
1998). The pDR5::GUS (Ulmasov et al., 1997), pDR5rev::GFP (Benkova et al., 2003) and
the pAUX1::AUX1-YFP (Swarup et al., 2004) lines were previously published. Seeds
were surface sterilised for 15 min. with a solution of 5% w/v calcium hypochlorite and
0.02% Triton X-100. After 3 washes in sterile water, they were left to dry under sterile
conditions. Seeds were sown on plates containing 1% w/v sucrose, half-strength MS
salts (Duchefa) and 12 g/l agar-agar (Roth) (pH 5.8). For myriocin (Sigma) containing
medium, the drug was dissolved initially in methanol (1 mg/ml) and diluted into
molten agar just prior to gelling. Control media received the same amount of methanol.
After two days of vernalisation at 4°C in darkness, plates were transferred to a growth
chamber (16h light/8h darkness, 22°C) for seed germination and were maintained in a
vertical position.
Root growth and gravitropism assays
Five days after germination induction, vertically positioned plates were scanned with a
CanonScan 9950F scanner and seedlings root length was measured from digital images
using the Image J software (http://rsbweb.nih.gov/ij/). For gravitropism assays, plants
were further incubated for 24 h in complete darkness maintaining the same growth
conditions. Next, plates were rotated clockwise through 90° and incubated for
additional 24 h in darkness. Successively, plates were scanned and the angle of
deviation from the gravity vector was determined at the root tip using the Image J
Chapter III
62
software. The angles of roots were grouped into twelve 30° sectors from -180° to +180°,
where a root completely realigned to the new gravity vector forms a 0° angle.
BFA treatment and immunocytochemistry
For monitoring PIN1 and PIN2 internalisation upon BFA treatment, five days old
seedlings were preincubated for 30 min with 50 μM cycloheximide (CHX) (Duchefa) in
half-strength MS salts, 1% (w/v) sucrose medium buffered at pH 5.8 with 50 mM MES
(Sigma). Subsequently, seedlings were transferred to the same medium containing
50 μM CHX and 10 μM BFA (Sigma). For myriocin-treated seedlings, 10 nM myriocin
was present in the medium throughout the experiment, while mock seedlings were
incubated in medium containing an equal amount of methanol. CHX was added from a
50 mM stock in ethanol and BFA from a 25 mM stock in DMSO. At the indicated timepoints seedlings were fixed with 3% (w/v) paraformaldehyde (Fluka) and 0.02% Triton
X-100 (Sigma) in MTSB buffer (pH 7.0) for 45 min and washed three times with dH2O.
For immunodetection of PIN1, PIN2, PIN3 and PIN4 in samples not treated with BFA,
five days old seedlings were directly transferred from the plate to the fixative solution.
Whole-mount immunocytochemistry was performed as described elsewhere (Chapter
II, p. 38) using an InsituPro VS robot (Intavis). The concentrations of primary
antibodies were 1:400 for rabbit anti-PIN1 (Galweiler et al., 1998), 1:1000 for guinea
pig anti-PIN2 (Ditengou et al., 2008), 1:400 for guinea pig anti-PIN3 (Men et al., 2008)
and 1:400 for rabbit anti-PIN4 (Friml et al., 2002). Different fluorescent Alexaconjugated secondary antibodies (Invitrogen) were employed at 1:600. Samples were
mounted in Prolong Gold antifade reagent containing DAPI (Molecular probes).
GUS staining
Five days old pDR5::GUS seedlings were fixed for 10’ in ice-cold 90% acetone and
subsequently incubated for 2.5 h at 37° in GUS staining solution (50 mM sodium
phosphate buffer, pH 7.0, 5 mM potassium ferrocyanide, 5 mM potassium ferricyanide,
0.1% Triton X-100 and 1 mg/ml X-Gluc). Seedlings were then rinsed in phosphate
buffer and transferred for 20 min to 100% ethanol for tissue clearing. After stepwise
Chapter III
63
rehydration in decreasing ethanol series (70%, 50% and 25%), samples were mounted
on slides with 50% glycerol for microscopy observations.
FM4-64 staining
Five days old seedlings were pulse-labelled for 10 min on ice with 50 μM FM4-64
(Invitrogen) in half-strength MS salts, 1% (w/v) sucrose medium buffered at pH 5.8
with 50 mM MES (Sigma). They were then washed twice and incubated for the
indicated time with the same medium. To monitor FM4-64 internalisation upon BFA
treatment, 10 μM BFA was added to the washing and incubation medium. For myriocin
treated samples, 10 nM myriocin was added to all the solutions of the experiment.
Confocal microscopy
Images were acquired using a Zeiss LSM 510 NLO confocal scanning microscope.
Excitation wavelengths were 488 nm (argon laser) for GFP and Alexa488-conjugated
antibodies, 514 nm for YFP and 543 nm (HeNe laser) for FM4-64 and Alexa555conjugated antibodies. Emission was detected at 500-550 nm for GFP and Alexa488conjugated antibodies, above 520 nm for YFP and above 575 nm for FM4-64 and
Alexa555-conjugated antibodies. DAPI was imaged using a 2-Photon module with
excitation at 730 nm and emission at 435-485 nm. All multi-labelling signals were
detected in multitracking mode to avoid fluorescence crosstalk. Images were analyzed
with the LSM image browser (Carl Zeiss MicroImaging).
FRAP analysis
FRAP experiments were performed as described elsewhere (Chapter II, p. 39). For
myriocin treated samples, 10 nM myriocin was added to the growth medium and to the
pre-incubation and mounting solutions.
Chapter III
64
Results
Inhibition of sphingolipid biosynthesis affects root growth and gravitropic response
The first step of sphingolipid biosynthesis is conserved among eukaryotes and involves
the condensation of palmitoyl-CoA and serine to form 3-ketosphinganine, the
precursor
of
long-chain
bases
(Fig.
1).
The
catalysing
enzyme,
serine
palmitoyltransferase (SPT) is the target of a potent antibiotic called myriocin, which
inhibits its activity in vitro at picomolar concentrations (Miyake et al., 1995) hence
affecting sphingolipid biosynthesis, also in plants (Spassieva et al., 2002). In order to
study the consequences of altered sphingolipid content on primary root growth, we
investigated the behaviour of seedlings grown on medium containing different
myriocin concentrations.
Figure 1. Biosynthesis of sphingolipid long chain bases and ceramides in plants. The first step of long
chain bases biosynthesis is the condensation of Palmitoyl Co-A and Serine to form 3-ketosphinganine.
The reaction is catalysed by Serine Palmitoyltransferase, which can be inhibited by the antibiotic
myriocin. 3-ketosphinganine is reduced to form sphinganine, the simplest long chain base. Addition of a
fatty-acyl chain (containing 16 to 26 carbon atoms) through an amide bond results in the production of
N-acyl-sphinganine, which can be further modified to form N-acyl-phytosphingosine. N-acylsphinganine and N-acyl-phytosphingosine can be both referred to as “ceramide”.
Chapter III
65
Figure 2. Inhibition of sphingolipid biosynthesis by myriocin impairs root growth and gravitropism. (A)
Five days old wild-type seedlings grown on medium supplemented with the indicated concentrations of
myriocin. The severity of primary root growth defects increases together with the drug concentration.
(B) Quantitative analysis of root gravitropic response for six days old seedlings grown on plates
containing the indicated concentrations of myriocin and rotated for 24 h to +135°. Root angles were
determined as deviations from 0° representing complete realignment to the gravity vector and were
grouped in twelve sectors of 30°. Bars represent the percentage of roots per sector in comparison to the
total number of roots analysed. Increasing concentrations of myriocin cause a progressive delay in root
tip reorientation. (C) Root length analysis of five days old seedlings grown on the indicated
concentrations of myriocin. Averages + SE are reported; n=3 and ten to twelve roots were used for each
experiment. Root length is significantly reduced at 5 nM or higher myriocin concentrations (Student’s ttest, P<0,001). (D-E) Expression of pDR5::GUS at the root tip of ungravistimulated five days old seedlings
grown on mock medium (D) and medium supplemented with 10 nM myriocin. Roots grown on
myriocin display an accumulation of expression for the auxin reporter. (F-G) Expression of
pDR5rev::GFP in gravistimulated (+90° for 2h) seedlings grown on mock medium (F) and medium
containing 10 nM myriocin (G). Arrows and arrowheads indicate the direction of the gravity vector and
the fluorescent signal at the bottom side of the root, respectively. In contrast to mock grown samples,
Chapter III
66
seedlings grown on myriocin failed to establish an asymmetric expression of the auxin reporter at the
bottom side of gravistimulated roots. Scale bars are 2 mm (A) and 20 μm (D-G).
A significant reduction in root length could be observed already at 5 nM myriocin and
was even stronger at higher concentrations of the drug (Fig. 2A and 2C). We asked
whether the inhibition of sphingolipid biosynthesis could as well have an effect on the
root response to gravistimulation. When rotated 90° for 24 hours in the dark, seedlings
grown on mock medium exhibited an almost complete realignment of the roots to the
new gravity vector (Fig. 2B). In contrast, 1 nM myriocin slightly altered root
gravitropic responses and plants grown at higher concentrations displayed a clear delay
in the realignment (Fig. 2B). It is known that auxin redistribution to the lower side of
gravistimulated roots plays a fundamental role in the correct gravisresponse
(Ottenschlager et al., 2003; Swarup et al., 2005). We therefore employed the auxin
response reporter lines pDR5::GUS and pDR5rev::GFP to monitor auxin gradients in
roots before and after gravistimulation. For comparison, seedlings were grown either
on mock medium or on medium supplemented with 10 nM myriocin, a concentration
clearly inhibiting root growth and root gravitropic response. When myriocin was
added, vertically grown seedlings displayed an accumulation of pDR5::GUS at the root
tip (Fig. 2D-E). After 2 hours of gravistimulation, roots grown on mock medium
presented an asymmetric expression of pDR5rev::GFP at their bottom flank (Fig. 2F), as
previously reported (Paciorek et al., 2005). In contrast, inhibition of sphingolipid
biosynthesis by myriocin impaired the formation of this asymmetric auxin gradient
(Fig. 2G). The accumulation of auxin at the root tip and the lack of redistribution upon
gravistimulation suggest the possibility of a defective basipetal auxin transport.
These results indicate that alterations in the sphingolipid content have a strong effect
on primary root growth and inhibit the correct root gravitropic response impairing
auxin redistribution.
Chapter III
67
PIN2 polarity is altered upon inhibition of sphingolipid biosynthesis
In order to identify the causes for the observed defects in auxin gradients before and
after gravistimulation, we analysed the localisation of several auxin influx and efflux
carriers in plants grown on 10 nM myriocin. The distribution of a YFP N-terminal
fusion of the influx carrier AUX1 did not display any difference between mock- and
myriocin-treated seedlings, as visualised by its PM labelling in stele, epidermal and
lateral root cap cells (Fig. 3K-P) Similarly, immunodetected PIN1, PIN3 and PIN4
exhibited mock-like localisation in treated roots (Fig. 3E-J). Only PIN2 distribution in
epidermis and cortex was clearly altered in seedlings grown on myriocin-containing
medium. Indeed, in contrast to the typical apical localisation in epidermal cells
displayed by control plants, we observed PIN2 immunolabelling also at the basal side of
the cells (Fig. 3A-D). Moreover, unfused cell plate structures labelled with PIN2 and
multinucleate cells, displayed by DAPI staining, revealed cytokinesis defects in several
cells (Fig. 3B and 3D). It is interesting to note that the obtained phenotype was very
similar to the effects of an altered membrane sterol composition in the cpi1-1 mutant
(Men et al., 2008).
Our data suggest that the specific defects observed for PIN2 localisation could account
for the described auxin accumulation at the root tip and for the lack of auxin
redistribution upon gravistimulation.
Chapter III
68
Figure 3. Inhibition of sphingolipid biosynthesis causes defects in PIN2 polarity and cytokinesis. (A-D)
Immunolocalisation of PIN2 (red) and DAPI staining (blue) in wild-type seedlings grown on mock
medium (A and C) and on medium supplemented with 10 nM myriocin (B and D). White and grey
arrowheads indicate respectively apical and basal localisation of PIN2. Arrows point at unfused cell plate
structures and asterisks mark multinucleate cells. (A) and (B) show epidermis (Ep) and cortex (Cx) cell
files. (C) and (D) represent epidermal cells. In contrast to samples grown on mock medium, seedlings
Chapter III
69
grown on 10 nM myriocin exhibit localisation of PIN2 to both apical and basal membranes of the same
cell and cytokinesis defects. (E-L) Immunolocalisation of PIN1 (green) (E and F), PIN4 (fuchsia) (G and
H) and PIN3 (pale blue) (I and J) and DAPI staining (blue) (E-H) in wild-type seedlings grown on mock
medium (E, G and I) and medium supplemented with 10 nM myriocin (F, H and J). Samples grown on
myriocin did not display any difference in the localisation of the three proteins. (K-P) Expression of
pAUX1::AUX1-YFP in the root tip (K and L) and protein localisation in epidermal (M and N) and lateral
root cap (O and P) cells of seedlings grown on mock medium (K, M and O) and medium containing
10 nM myriocin. No localisation defects are visible for samples grown on myriocin. Scale bars are 5 μm
(A-D and M-P), 10 μm (E-J) and 20 μm (K and L).
Altered sphingolipid content does not affect PIN2-GFP lateral diffusion
A possible explanation for the observed changes in PIN2 distribution was that
sphingolipid membrane composition influenced PIN2 lateral motility. To test this
hypothesis, we monitored the membrane lateral diffusion of PIN2-GFP fluorescent
fusion proteins upon sphingolipid biosynthesis inhibition.
Figure 4. Alterations in sphingolipid content do not affect PIN2 lateral diffusion. Quantitative analysis of
FRAP experiments for PIN2-GFP. A comparison between pPIN2::PIN2-GFP seedlings grown on mock
medium and on medium supplemented with 10 nM myriocin reveals no changes in PIN2-GFP lateral
motility upon inhibition of sphingolipid biosynthesis. Data were corrected for background fluorescence
and loss-of-fluorescence intensity due to excitation during image acquisition. The y-axis reports values of
relative fluorescence calculated setting pre-bleach intensities to 1 and immediate post-bleach intensities
Chapter III
70
to 0. Data points and error bars represent averages ± SE from three independent experiments involving a
total of 12 ROIs (Regions of Interest).
FRAP measurements were performed with seedlings grown on and incubated with
medium containing 10 nM myriocin during the experiment. A comparison between
mock and inhibitor-treated plants displayed no difference in the behaviour of PIN2GFP: in both cases the same fraction of the initial fluorescence was recovered with
comparable speed in photobleached regions of the PM (Fig. 4). These results indicate
that the changes in PIN2 polarity observed upon inhibition of sphingolipid biosynthesis
are not due to alterations in the lateral diffusion of the protein. Moreover, it can also be
concluded that the sphingolipid composition of the PM is not the factor determining
the slow lateral motility of PIN2 described elsewhere (Chapter II).
A reduced endocytosis rate caused by sphingolipid biosynthesis inhibition impairs
PIN2 polarity establishment
During cytokinesis PIN2 is targeted to the cell plate and it is subsequently removed by
endocytosis from one of the daughter membranes (Men et al., 2008). In seedlings with
an altered sphingolipid composition, PIN2 remained at the basal side of epidermal cells
after cytokinesis. Therefore, we examined whether this persistence could be due to
defects in the internalisation of the protein. The fungal toxin Brefeldin A (BFA)
interferes with vesicle-mediated protein recycling from endosomes to the PM. It has
been used as a tool to monitor endocytosis following the accumulation of internalised
material in endomembrane agglomerations called “BFA compartments” (Steinmann et
al., 1999). Plants were pre-treated for 30’ with 50 μM of the protein biosynthesis
inhibitor cycloheximide (CHX) and subsequently with 10 μM BFA and the same
concentration of CHX. This set up allowed monitoring the internalisation of PMlocalised PIN2 while preventing its de novo synthesis.
Chapter III
71
Figure 5. Inhibition of sphingolipid biosynthesis causes a general reduction of the endocytosis rate. (AD) Immunolocalisation of PIN2 (A and B) and PIN1 (C and D) in roots of wild-type seedlings grown on
mock medium (A and C) or on medium supplemented with 10 nM myriocin (B and D) and incubated for
the indicated time with 10 μM BFA. Samples grown on myriocin display a slower intracellular
accumulation of protein internalised from the PM. (E-H) Internalisation of FM4-64 at the indicated time
points in the absence (E and F) or in the presence (G and H) of 10 μM BFA in root cells of seedlings
grown on mock medium (E and G) and on medium supplemented with 10 nM myriocin (F and H).
Inhibition of sphingolipid biosynthesis by myriocin causes a delay in FM4-64 internalisation, as
Chapter III
72
visualised by the number and the size of fluorescent intracellular compartments. Scale bars are 5 μm (AH).
In plants grown on mock medium, immunodetected PIN2 progressively labelled
intracellular compartments upon BFA treatment (Fig. 5A). In contrast, 10 nM myriocin
added to the growth medium and to the BFA treatment, almost completely inhibited
the formation of PIN2-positive intracellular compartments, indicating a strong
reduction of the PIN2 internalisation rate (Fig. 5B). Next, we investigated whether this
slowing down of endocytosis caused by the inhibition of sphingolipid biosynthesis was
specific to PIN2 or common to other PM proteins and PM material. First, we examined
the internalisation of PIN1 upon BFA treatment. Similarly to PIN2, immunolabelled
PIN1 exhibited a reduced internalisation rate in myriocin-treated seedlings, with the
protein still persisting at the PM, while in control plants it was exclusively present in
intracellular compartments or had already been targeted for degradation (Fig. 5C-D).
We then monitored the internalisation of the general endocytic tracer FM4-64, which
is inserted into the bilayer of the PM upon pulse-labelling and subsequently
incorporated into endocytic vesicles. In the absence of BFA, control seedlings displayed
a faster appearance and a higher number of intracellular fluorescent compartments in
comparison with myriocin-treated plants (Fig. 5E-F). Similarly, in the presence of BFA,
growth and treatment of seedlings with myriocin clearly slowed down the formation of
BFA compartments and prevented the internalisation of the PM fluorescent signal (Fig.
5G-H). Thus, the inhibition of sphingolipid biosynthesis affects the general mechanism
of endocytosis and not only PIN2 internalisation.
In conclusion, our data indicate that membrane sphingolipid composition is
fundamental to a correct rate of endocytosis, in turn necessary for PIN2 polarity
establishment.
Chapter III
73
Discussion
Changes in the relative amounts of different lipid species in cell membranes affect
many developmental and physiological functions also in plants (Souter et al., 2002;
Bach et al., 2008; Dietrich et al., 2008; Men et al., 2008; Teng et al., 2008). In this study,
we investigated the role played by sphingolipids in root development. We employed an
inhibitor of sphingolipid biosynthesis, myriocin, and characterised its effects in a doseresponse manner. Arabidopsis seedlings grown on medium containing myriocin
displayed a significant reduction of root length at very low concentrations (nanomolar
range) of the drug. The strong effect of myriocin in Arabidopsis seedlings is comparable
to the severe growth defects observed for other eukaryotic cells treated with the drug
(Nakamura et al., 1996; Sun et al., 2000; Blank et al., 2005). A mutation in the
Arabidopsis gene coding for one of the two subunits of SPT (LCB1), lcb1-1, confers an
embryo lethal phenotype to homozygous plants and partial suppression of AtLCB1
expression by RNA interference causes a general reduction of plant growth (Chen et
al., 2006). Thus, the root growth defects obtained by pharmacological inhibition of SPT
with myriocin are in agreement with the phenotype of plants genetically impaired in
SPT activity.
Inhibition of sphingolipid biosynthesis by myriocin hindered the correct response of
roots upon gravistimulation. The delay in the realignment to the new gravity vector
was progressively higher with increasing concentrations of the drug. Expression of
pDR5::GUS and pDR5rev::GFP revealed an accumulation of auxin at the root tip before
gravistimulation and a failure in the redistribution of auxin to the lower side of
gravistimulated roots. This indicates a role for membrane sphingolipid composition in
the basipetal transport of auxin that mediates the proper response of roots to gravity
changes (Rashotte et al., 2000). Indeed, the localisation of PIN2 in epidermal and
cortical cells, which significantly contributes to this specific route of auxin transport,
was disturbed in roots grown on myriocin. The protein lost its asymmetric distribution
to one side of the cell, labelling both apical and basal sides and partially expanding its
localisation to the lateral membranes. Other PIN proteins and the auxin influx carrier
AUX1 did not display any change in their localisation, suggesting that the observed
Chapter III
74
variations in auxin distribution before and after gravistimulation are probably due to
the specific defects in PIN2 polarity.
Our FRAP experiments revealed that sphingolipid biosynthesis inhibition does not
affect PIN2 lateral motility along the PM. This excludes the possibility that PIN2
polarity defects might be caused by changes in the lateral diffusion of the protein,
when the sphingolipid content is reduced. Furthermore, it can be concluded that PIN2
slow motility does not depend on the sphingolipid composition of the PM. A similar
result had been obtained when the sterol content of the PM was reduced (Men et al.,
2008). Taken together, these two observations indicate that the lipid composition of the
PM is not the factor determining the speed of PIN2 lateral diffusion. A different
mechanism must exist for anchoring PIN2 proteins in their positions, preventing them
from free movement.
In roots grown on myriocin, the polarity defects of PIN2 could have been brought
about by the observed reduction in the endocytosis rate. Acquisition of PIN2
asymmetric distribution requires the internalisation of the protein from one of the two
daughter membranes of the cell plate at the end of cytokinesis (Men et al., 2008). PIN2
endocytosis was considerably reduced upon inhibition of sphingolipid biosynthesis.
The persistence of the protein to both sides of the cell might therefore be caused by its
failed internalisation from one daughter membrane of the cell plate at the end of
cytokinesis. The reduced rate of endocytosis was not specific to PIN2 but common to
PIN1 and to the general endocytic tracer FM4-64. The defects in PIN1 internalisation
indicate that myriocin is able to penetrate up to the inner cell layers of the root.
Noteworthy, the absence of changes in PIN1 localisation suggests that PIN1 is less
sensitive than PIN2 to an altered content of sphingolipids. Defects in endocytic
membrane trafficking have been previously reported for mutant plants with an altered
content of very-long-chain fatty acids in sphingolipid molecules (Zheng et al., 2005).
Our data confirm the importance of a correct sphingolipid composition of cell
membranes for endocytosis.
The effects of sphingolipid biosynthesis inhibition on the endocytosis rate and on PIN2
polarity, together with the observed cytokinesis defects, are very similar to the
Chapter III
75
situation in the sterol-deficient mutant cpi1-1 (Men et al., 2008). Moreover, the defects
in root growth and gravitropism caused by inhibition of sphingolipid biosynthesis are
comparable to the effects of a reduced content of sterols in cell membranes (Men et al.,
2008; Pan et al., 2009). Sphingolipids and sterols are both enriched in lipid
microdomains (Borner et al., 2005). Thus, it could be hypothesised that independent
alterations in the membrane content of each of the two lipid classes bring about the
same destabilisation of lipid microdomains. This would in turn affect cellular processes
like endocytosis, fundamental to correct root development. Indeed, sphingolipids and
sterols are both instrumental in the formation of lipid microdomains, as measured by
the recovery of DRMs from membranes with different lipid compositions (Laloi et al.,
2007; Roche et al., 2008). However, an alternative explanation for the similar effects
obtained by independent variations in the content of the two lipid species might be the
possible cross-talk between the two biosynthetic pathways. The inhibition of one
pathway could reduce the flow of intermediates through the other. In this case, it
would remain unclear whether it is the reduction of sphingolipids or rather that of
sterols that primarily determines the defects in root development. A number of
sphingolipid-derived molecules have been shown to function as cellular signals for
induction of programmed cell death (Liang et al., 2003; Wang et al., 2008). Although
we can not rule out the possibility that inhibition of sphingolipid biosynthesis by
myriocin induces an accumulation of these signalling molecules, the endocytosis
defects point towards a primary effect of the drug on the sphingolipid composition of
the PM. Our current experiments aim at obtaining a lipid profile of plants treated with
myriocin in order to identify the lipid species subjected to changes in their content.
PIN2 is absent from DRMs (unpublished data from Cho, Y.J., Teale, W., Palme, K.;
Titapiwatanakun et al., 2009) and the speed of PIN2 lateral diffusion is unaffected
when the membrane content of sterols and sphingolipids is altered. The observed
defects in PIN2 polarity therefore cannot be directly linked to changes in the
organisation of lipid microdomains. It is possible that independent variations in sterols
and sphingolipids contents, either affecting or not lipid microdomain clustering, lead to
Chapter III
76
the same loss of membrane integrity. These would in turn explain the general defects
in endocytic trafficking.
In summary, our data reveal a significant role for sphingolipids in the process of PIN2
polarity establishment and demonstrate their function in basipetal auxin transport.
Additional studies are required to determine the relation between sphingolipids and
sterols in the process of cell polarity acquisition.
Acknowledgements
We are grateful to X. Li for the production of PIN antibodies, to M.J. Bennet for providing seeds of the
pAUX1::AUX1-YFP line, to R. Nitschke and the Life Imaging Center (University of Freiburg) for the use
of confocal microscopes, to P. Kochersperger for the assistance with GUS staining and to Y.J. Cho for
sharing unpublished results. We wish to thank in particular F. Santos Schröter, C. Becker and A.
Dovzhenko for critical reading of the manuscript and helpful comments.
References
Bach L, Michaelson LV, Haslam R, Bellec Y, Gissot L, Marion J, Da Costa M, Boutin JP, Miquel M,
Tellier F, Domergue F, Markham JE, Beaudoin F, Napier JA, Faure JD (2008) The very-longchain hydroxy fatty acyl-CoA dehydratase PASTICCINO2 is essential and limiting for plant
development. Proc Natl Acad Sci U S A 105: 14727-14731
Beaudoin F, Wu X, Li F, Haslam RP, Markham JE, Zheng H, Napier JA, Kunst L (2009) Functional
characterization of the Arabidopsis beta-ketoacyl-coenzyme A reductase candidates of the fatty
acid elongase. Plant Physiol 150: 1174-1191
Benjamins R, Scheres B (2008) Auxin: the looping star in plant development. Annu Rev Plant Biol 59:
443-465
Benkova E, Michniewicz M, Sauer M, Teichmann T, Seifertova D, Jurgens G, Friml J (2003) Local,
efflux-dependent auxin gradients as a common module for plant organ formation. Cell 115: 591602
Bennett MJ, Marchant A, Green HG, May ST, Ward SP, Millner PA, Walker AR, Schulz B, Feldmann
KA (1996) Arabidopsis AUX1 gene: a permease-like regulator of root gravitropism. Science 273:
948-950
Bhalerao RP, Bennett MJ (2003) The case for morphogens in plants. Nat Cell Biol 5: 939-943
Chapter III
77
Blank N, Schiller M, Gabler C, Kalden JR, Lorenz HM (2005) Inhibition of sphingolipid synthesis impairs
cellular activation, cytokine production and proliferation in human lymphocytes. Biochem
Pharmacol 71: 126-135
Borner GH, Sherrier DJ, Weimar T, Michaelson LV, Hawkins ND, Macaskill A, Napier JA, Beale MH,
Lilley KS, Dupree P (2005) Analysis of detergent-resistant membranes in Arabidopsis. Evidence
for plasma membrane lipid rafts. Plant Physiol 137: 104-116
Chen M, Han G, Dietrich CR, Dunn TM, Cahoon EB (2006) The essential nature of sphingolipids in
plants as revealed by the functional identification and characterization of the Arabidopsis LCB1
subunit of serine palmitoyltransferase. Plant Cell 18: 3576-3593
Chen R, Hilson P, Sedbrook J, Rosen E, Caspar T, Masson PH (1998) The arabidopsis thaliana
AGRAVITROPIC 1 gene encodes a component of the polar-auxin-transport efflux carrier. Proc
Natl Acad Sci U S A 95: 15112-15117
Dhonukshe P, Aniento F, Hwang I, Robinson DG, Mravec J, Stierhof YD, Friml J (2007) Clathrinmediated constitutive endocytosis of PIN auxin efflux carriers in Arabidopsis. Curr Biol 17: 520527
Dietrich CR, Han G, Chen M, Berg RH, Dunn TM, Cahoon EB (2008) Loss-of-function mutations and
inducible RNAi suppression of Arabidopsis LCB2 genes reveal the critical role of sphingolipids
in gametophytic and sporophytic cell viability. Plant J 54: 284-298
Ditengou FA, Teale WD, Kochersperger P, Flittner KA, Kneuper I, van der Graaff E, Nziengui H, Pinosa
F, Li X, Nitschke R, Laux T, Palme K (2008) Mechanical induction of lateral root initiation in
Arabidopsis thaliana. Proc Natl Acad Sci U S A 105: 18818-18823
Feraru E, Friml J (2008) PIN polar targeting. Plant Physiol 147: 1553-1559
Friml J, Benkova E, Blilou I, Wisniewska J, Hamann T, Ljung K, Woody S, Sandberg G, Scheres B,
Jurgens G, Palme K (2002) AtPIN4 mediates sink-driven auxin gradients and root patterning in
Arabidopsis. Cell 108: 661-673
Galweiler L, Guan C, Muller A, Wisman E, Mendgen K, Yephremov A, Palme K (1998) Regulation of
polar auxin transport by AtPIN1 in Arabidopsis vascular tissue. Science 282: 2226-2230
Geldner N, Friml J, Stierhof YD, Jurgens G, Palme K (2001) Auxin transport inhibitors block PIN1
cycling and vesicle trafficking. Nature 413: 425-428
Laloi M, Perret AM, Chatre L, Melser S, Cantrel C, Vaultier MN, Zachowski A, Bathany K, Schmitter
JM, Vallet M, Lessire R, Hartmann MA, Moreau P (2007) Insights into the role of specific lipids
in the formation and delivery of lipid microdomains to the plasma membrane of plant cells.
Plant Physiol 143: 461-472
Liang H, Yao N, Song JT, Luo S, Lu H, Greenberg JT (2003) Ceramides modulate programmed cell death
in plants. Genes Dev 17: 2636-2641
Chapter III
78
Luschnig C, Gaxiola RA, Grisafi P, Fink GR (1998) EIR1, a root-specific protein involved in auxin
transport, is required for gravitropism in Arabidopsis thaliana. Genes Dev 12: 2175-2187
Lynch DV, Dunn TM (2004) An introduction to plant sphingolipids and a review of recent advances in
understanding their metabolism and function. New Phytologist 161: 677-702
Lynch DV, Steponkus PL (1987) Plasma Membrane Lipid Alterations Associated with Cold Acclimation
of Winter Rye Seedlings (Secale cereale L. cv Puma). Plant Physiol 83: 761-767
Men S, Boutte Y, Ikeda Y, Li X, Palme K, Stierhof YD, Hartmann MA, Moritz T, Grebe M (2008) Steroldependent endocytosis mediates post-cytokinetic acquisition of PIN2 auxin efflux carrier
polarity. Nat Cell Biol 10: 237-244
Miyake Y, Kozutsumi Y, Nakamura S, Fujita T, Kawasaki T (1995) Serine palmitoyltransferase is the
primary target of a sphingosine-like immunosuppressant, ISP-1/myriocin. Biochem Biophys Res
Commun 211: 396-403
Muller A, Guan C, Galweiler L, Tanzler P, Huijser P, Marchant A, Parry G, Bennett M, Wisman E,
Palme K (1998) AtPIN2 defines a locus of Arabidopsis for root gravitropism control. Embo J 17:
6903-6911
Nakamura S, Kozutsumi Y, Sun Y, Miyake Y, Fujita T, Kawasaki T (1996) Dual roles of sphingolipids in
signaling of the escape from and onset of apoptosis in a mouse cytotoxic T-cell line, CTLL-2. J
Biol Chem 271: 1255-1257
Noh B, Murphy AS, Spalding EP (2001) Multidrug resistance-like genes of Arabidopsis required for auxin
transport and auxin-mediated development. Plant Cell 13: 2441-2454
Ottenschlager I, Wolff P, Wolverton C, Bhalerao RP, Sandberg G, Ishikawa H, Evans M, Palme K (2003)
Gravity-regulated differential auxin transport from columella to lateral root cap cells. Proc Natl
Acad Sci U S A 100: 2987-2991
Paciorek T, Zazimalova E, Ruthardt N, Petrasek J, Stierhof YD, Kleine-Vehn J, Morris DA, Emans N,
Jurgens G, Geldner N, Friml J (2005) Auxin inhibits endocytosis and promotes its own efflux
from cells. Nature 435: 1251-1256
Pan J, Fujioka S, Peng J, Chen J, Li G, Chen R (2009) The E3 ubiquitin ligase SCFTIR1/AFB and
membrane sterols play key roles in auxin regulation of endocytosis, recycling, and plasma
membrane accumulation of the auxin efflux transporter PIN2 in Arabidopsis thaliana. Plant Cell
21: 568-580
Pike LJ (2006) Rafts defined: a report on the Keystone Symposium on Lipid Rafts and Cell Function. J
Lipid Res 47: 1597-1598
Rashotte AM, Brady SR, Reed RC, Ante SJ, Muday GK (2000) Basipetal auxin transport is required for
gravitropism in roots of Arabidopsis. Plant Physiol 122: 481-490
Robert HS, Friml J (2009) Auxin and other signals on the move in plants. Nat Chem Biol 5: 325-332
Chapter III
79
Roche Y, Gerbeau-Pissot P, Buhot B, Thomas D, Bonneau L, Gresti J, Mongrand S, Perrier-Cornet JM,
Simon-Plas F (2008) Depletion of phytosterols from the plant plasma membrane provides
evidence for disruption of lipid rafts. Faseb J 22: 3980-3991
Souter M, Topping J, Pullen M, Friml J, Palme K, Hackett R, Grierson D, Lindsey K (2002) hydra
Mutants of Arabidopsis are defective in sterol profiles and auxin and ethylene signaling. Plant
Cell 14: 1017-1031
Spassieva SD, Markham JE, Hille J (2002) The plant disease resistance gene Asc-1 prevents disruption of
sphingolipid metabolism during AAL-toxin-induced programmed cell death. Plant J 32: 561-572
Sperling P, Heinz E (2003) Plant sphingolipids: structural diversity, biosynthesis, first genes and
functions. Biochim Biophys Acta 1632: 1-15
Steinmann T, Geldner N, Grebe M, Mangold S, Jackson CL, Paris S, Galweiler L, Palme K, Jurgens G
(1999) Coordinated polar localization of auxin efflux carrier PIN1 by GNOM ARF GEF. Science
286: 316-318
Sun Y, Taniguchi R, Tanoue D, Yamaji T, Takematsu H, Mori K, Fujita T, Kawasaki T, Kozutsumi Y
(2000) Sli2 (Ypk1), a homologue of mammalian protein kinase SGK, is a downstream kinase in
the sphingolipid-mediated signaling pathway of yeast. Mol Cell Biol 20: 4411-4419
Swarup K, Benkova E, Swarup R, Casimiro I, Peret B, Yang Y, Parry G, Nielsen E, De Smet I, Vanneste S,
Levesque MP, Carrier D, James N, Calvo V, Ljung K, Kramer E, Roberts R, Graham N,
Marillonnet S, Patel K, Jones JD, Taylor CG, Schachtman DP, May S, Sandberg G, Benfey P,
Friml J, Kerr I, Beeckman T, Laplaze L, Bennett MJ (2008) The auxin influx carrier LAX3
promotes lateral root emergence. Nat Cell Biol 10: 946-954
Swarup R, Friml J, Marchant A, Ljung K, Sandberg G, Palme K, Bennett M (2001) Localization of the
auxin permease AUX1 suggests two functionally distinct hormone transport pathways operate in
the Arabidopsis root apex. Genes Dev 15: 2648-2653
Swarup R, Kargul J, Marchant A, Zadik D, Rahman A, Mills R, Yemm A, May S, Williams L, Millner P,
Tsurumi S, Moore I, Napier R, Kerr ID, Bennett MJ (2004) Structure-function analysis of the
presumptive Arabidopsis auxin permease AUX1. Plant Cell 16: 3069-3083
Swarup R, Kramer EM, Perry P, Knox K, Leyser HM, Haseloff J, Beemster GT, Bhalerao R, Bennett MJ
(2005) Root gravitropism requires lateral root cap and epidermal cells for transport and response
to a mobile auxin signal. Nat Cell Biol 7: 1057-1065
Tanaka H, Dhonukshe P, Brewer PB, Friml J (2006) Spatiotemporal asymmetric auxin distribution: a
means to coordinate plant development. Cell Mol Life Sci 63: 2738-2754
Teng C, Dong H, Shi L, Deng Y, Mu J, Zhang J, Yang X, Zuo J (2008) Serine palmitoyltransferase, a key
enzyme for de novo synthesis of sphingolipids, is essential for male gametophyte development
in Arabidopsis. Plant Physiol 146: 1322-1332
Chapter III
80
Titapiwatanakun B, Blakeslee JJ, Bandyopadhyay A, Yang H, Mravec J, Sauer M, Cheng Y, Adamec J,
Nagashima A, Geisler M, Sakai T, Friml J, Peer WA, Murphy AS (2009) ABCB19/PGP19
stabilises PIN1 in membrane microdomains in Arabidopsis. Plant J 57: 27-44
Ulmasov T, Murfett J, Hagen G, Guilfoyle TJ (1997) Aux/IAA proteins repress expression of reporter
genes containing natural and highly active synthetic auxin response elements. Plant Cell 9:
1963-1971
Utsuno K, Shikanai T, Yamada Y, Hashimoto T (1998) Agr, an Agravitropic locus of Arabidopsis thaliana,
encodes a novel membrane-protein family member. Plant Cell Physiol 39: 1111-1118
Vanneste S, Friml J (2009) Auxin: a trigger for change in plant development. Cell 136: 1005-1016
Wang W, Yang X, Tangchaiburana S, Ndeh R, Markham JE, Tsegaye Y, Dunn TM, Wang GL, Bellizzi M,
Parsons JF, Morrissey D, Bravo JE, Lynch DV, Xiao S (2008) An inositolphosphorylceramide
synthase is involved in regulation of plant programmed cell death associated with defense in
Arabidopsis. Plant Cell 20: 3163-3179
Willemsen V, Friml J, Grebe M, van den Toorn A, Palme K, Scheres B (2003) Cell polarity and PIN
protein positioning in Arabidopsis require STEROL METHYLTRANSFERASE1 function. Plant
Cell 15: 612-625
Wisniewska J, Xu J, Seifertova D, Brewer PB, Ruzicka K, Blilou I, Rouquie D, Benkova E, Scheres B,
Friml J (2006) Polar PIN localization directs auxin flow in plants. Science 312: 883
Yoshida S, Uemura M (1986) Lipid Composition of Plasma Membranes and Tonoplasts Isolated from
Etiolated Seedlings of Mung Bean (Vigna radiata L.). Plant Physiol 82: 807-812
Zappel NF, Panstruga R (2008) Heterogeneity and lateral compartmentalization of plant plasma
membranes. Curr Opin Plant Biol 11: 632-640
Zheng H, Rowland O, Kunst L (2005) Disruptions of the Arabidopsis Enoyl-CoA reductase gene reveal
an essential role for very-long-chain fatty acid synthesis in cell expansion during plant
morphogenesis. Plant Cell 17: 1467-1481
- Chapter IV -
Analysis of AtMob1 function in plant development
Chapter IV
82
Abstract
The MOB1 (MPS one binder 1) family includes a group of proteins conserved
throughout eukaryotes. The family founding member was originally identified in yeast
as a component of a signalling pathway regulating exit from mitosis and cytokinesis. In
multicellular organisms, MOB1 proteins play a role in fundamental processes like cell
proliferation and apoptosis, thus controlling appropriate cell number and organ size. In
this study, we investigate the role of two Arabidopsis MOB1-like genes, namely
MOB1A and MOB1B, in plant development. An AtMOB1B loss-of-function mutant did
not show any defect. In contrast, suppression of AtMOB1A expression affected growth
and reproduction and led to defects in ovule development as well as in the tissue
pattern of the root apex. The nuclear localisation of MOB1 proteins supports the
hypothesis that they might have maintained a function in cell division control,
similarly to their yeast and animal orthologs.
Introduction
In plants, embryogenesis generates only a basic body organisation with an apical-basal
pattern rather than the complete organism, as it occurs in animals (reviewed
by(Jurgens, 2001; Willemsen and Scheres, 2004; Jenik et al., 2007). Most of the plant
body is formed post-embryonically with the continuous generation of new tissues and
structures throughout plant life. The process of organogenesis occurs in a reiterative
form and depends on the activity of meristematic zones located at the tip of plant
organs (reviewed by(Baurle and Laux, 2003; Jurgens, 2003). Meristems are constituted
by cells that complete several rounds of cell division before undergoing expansion and
differentiation. Thus, a tight regulation of cell division is crucial to sustain organ
outgrowth and ultimately enables the plant to complete its developmental program.
During mitosis, complete copies of the genome have to successfully segregate between
the two daughter nuclei. As control mechanism, eukaryotic cells evolved signalling
components that coordinate exit from mitosis with cytokinesis. Knowledge about this
mechanism mostly derives from extensive studies in the fission and budding yeasts,
Schizosaccharomyces pombe and Saccharomyces cerevisiae, respectively. S. pombe
Chapter IV
83
cells divide by constriction of an actomyosin ring and concomitant assembly of a
division septum, corresponding to a new cell wall (Gould and Simanis, 1997). S.
cerevisiae divides by forming a bud (Chant and Pringle, 1995). The onsets of septation
in S. pombe and budding in S. cerevisiae are signalled through the septation initiation
network (SIN) and the mitotic exit network (MEN) signalling pathways, respectively
(reviewed by(Bardin and Amon, 2001). SIN and MEN are similar signalling networks
using orthologous proteins that control events at the end of mitosis. Both networks
consist of a GTPase-activated kinase cascade. In the case of MEN, the activated form of
the RAS-like GTPase Tem1 is thought to propagate a signal to the protein kinase
Cdc15, which in turn activates the protein kinase Dbf2. It is known that Dbf2 kinase
activity requires the Dbf2-associated factor Mob1 (Mah et al., 2001). The Mob1-Dbf2
interaction leads to release from the nucleolus and subsequent activation of Cdc14
phosphatase during anaphase (Stegmeier and Amon, 2004; Mohl et al., 2009). The
release of Cdc14 from its inhibitor complex (Shou et al., 1999) promotes the
inactivation of the mitotic Cdk1-cyclin B complex finally driving exit from mitosis
(Visintin et al., 1998). Besides its primary role as promoter of mitotic exit, the MEN has
been shown to control also cytokinesis (Lee et al., 2001; Lippincott et al., 2001; Luca et
al., 2001). The SIN signalling cascade is organised similarly to the MEN but its main
role is to control cytokinesis by initiating contraction of the actin ring and synthesis of
the septum (reviewed by(Krapp and Simanis, 2008). In S. pombe, the ortholog of S.
cerevisiae Dbf2 kinase is represented by Sid2, whose activity similarly requires the
interaction with Mob1 (Hou et al., 2000). Yeast Mob1 proteins do not function solely as
activators of Dbf2/Sid2, but are also required for Dbf2/Sid2 localisation to activation
sites (Frenz et al., 2000; Lee et al., 2001; Hou et al., 2004). Indeed, in agreement with
their functions in mitosis exit and cytokinesis, Dbf2/Sid2-Mob1 complexes localise to
the spindle pole body (SPB) in anaphase and move to the division site in late mitosis
(Yoshida and Toh-e, 2001). Different conditional mutations of yeast Mob1 cause a late
nuclear division arrest at restrictive temperature and result in a quantal increase in
ploidy at the permissive temperature (Luca and Winey, 1998).
Chapter IV
84
Several components of the MEN and SIN pathways are conserved among eukaryotes
and are similarly involved in the regulation of cell division in multicellular organisms
(Mailand et al., 2002; Bothos et al., 2005; Hergovich et al., 2006; Bedhomme et al.,
2008). Studies in Drosophila have related Mob proteins also to a different signalling
pathway that plays a crucial role in tissue growth and cell number control. Two protein
kinases Hippo (Hpo) and Warts (Wts)/large tumor suppressor (Lats), Hpo-scaffold
protein Salvador (Sav) and Mats (Mob as tumor suppressor, dMob1) are the key
components of this pathway (Justice et al., 1995; Tapon et al., 2002; Harvey et al., 2003;
Lai et al., 2005). Loss of any of these factors results in increased cell proliferation and
decreased cell death, indicating that Sav, Hpo, Lats and Mats all function as tumor
suppressors. The Dbf2-related Lats is phosphorylated by Hpo and needs to bind to its
co-activator Mats to properly coordinate cell death and proliferation (reviewed by(Pan,
2007). The components of the Hippo-pathway are conserved from yeast to flies and
humans, suggesting that this signalling cascade plays a fundamental role in cellular
regulation.
Cell division is more complex in plants than in animals due to the presence of a rigid
external cell wall. In contrast to yeast and animal cells, plant cells undergoing cell
division display two unique cytoskeletal structures, namely the pre-prophase band
(PPB) and the phragmoplast, which are necessary to assure adequate positioning and
assembly of a new cell wall between the separating sister nuclei (Verma, 2001). On the
other hand, plants do not possess SPBs and centrosomes. Despite these differences,
several components of the MEN/SIN pathways are conserved in plants (Bedhomme et
al., 2008). In particular, several genes encoding putative proteins homologous to yeast
Mob1 have been identified in different plant species (Vitulo et al., 2007). In Medicago
sativa L., MOB1-like genes were shown to be constitutively expressed with a maximum
in proliferating tissues (Citterio et al., 2006). The Arabidopsis genome contains four
different MOB1-like genes that can be divided into two subgroups according to their
homology (Citterio et al., 2006; Vitulo et al., 2007). We have recently started the
characterisation of these genes investigating their role during gametophytic
development (Galla et al., 2009). Here, we report the isolation of different T-DNA
Chapter IV
85
insertion alleles for two MOB1-like genes (AtMOB1A and AtMOB1B) and show that
proper expression of AtMOB1A is required for plant growth and reproduction and for
the maintenance of the root meristem pattern.
Materials and methods
Alignment of MOB1A and MOB1B protein sequences
Amino acid sequences were aligned by Clustal W method (Gonnet protein weight
matrix, gap penalty = 10, gap length penalty = 0.2, delay divergent seqs = 30%) using
the MegAlign software (DNASTAR).
Plasmid construction and plant transformation
The generation of MOB1A RNAi and p35S::GFP-MOB1A constructs have been
described by Galla et al. (2009). For MOB1A RNAi construct, a unique 158 bp cDNA
fragment was amplified using specific primers designed in the 3’-UTR of MOB1A gene
(At5g45550) and the PCR product was cloned into a pENTRTM/D-TOPO_vector
(Invitrogen,). This was then used for LR recombination using the RNAi Gateway
destination vector pK7GWIWG2(II) (Karimi et al., 2002) to produce the MOB1A-RNAi
vector. For p35S::GFP-MOB1A construct the coding sequence of MOB1A was
amplified from Arabidopsis leaf cDNA. The PCR product was cloned into pENTRTM/DTOPO_vector and subsequently transferred to the destination vector pK7FWG2
(Karimi et al., 2002) to create a N-terminal gfp fusion. Binary vectors were introduced
into Agrobacterium tumefaciens EHA105 by electroporation. Arabidopsis plants
(Col-0) were transformed by a modified version of the floral dip method (Clough and
Bent, 1998), in which the Agrobacterium culture was applied directly to flower buds
using a pipette.
Plant material and growth conditions
Arabidopsis thaliana (L.) Heynh. (Col-0) was used as wild type. The SALK_076775,
SALK_062070 and GK719G04 lines were obtained from the Nottingham Arabidopsis
Stock Centre (NASC) (Scholl et al., 2000). MOB1A RNAi and p35S::GFP-MOB1A lines
Chapter IV
86
have been described by Galla et al. (2009). The J2341 enhancer trap line belongs to the
Hasselhoff collection and was provided by the NASC. The pWOX5::GFP line was
described by Ditengou et al. (2008). Seeds were surface sterilised for 15 min. with a
solution of 5% w/v calcium hypochlorite and 0.02% Triton X-100. After 3 washes in
sterile water, they were left to dry under sterile conditions. Seeds were sown on plates
containing 1% w/v sucrose, half-strength MS salts (Duchefa) and 12 g/l agar-agar
(Roth) (pH 5.8). After two days of vernalisation at 4°C in darkness, plates were
transferred to a growth chamber (16h light/8h darkness, 22°C) for seed germination
and were maintained in a vertical position.
Characterisation of homozygous T-DNA insertion mutants
Individual T3 plants of SALK_076775, SALK_062070 and GK719G04 were screened by
PCR. The T-DNA insertion alleles were verified using a primer annealing to a sequence
of the T-DNA left border (LBb1 for the SALK lines and GK8409 for GK719G04) and a
gene specific primer (P2, P7 and P4 for SALK_076775, SALK_062070 and GK719G04,
respectively). The wild-type alleles were amplified using a pair of gene specific primers
(P1 and P2 for SALK_076775; P7 and P8 for SALK_062070; P3 and P4 for GK719G04).
The primer sequences and their annealing positions are indicated in Table 1 and Fig. 1,
respectively.
RT-PCR analysis
Total RNA was isolated from one week old seedlings using the RNeasy plant kit
(Qiagen) according to the manufacturer’s protocol. Total RNA (1 μg) was first treated
with DNase (Qiagen) and first-strand cDNA was subsequently synthesised using
RevertAid™ M-MuLV Reverse Transcriptase (Fermentas) and oligo(dT) primer,
according to manufacturer’s instructions. One and a half microliter of first-strand
cDNA was used as template for PCR amplification in a 25 μl reaction employing a
home made Taq DNA polymerase. The primer pairs and the relative annealing
temperatures used for the amplification of MOB1A, MOB1B and ACTIN 2 are reported
in Table 1. Reactions were performed at 95° for 30 s, annealing temperature for 30 s
Chapter IV
87
and 72° for 1 min (30 cycles for MOB1A and MOB1B and 25 cycles for ACTIN 2). The
ACTIN 2 gene (At5g09810) was used as an internal control.
Plant growth observations and root length measurements
After 1 week of growth in plates, seedlings were transferred to soil in pots. Images of
rosettes, rosette leaves and siliques were taken with a digital camera. For the
examination of seeds contained in siliques, the latter were dissected on a slide under a
Zeiss Stemi SV11 Apo stereomicroscope (Carl Zeiss MicroImaging,) and images were
acquired with an AxioCam MRc camera (Zeiss). Length measurements of whole root,
meristem and elongation zone were performed on seedlings five days after germination
induction. For whole root length measurements, plates containing seedlings were
scanned with a CanonScan 9950F scanner. For length measurements of meristem and
elongation zone, seedlings were mounted on slides in chloral hydrate:glycerol:water
(8:3:1, w/vol/vol). Images were subsequently acquired with an Axiovert 200M MAT
microscope (Zeiss) equipped with DIC optics and an AxioCam ICc 1 camera (Zeiss).
Root sizes were determined with the Image J software (http://rsbweb.nih.gov/ij/).
Immunocytochemistry
Whole-mount immunodetections with four days old seedlings were performed as
described elsewhere (Chapter II, p. 38). The concentrations of primary antibodies were
1:400 for rabbit anti-PIN4 (Friml et al., 2002), 1:300 for rabbit anti-MOB1 (Citterio et
al., 2005) and 1:200 for mouse anti-α-tubulin (Molecular Probes). Different fluorescent
Alexa-conjugated secondary antibodies (Invitrogen) were employed at 1:600 (anti
rabbit-A488 for MOB1, anti rabbit-A555 for PIN4 and anti mouse-A555 for α-tubulin).
Samples were mounted in Prolong Gold antifade reagent containing DAPI (Molecular
probes). For immunolocalisations of wild-type ovules, flowers were collected and
dissected under a Zeiss Stemi SV11 Apo stereomicroscope (Carl Zeiss MicroImaging,
Germany). Carpels were fixed with 4% paraformaldehyde/MTSB (pH 7.0) for 1 h and
washed three times with ddH2O. Tissue clearing was obtained with two washes in
methanol for 20 min and progressive substitution with distilled water. Permeabilisation
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was achieved by 30 min incubation in 0.15% Driselase (Sigma), 0.15% Macerozyme
(Sigma) in 10 mM MES (pH 5.3) at 37°C, followed by one wash in MTSB and successive
treatment with 10% DMSO, 3% Nonidet P40 (Fluka). After two washes in MTSB,
blocking was performed with 3% BSA (Roth) in MTSB for one hour at RT. Rabbit antiMOB primary antibody (1:200) in 3% BSA in MTSB was applied for 1.5 hours at RT,
followed by two washes in MTSB. Goat anti-rabbit A555-conjugated (1:600) secondary
antibody (Invitrogen) was applied for 1.5 h at RT, followed by three washes in MTSB.
Carpels were mounted in Prolong Gold antifade reagent containing DAPI (Molecular
Probes).
Lugol and propidium iodide straining
For Lugol staining, five days old seedlings were dipped in Lugol’s staining reagent
(Roth) for 10 min, rinsed twice in water and mounted on slides in chloral
hydrate:glycerol:water (8:3:1, w/vol/vol). Images were acquired with an Axiovert 200M
MAT microscope (Zeiss) equipped with DIC optics and an AxioCam ICc 1 camera
(Zeiss). For propidium iodide (PI) staining, five days old seedlings were incubated for
10 min in a 10 μg/ml PI solution and mounted on slides in water.
Confocal microscopy
Images were acquired using a Zeiss LSM 510 NLO confocal scanning microscope.
Excitation wavelengths were 488 nm (argon laser) for GFP, Alexa488-conjugated
antibodies
and
propidium
iodide
and
543 nm
(Helium-Neon-Laser)
for
Alexa555-conjugated antibodies. Emission was detected at 500-550 nm for GFP and
Alexa488-conjugated antibodies, above 560 nm for propidium iodide and above 575 nm
for Alexa555-conjugated antibodies. DAPI was imaged using a 2-Photon module with
excitation at 730 nm and emission at 435-485 nm. All multi-labelling signals were
detected in multitracking mode to avoid fluorescence crosstalk. Images were analyzed
with the LSM image browser (Carl Zeiss MicroImaging) and Adobe Photoshop CS2.
Chapter IV
89
Results (Hruz T, 2008; Galla, 2009)
Sequence comparison and expression analysis of two MOB1-like genes in Arabidopsis
Blast analysis revealed that, among the four MOB1-like genes present in the
Arabidopsis genome, At5g45550 and At4g19045 encode predicted proteins with the
highest similarity to S. cerevisiae Mob1 (E-values: 9e-51 and 1e-49, respectively). The
two protein sequences contain both 215 amino acids and share 93% of identity (Fig.
1A). We renamed the two genes MOB1A (At5g45550) and MOB1B (At4g19045).
Figure 1. Sequence, expression pattern, gene structure and mutant alleles of Arabidopsis Mob1A and
Mob1B. (A) Alignment of AtMob1A and AtMob1B protein sequences. Identical residues are shaded
black and correspond to the 93% of the whole sequences. (B) Microarray data for expression of
AtMob1A in different organs of Arabidopsis. Data were obtained from the public microarray database
AtGenExpress. AtMob1A expression was detected in every organ at comparable levels. (C) Schematic
representation of AtMob1A and AtMob1B intron-exon structure and T-DNA insertion sites for different
SALK and GABI-Kat lines. Black and grey boxes indicate exons and UTR-regions, respectively. Arrows
indicate primers used for genotyping and RT-PCR analysis of T-DNA insertion lines. The primer pairs
employed for PCR-amplification of each allele are described in “Material and methods” and the relative
sequences are reported in Table 1. (D) Semiquantitative RT-PCR analysis of Mob1A and Mob1B T-DNA
Chapter IV
90
lines. The three panels show the amplification of Mob1A coding sequence (top), Mob1B coding sequence
(middle) and a fragment of the ACTIN 2 gene (At5g09810; internal control) (bottom) in wild-type Col-0
(wt), mob1A-2 (SALK_076775), mob1A-1 (GK719G04) and mob1B-1 (SALK_062070). The primer pairs
used for the analysis are reported and their relative positions are indicated in (C). RT-PCR cycle numbers
are indicated.
We have recently shown the presence of MOB1A gene transcript in roots, leaves,
flowers and siliques by real time PCR analysis (Galla et al., 2009). Consistently,
microarray
data
from
the
public
AtGenExpress
database
(http://www.weigelworld.org/resources/microarray/AtGenExpress/; (Schmid et al.,
2005) displayed ubiquitous expression of MOB1A in Arabidopsis tissues with a
maximum in floral organs (Fig. 1B). Expression data were also retrieved from the
publicly
available
microarray
database
GENEVESTIGATOR
(https://www.genevestigator.com/gv/index.jsp; Hruz et al., 2008). The largest changes
in expression of MOB1A, as observed from the analysis of over 250 treatments, were an
approximately two-fold increase in response to transformation of rosette leaves with
cabbage leaf curl virus DNA and a two-fold decrease in response to application of
brassinolide and boric acid to cell cultures. Locus At4g19045 has recently been
annotated as predicted gene by The Arabidopsis Information Resource (TAIR;
http://www.arabidopsis.org/) and microarray data have not yet been available for it.
Isolation and characterisation of T-DNA insertion lines and RNAi lines for AtMOB1
genes
In order to assess the effects of a reduced expression of MOB1 genes in plants, we
employed two different strategies. First, we examined putative T-DNA disruption
mutants of the two genes. Two independent GABI-Kat lines (GK719G04 and
GK295E12; Rosso et al., 2003) and one SALK T-DNA line (SALK_076775; Alonso et al.,
2003) were available for MOB1A. We confirmed the presence of a T-DNA insertion in
GK719G04 and SALK_076775, respectively in the first intron and in the promoter
region of the gene (Fig. 1C). These two T-DNA disruption alleles were designated
Chapter IV
91
mob1A-1 and mob1A-2, respectively. RT-PCR analysis with primers flanking MOB1A
coding sequence (P5 and P6) revealed a level of transcript similar to wild-type in
homozygous mob1A-2 plants (Fig. 1D). In contrast, we could not detect MOB1A
mRNA (primers P5 and P6) in homozygous mob1A-1 plants, suggesting that mob1A-1
represents a null allele (Fig. 1D). For MOB1B only the SALK_062070 line was available
and could be confirmed to contain a T-DNA insertion in the fourth exon (Fig. 1C). This
T-DNA disruption allele was named mob1B-1. RT-PCR analysis of mob1B-1
homozygous plants with primers flanking the ORF (P9 and P10) displayed the absence
of MOB1B transcript, demonstrating mob1B-1 as a null allele (Fig. 1D). Moreover, the
expression of MOB1B in mob1A-1 and that of MOB1A in mob1B-1 was not increased
in comparison to the wild-type levels (Fig. 1D).
Table 1. Nucleotide sequences and employed annealing temperatures of primers used for genotyping and
RT-PCR analysis of Mob1A and Mob1B T-DNA lines.
Primer
Sequence
Annealing
T (°C)
Primers for T-DNA insertion lines genotyping
LBb1
GCGTGGACCGCTTGCTGCAACT
GK_8409
ATATTGACCATCATACTCATTGC
P1
TCTGATATTAACCGCAAACGC
P2
AATCAACCATGAACCTGAATCC
P3
GGATTCGTGTGGCTTTCTTC
P4
GGATTCGTGTGGCTTTCTTC
P7
TCCTGCAAAACAAAACCAGAC
P8
CTAAGAAGAGCGCACCATCAG
57
Primers for RT-PCR analysis
P5
TGAGTCTCTTTGGGTTAGG
P6
TGAGTCTCTTTGGGTTAGG
P9
CGTGTCTCACTCCGATAAAGC
P10
TGGACAGAGGATTTGGGTTT
Act2_for
TGTTCACCACTACCGCAGAA
Act2_rev
GGTGCAACCACCTTGATCTT
50
57
57
Chapter IV
92
In a second approach, we studied MOB1A loss-of-function effects in three independent
RNAi lines, whose generation has been recently described (Galla et al., 2009). The
reduction of MOB1A transcript level in these lines was between 50% and 70% in
comparison to the wild-type level.
Reduced MOB1A expression causes delayed plant growth and reproductive defects
As an initial step to analyse the effects caused by the reduction of MOB1 gene
expression, the phenotypes of mob1A-1 and mob1B-1 plants grown in soil were
examined. mob1B-1 plants did not display any major defect in plant growth and
development and reproduced normally to the next generation (data not shown). In
contrast, mob1A-1 exhibited several growth and reproductive defects.
Figure 2. mob1A-1 displays defects in plant growth and reproduction. (A) Rosette phenotype of three
weeks-old wild-type Col-0 (wt) and mob1A-1 plants. Note the reduced size of mob1A-1 mutant. (B)
Arrangement of all leaves from three weeks-old wild-type (wt) and mob1A-1 plants reveals a reduced
Chapter IV
93
number of leaves for mob1A-1. (C-D) Flowers of wild-type (C) and mob1A-1 (D). One sepal and one
petal were removed to allow the inspection of carpels and stamens. mob1A-1 does not display major
floral organ defects. (E) Siliques of wild-type (wt) and mob1A-1 plants 10 days after flower opening.
mob1A-1 siliques display a clear reduction in size. (F-G) Silique content from wild-type (F) and
mob1A-1 (G) plants. Shown are siliques at 8 to 10 days after flowering. mob1A-1 siliques contain several
aborted ovules (arrowheads). (H) Average percentage of aborted ovules from siliques of wild-type and
mob1A-1 plants. Average ± SD are reported; n=3 (10 siliques per plant; >370 total seeds were examined).
Scale bars are 500 μm (C-D and F-G).
mob1A-1 rosettes contained a reduced number of leaves when compared to wild-type
plants of the same age (Fig. 2A-B) and were characterised by delayed bolting. Although
mob1A-1 carpels and stamens did not display any major morphological alteration
(Fig. 2C-D), siliques collected 8 to 10 days after flowering presented a strongly reduced
size (Fig. 2E). An inspection of siliques content revealed a dramatically high proportion
of aborted ovules for mob1A-1 plants (Fig. 2F-H).
In agreement with the phenotype of mob1A-1, defects in correct ovule development
were shown for all three RNAi lines of MOB1A (Galla et al., 2009). Moreover, one of
the lines exhibited also a reduced growth of the vegetative organs.
Taken together, these results indicate the importance of MOB1A in plant growth and
reproductive development.
Reduced MOB1A expression causes defects in root growth and root meristem
patterning
Given the ubiquitous expression of MOB1A in Arabidopsis tissues (Galla et al., 2009),
we next examined whether its reduced expression affected also root development.
Five-days-old mob1A-1 plants exhibited a shorter root length in comparison to wildtype seedlings (Fig. 3A). Furthermore, length measurements of mob1A-1 roots revealed
a significant reduction in the meristem size, while the size of the elongation zone was
not affected (Fig. 3A). This observation suggests that the reduced root length
phenotype might be caused by defects in cell proliferation in the meristem region. In
contrast, mob1B-1 plants did not show any change in root length (Fig. 3A).
Chapter IV
94
Figure 3. Reduced levels of Mob1A expression affect root length, root meristem size and root tip cellular
pattern. (A) Analysis of whole root length (left panel), meristem size (middle panel) and elongation zone
size (right panel) for five days old wild-type (wt) and mob1A-1 seedlings. Averages ± SE are reported;
n=3 and at least fifteen roots were used for each experiment. Whole root measurements were performed
also on mob1B-1 seedlings and did not reveal significant differences with wt. mob1A-1 roots exhibit a
significant reduction in root length and meristem size (Student’s t-test, P<0,001). (B-E) Propidium iodide
staining of wild-type (B), Mob1A RNAi (C and E) and mob1A-1 (D) roots. The cellular patterns of
Mob1A RNAi and mob1A-1 root tips appear altered in comparison to the situation in wt, which shows
aligned cell files. In some cases (E) the pattern defects were particularly severe and the whole root tip
Chapter IV
95
morphology was affected. (F-G) Lugol staining of wild-type (F) and Mob1A RNAi (G) root tips. Starch
granules accumulated in the columella region of Mob1A RNAi roots differently from the clear
distribution displayed by wt roots. (H-I) Immunolocalised PIN4 (red) and DAPI staining (blue) in
wild-type (H) and Mob1A RNAi (I) root tips. Cells labelled with PIN4 show a disordered pattern in
Mob1A RNAi roots in comparison to wild-type roots. (J-K) Root tip GFP distribution in pWOX5::GFP
reporter line (J) and in a F2 population from a pWOX5::GFP x Mob1A RNAi crossing (K). Seedlings were
incubated for 10 min in 4 μm FM4-64 prior to microscopy observation to visualise the cell plasma
membrane. Note the expanded GFP expression in the sample from the F2 crossing population. (L-M)
Root tip GFP distribution in J2341 columella initials marker line (L) and in a F2 population from a J2341
x Mob1A RNAi crossing (M). Plasma membranes were stained with FM4-64 as described above. Cells
expressing GFP in the F2 crossing population failed to display the same alignment as in J2341 roots. Scale
bars are 20 μm (A-M).
Root tip microscopic inspections of mob1A-1 seedlings and T2 plants from three
independent MOB1A RNAi lines revealed severe defects in tissue patterning around
the quiescent centre (QC) and the stem cell niche. Wild-type root tips stained with
propidium iodide exhibited an ordered cellular organisation in the great majority of the
examined seedlings (90%). Typically, the QC was flanked by cortex and endodermis
stem cells. Apically to the QC a file of columella stem cells (CSCs) and different
columella cell-layers could be observed (Fig. 3B). In contrast, the root tips of mob1A-1
and MOB1A RNAi showed a disordered cellular pattern with cells not aligned in files
(Fig. 3C-D). In some cases the whole morphology of the root tip was perturbed
(Fig. 3E). This phenotype was not common to all the roots analysed but had a different
penetrance, between 20 and 33.3% in MOB1A RNAi lines and 43.3% in mob1A-1, as
illustrated in Table 2. To further characterise the observed disorder in the cellular
organisation of the root tips, we employed several tissue markers that label this region.
First, the distribution of starch granules was examined by means of Lugol staining in
columella cells. While wild-type roots displayed an ordered distribution reflecting
columella organisation in different cell files, in MOB1A RNAi roots starch granules
accumulated in the columella region without any order (Fig. 3F-G). A suitable marker
labelling the plasma membrane of the cells around the QC is the auxin efflux carrier
Chapter IV
96
Table 2. Frequency of root tips with a disordered cellular pattern in wild-type, Mob1A RNAi lines and
mob1-A mutant line.
Mob1A RNAi lines
wt
2F_1
4G_5
6E_7
mob1A-1
Root tips with
disordered cell. pattern
3
8
6
10
13
Observed root tips
30
30
30
30
30
Phenotype penetrance
10%
26.7%
20%
33.3%
43.3%
PIN4 (Friml et al., 2002). Indeed, immunolocalisation analysis in MOB1A RNAi plants
showed a disordered pattern of the cells expressing PIN4 around the stem cell niche
(Fig. 3H-I). Next, MOB1A RNAi lines were crossed with pWOX5::GFP (Ditengou et al.,
2008) and the enhancer trap line J2341 (described by(Sabatini et al., 2003), which show
GFP expression in QC cells and columella initials, respectively. Analysis of a F2
segregating population revealed that in root tips displaying a disordered cellular pattern
the GFP expression domains were expanded or reflected the failed alignment of cells in
files (Fig. 3J-M).
Overall, our data demonstrate a role for MOB1A in root growth and patterning.
MOB1 proteins display a preferential nuclear localisation.
To get an initial clue about the cellular function of MOB1 proteins in plants, we
addressed the question about their sub-cellular localisation. To this end, two different
approaches were employed, consisting in the analysis of five independent 35S::GFPMOB1A lines and in the immunolocalisation of MOB1 proteins in wild-type plants.
GFP-MOB1A fluorescent fusion displayed a nuclear and cytoplasmic localisation in all
examined cell types (Fig. 4) for all the five independent lines analysed. Whole-mount
immunocytochemistry experiments were performed using an antibody previously
described (Citterio et al., 2005) and recognising both MOB1A and MOB1B proteins.
Chapter IV
A
B
C
97
E
D
Figure 4. Constitutively expressed Mob1A-GFP localises to nucleus and cytoplasm. (A-E) Subcellular
localisation of Mob1A-GFP in different tissues of stably transformed p35S::Mob1A-GFP Arabidopsis
plants. (A-B) Root cells. (C) Root hair. (D) Stomata cells. (E) Cotyledon epidermal cells. All cell types
display a nuclear and cytoplasmic localisation of Mob1A-GFP. Scale bars are 5 μm (B) and 10 μm (A,
C-E).
All examined organs, comprising roots, shoot apical meristems, cotyledons, leaf
primordia and ovules, were positively labelled with the antibody (data not shown).
Given the observed effects of the reduced expression of MOB1A on roots and ovules,
we focused our attention on the sub-cellular localisation of MOB1 proteins in these
tissues. Within ovules, immunolocalised MOB1 proteins exhibited a predominantly
nuclear targeting and a weak cytoplasmic signal (Fig. 5). Interestingly, MOB1 nuclear
localisation in the megaspore mother cell was not detected at meiosis during nuclear
division (Fig. 5B), indicating a cell cycle regulation of MOB1 distribution. In root
meristematic cells, MOB1 proteins displayed a similar localisation pattern with a
preferential
nuclear
targeting
(Fig. 6).
Co-localisation
experiments
with
immunodetected MOB1 and α-tubulin and DAPI staining allowed us to follow protein
localisation during the different phases of cell division. In interphase cells and during
DNA condensation in prophase, MOB1 was clearly targeted to the nucleus (Fig. 6A-B).
When the chromosomes aligned in the centre of the cell, MOB1 followed the nuclear
envelope breakdown and exhibited a diffuse cytosolic localisation (Fig. 6C).
Chapter IV
98
Figure 5. Mob1 proteins exhibit a cell-cycle-dependent localisation in ovules during megasporogenesis.
(A-D) DIC images of Arabidopsis ovules at different stages of megasporogenesis and correspondent Mob1
immunolocalisation (‘), DAPI staining (‘’) and merged Mob1/DAPI images (‘’’). Arrowheads indicate the
positions of the megaspore mother cell (A’ and B’) and of the first two megaspores (C’ and D’). Mob1
proteins display a clear nuclear localisation in the megaspore mother cell (A-A’’’) but are not detected
during DNA condensation at meiosis (B-B’’’). Upon completion of the first meiotic division, Mob1
proteins progressively reappear around the two daughter nuclei (C-D). Scale bars are 5 μm (A-D).
Finally, when the two chromosome sets were split apart, MOB1 signal appeared around
them, increasing its intensity with the progression of cytokinesis (Fig. 6D-E).
In conclusion, the distribution of GFP-MOB1A fusion and the immunodetection of
MOB1 indicate a preferential nuclear localisation of MOB1 proteins and suggest their
possible involvement in cell division, as demonstrated for their orthologs in other
eukaryotes.
Chapter IV
99
Figure 6. Mob1 proteins exhibit a cell-cycle-dependent localisation in meristematic root cells. (A-E)
Images of Arabidopsis root meristematic cells displaying immunolocalised Mob1 proteins, DAPI staining
(‘), immunolocalised α-tubulin (‘’) and merged Mob1/DAPI/α-tubulin fluorescent signals (‘’’). The
different phases of mitosis are indicated, as assumed by DAPI staining and microtubular arrays. Mob1
proteins localise to the nucleus in interphase and prophase (A-B) and exhibit a diffuse cytoplasmic signal
during chromosome alignment at metaphase (C). At anaphase, Mob1 reappears around the separating
chromosome sets (D) and subsequently labels the two daughter nuclei (E). Scale bars are 5 μm (A-E).
Discussion
Developmental patterning and morphogenesis of multicellular organisms are
determined by coordinated cell proliferation, cell differentiation and programmed cell
death. MOB1 proteins are conserved among eukaryotes and are essential components
of pathways that control fundamental cellular processes such as mitotic exit,
cytokinesis and apoptosis (reviewed by(Vitulo et al., 2007). In this study, we
investigated the role played by two MOB1-like genes during plant development in
Chapter IV
100
Arabidopsis. MOB1A (At5g45550) and MOB1B (At4g19045) are the closest Arabidopsis
orthologs to S. cerevisiae Mob1 and their predicted protein sequences share a very high
identity level (93%). Yeast Mob1 is a key player of MEN and SIN, two signalling
pathways that coordinate exit from mitosis with cytokinesis in S. cerevisiae and S.
pombe, respectively. Recently, Bedhomme et al. (2008) have shown that, for several
MEN/SIN components, the Arabidopsis genome comprises two paralogues for each of
the yeast orthologous genes and have suggested a certain level of functional
redundancy.
Expression data from publicly available databases and recently published real time PCR
analysis (Galla et al., 2009) demonstrated that MOB1A is ubiquitously expressed in all
Arabidopsis organs. Similarly, immunocytochemistry experiments revealed the
presence of MOB1 proteins in all tissues analysed. Our results are in agreement with a
previous study showing that transcripts and proteins of two MOB1-like genes in
Medicago sativa are present in roots, stems, leaves, flowers and pods and they are
mostly produced in actively proliferating tissues (Citterio et al., 2006). Interestingly,
the expression of some SIN components conserved in plants is limited to a restricted
subset of cells related to differentiation events, although present in several organs
(Bedhomme et al., 2009). It was suggested that these proteins have evolved in plants to
perform a function different from the SIN pathway. It will be thus interesting to
evaluate the expression of MOB1A and MOB1B genes with a detailed spatial-temporal
microscopic analysis in order to study their potential expression overlap and to help
clarifying their function in plants.
The high identity level of MOB1A and MOB1B amino acid sequences suggests a
common function for the two proteins. In contrast to the major growth and
developmental defects of a MOB1A knock-out mutant (mob1A-1), a null allele of
MOB1B (mob1B-1) did not reveal any phenotypical effect. Additionally,
semiquantitative RT-PCR analysis of MOB1A and MOB1B expression showed that
loss-of-function in one of the two genes did not induce an up-regulation of the other
homolog. These results might indicate that the potential common biological function of
MOB1A and MOB1B is mainly accomplished by the first or that MOB1B expression is
Chapter IV
101
controlled by a not yet identified external signal or has different spatial-temporal
expression kinetics. Therefore, the ongoing isolation of mob1A-1,mob1B-1 double
mutant will provide an insight into the functional redundancy between the two genes.
The phenotype of mob1A-1 included defects in the number of rosette leaves, bolting
time, ovule development, root growth and root tip cellular organisation. All these traits
could be potentially linked to the hypothesised function of Arabidopsis MOB1A in cell
division regulation and cell proliferation. The defects in ovule development observed in
mob1A-1 have been described also for MOB1A RNAi lines (Galla et al., 2009). In
particular, post-transcriptional silencing of MOB1A affected the normal progression of
both female meiosis and megagametogenesis, leading to multinucleated megaspores and
embryo sacs carrying cellularisation defects. Thus, our data support the role proposed
by Galla et al. (2009) for MOB1A during reproductive processes.
When compared to wild-type, mob1A-1 seedlings exhibited shorter roots and a
reduced size of the root meristem. Given the possible role of MOB1A in the regulation
of cell division, a reduced cell proliferation in the meristem region of mob1A-1 might
explain its shorter size and account for the overall reduced root length. Moreover, the
roots of mob1A-1 and MOB1A RNAi lines displayed a disordered cellular pattern in the
root tip area around the stem cell niche. Cells normally aligned in adjacent cell files in
wild-type occupied anomalous positions exhibiting a failure in the coordinated growth
of the tissue. The positioning of starch granules, the expression pattern of PIN4 and
pWOX5 and the columella stem cell identity marker J2341 similarly demonstrated the
disordered cellular architecture of this area. This phenotype displayed a variable degree
of penetrance in the different MOB1A RNAi lines and was maximal in mob1A-1,
which correlated with the higher reduction of MOB1A expression. Different
hypotheses can be advanced about the causes of the observed phenotype. Similarly to
the role of yeast MOB1 in cytokinesis, Arabidopsis MOB1A might control the correct
alignment of the cell plate during cell division. Thus, absent or reduced levels of the
protein could cause anomalous orientations of cell division, which in turn might
determine the failed alignment of cells in contiguous cell files. However, the disordered
cellular pattern was restricted to the area of the root tip around the stem cell niche.
Chapter IV
102
This observation is interesting because root stem cells are known to perform an
asymmetric cell division, which generates two cells with different fates: while one
daughter cell keeps the stem cell identity, the other undergoes differentiation
(reviewed by(Jiang and Feldman, 2005). This kind of division might involve the
asymmetric distribution of certain cellular determinants between the two daughter
cells at cytokinesis. Budding yeast also undergoes an asymmetric cell division.
Interestingly, MEN components localise to the spindle pole body that migrates into the
daughter cell during anaphase but are largely absent from the SPB that remains in the
mother cell (Bardin et al., 2000; Pereira et al., 2000). It could be therefore hypothesised
that MOB1A plays a particular role during asymmetric cell divisions in plants and the
correct fulfilment of root stem cell divisions might be particularly sensitive to MOB1A
protein levels.
In some cases, mob1A-1 and MOB1A RNAi seedlings exhibited an extremely severe
root tip phenotype characterised by morphological defects. This phenotype is
reminiscent of the effects caused by loss of Mats (Mob as tumor suppressor) function in
Drosophila, consisting in increased cell proliferation, defective apoptosis, and induction
of tissue overgrowth (Lai et al., 2005; Shimizu et al., 2008). Mats is an ortholog of yeast
Mob1 and has been involved in the Hippo (Hpo) signalling pathway, which participates
in the control of tissue growth (reviewed by(Hariharan and Bilder, 2006; Harvey and
Tapon, 2007). The morphology of the root tip is assured by the programmed cell death
of distal columella cell layers, which are progressively shed from the root cap. Similarly
to Mats role in Drosophila, MOB1A might perform a fundamental function in the
coordinated growth of columella tissue. Reduced protein levels could affect the correct
balance between cell proliferation and programmed cell death, hence causing the
observed phenotype in mob1A-1 and MOB1A RNAi roots.
The subcellular localisation of MOB1A and MOB1B supports the hypothesis that plant
MOB1 proteins might play a function in the regulation of cell division, similarly to the
situation of their orthologs in other eukaryotes. Analysis of a transgenic Arabidopsis
line constitutively expressing a MOB1A-GFP translational fusion revealed a nuclear
and cytoplasmic localisation of the protein. The nuclear targeting of MOB1A is in
Chapter IV
103
agreement with the results of Van Damme et al. (2004), who overexpressed MOB1A
fluorescent fusion in tobacco BY2 cells. As our immunocytochemistry experiments
with an antibody recognising both MOB1A and MOB1B also displayed a preferentially
nuclear localisation of the two proteins, the cytosolic signal exhibited by MOB1A-GFP
might be due to the artificial overexpression levels of the transgene. Sequence analysis
of MOB1A did not indicate the presence of any known nuclear localisation signal (data
not shown). The nuclear targeting of the protein must therefore rely on a different
mechanism, possibly the association with a second protein that drives the import into
the nucleus. Interestingly, the subcellular localisation of MOB1 proteins seemed to
follow the fate of the nuclear envelope. MOB1 nuclear localisation disappeared at the
end of prophase in parallel to nuclear envelope break-down, remained cytosolic until
the two chromosome sets were completely separated and finally reappeared around the
two new nuclei. In yeast, Mob1 associates to the SPB (Yoshida and Toh-e, 2001). As
plant cells do not possess a SPB, the nuclear targeting of Mob1 can be taken as an
argument to consider the nucleus as an alternative centre for the coordination of cell
division. Indeed, the nuclear surface is considered as the main functional plant
microtubule-organising centre and is instrumental in the formation of mitotic
cytoskeleton arrays like preprophase band and spindle (reviewed by(Schmit, 2002). The
presence
of
MOB1
proteins
in
ovules
and
root
cells,
documented
by
immunocytochemistry experiments, provide evidence for their role in the correct
development of these tissues, consistently with the phenotype of mob1A-1 and
MOB1A RNAi plants.
In summary, our results provide new insights into the function of MOB1 proteins in
plants. It emerges that MOB1 plays a role in the control of cell division and cell
proliferation, similarly to its orthologs in other eukaryotes. Our current investigations
are aiming at clarifying the exact role of MOB1 during cell division and to explore the
redundancy level among the different MOB1 genes.
Chapter IV
104
Acknowledgements
We thank F. Ditengou for providing us the pWOX5::GFP line and for helpful discussions; the NASC for
providing the SALK T-DNA mutant lines and the J2341 GFP marker line, which was generously made
available by J. Haseloff; C. Becker for assistance with genotyping of T-DNA lines and RT-PCR analysis;
X. Li for the production of PIN4 antibody; R. Nitschke and the Life Imaging Center (University of
Freiburg) for the use of confocal microscopes. We are particularly grateful to F. Santos Schröter and C.
Becker for critical reading of the manuscript and helpful comments.
References
Bardin AJ, Amon A (2001) Men and sin: what's the difference? Nat Rev Mol Cell Biol 2: 815-826
Bardin AJ, Visintin R, Amon A (2000) A mechanism for coupling exit from mitosis to partitioning of the
nucleus. Cell 102: 21-31
Baurle I, Laux T (2003) Apical meristems: the plant's fountain of youth. Bioessays 25: 961-970
Bedhomme M, Jouannic S, Champion A, Simanis V, Henry Y (2008) Plants, MEN and SIN. Plant Physiol
Biochem 46: 1-10
Bedhomme M, Mathieu C, Pulido A, Henry Y, Bergounioux C (2009) Arabidopsis monomeric Gproteins, markers of early and late events in cell differentiation. Int J Dev Biol 53: 177-185
Bothos J, Tuttle RL, Ottey M, Luca FC, Halazonetis TD (2005) Human LATS1 is a mitotic exit network
kinase. Cancer Res 65: 6568-6575
Chant J, Pringle JR (1995) Patterns of bud-site selection in the yeast Saccharomyces cerevisiae. J Cell Biol
129: 751-765
Citterio S, Albertini E, Varotto S, Feltrin E, Soattin M, Marconi G, Sgorbati S, Lucchin M, Barcaccia G
(2005) Alfalfa Mob 1-like genes are expressed in reproductive organs during meiosis and
gametogenesis. Plant Mol Biol 58: 789-807
Citterio S, Piatti S, Albertini E, Aina R, Varotto S, Barcaccia G (2006) Alfalfa Mob1-like proteins are
involved in cell proliferation and are localized in the cell division plane during cytokinesis. Exp
Cell Res 312: 1050-1064
Clough SJ, Bent AF (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of
Arabidopsis thaliana. Plant J 16: 735-743
Ditengou FA, Teale WD, Kochersperger P, Flittner KA, Kneuper I, van der Graaff E, Nziengui H, Pinosa
F, Li X, Nitschke R, Laux T, Palme K (2008) Mechanical induction of lateral root initiation in
Arabidopsis thaliana. Proc Natl Acad Sci U S A 105: 18818-18823
Frenz LM, Lee SE, Fesquet D, Johnston LH (2000) The budding yeast Dbf2 protein kinase localises to the
centrosome and moves to the bud neck in late mitosis. J Cell Sci 113 Pt 19: 3399-3408
Chapter IV
105
Friml J, Benkova E, Blilou I, Wisniewska J, Hamann T, Ljung K, Woody S, Sandberg G, Scheres B,
Jurgens G, Palme K (2002) AtPIN4 mediates sink-driven auxin gradients and root patterning in
Arabidopsis. Cell 108: 661-673
Galla G, Zenoni, S., Marconi, G., Marino, G., Botton, A., Pinosa, F., Citterio, S.,
Ruperti, B., Palme, K.,
Albertini, E., Pezzotti, M., Martin, M., Barcaccia, G. (2009) The Cell Cycle-Associated Mob1
Gene is Responsible for Defective Female Meiosis and Gametogenesis in Arabidopsis thaliana L..
Submitted
Gould KL, Simanis V (1997) The control of septum formation in fission yeast. Genes Dev 11: 2939-2951
Hariharan IK, Bilder D (2006) Regulation of imaginal disc growth by tumor-suppressor genes in
Drosophila. Annu Rev Genet 40: 335-361
Harvey K, Tapon N (2007) The Salvador-Warts-Hippo pathway - an emerging tumour-suppressor
network. Nat Rev Cancer 7: 182-191
Harvey KF, Pfleger CM, Hariharan IK (2003) The Drosophila Mst ortholog, hippo, restricts growth and
cell proliferation and promotes apoptosis. Cell 114: 457-467
Hergovich A, Stegert MR, Schmitz D, Hemmings BA (2006) NDR kinases regulate essential cell processes
from yeast to humans. Nat Rev Mol Cell Biol 7: 253-264
Hou MC, Guertin DA, McCollum D (2004) Initiation of cytokinesis is controlled through multiple modes
of regulation of the Sid2p-Mob1p kinase complex. Mol Cell Biol 24: 3262-3276
Hou MC, Salek J, McCollum D (2000) Mob1p interacts with the Sid2p kinase and is required for
cytokinesis in fission yeast. Curr Biol 10: 619-622
Hruz T LO, Szabo G, Wessendorp F, Bleuler S, Oertle L, Widmayer P, Gruissem W and P Zimmermann
(2008) Genevestigator V3: a reference expression database for the meta-analysis of
transcriptomes. Advances in Bioinformatics
Jenik PD, Gillmor CS, Lukowitz W (2007) Embryonic patterning in Arabidopsis thaliana. Annu Rev Cell
Dev Biol 23: 207-236
Jiang K, Feldman LJ (2005) Regulation of root apical meristem development. Annu Rev Cell Dev Biol 21:
485-509
Jurgens G (2001) Apical-basal pattern formation in Arabidopsis embryogenesis. Embo J 20: 3609-3616
Jurgens G (2003) Growing up green: cellular basis of plant development. Mech Dev 120: 1395-1406
Justice RW, Zilian O, Woods DF, Noll M, Bryant PJ (1995) The Drosophila tumor suppressor gene warts
encodes a homolog of human myotonic dystrophy kinase and is required for the control of cell
shape and proliferation. Genes Dev 9: 534-546
Karimi M, Inze D, Depicker A (2002) GATEWAY vectors for Agrobacterium-mediated plant
transformation. Trends Plant Sci 7: 193-195
Krapp A, Simanis V (2008) An overview of the fission yeast septation initiation network (SIN). Biochem
Soc Trans 36: 411-415
Chapter IV
106
Lai ZC, Wei X, Shimizu T, Ramos E, Rohrbaugh M, Nikolaidis N, Ho LL, Li Y (2005) Control of cell
proliferation and apoptosis by mob as tumor suppressor, mats. Cell 120: 675-685
Lee SE, Frenz LM, Wells NJ, Johnson AL, Johnston LH (2001) Order of function of the budding-yeast
mitotic exit-network proteins Tem1, Cdc15, Mob1, Dbf2, and Cdc5. Curr Biol 11: 784-788
Lippincott J, Shannon KB, Shou W, Deshaies RJ, Li R (2001) The Tem1 small GTPase controls
actomyosin and septin dynamics during cytokinesis. J Cell Sci 114: 1379-1386
Luca FC, Mody M, Kurischko C, Roof DM, Giddings TH, Winey M (2001) Saccharomyces cerevisiae
Mob1p is required for cytokinesis and mitotic exit. Mol Cell Biol 21: 6972-6983
Luca FC, Winey M (1998) MOB1, an essential yeast gene required for completion of mitosis and
maintenance of ploidy. Mol Biol Cell 9: 29-46
Mah AS, Jang J, Deshaies RJ (2001) Protein kinase Cdc15 activates the Dbf2-Mob1 kinase complex. Proc
Natl Acad Sci U S A 98: 7325-7330
Mailand N, Lukas C, Kaiser BK, Jackson PK, Bartek J, Lukas J (2002) Deregulated human Cdc14A
phosphatase disrupts centrosome separation and chromosome segregation. Nat Cell Biol 4: 317322
Mohl DA, Huddleston MJ, Collingwood TS, Annan RS, Deshaies RJ (2009) Dbf2-Mob1 drives
relocalization of protein phosphatase Cdc14 to the cytoplasm during exit from mitosis. J Cell
Biol 184: 527-539
Pan D (2007) Hippo signaling in organ size control. Genes Dev 21: 886-897
Pereira G, Hofken T, Grindlay J, Manson C, Schiebel E (2000) The Bub2p spindle checkpoint links
nuclear migration with mitotic exit. Mol Cell 6: 1-10
Sabatini S, Heidstra R, Wildwater M, Scheres B (2003) SCARECROW is involved in positioning the stem
cell niche in the Arabidopsis root meristem. Genes Dev 17: 354-358
Schmid M, Davison TS, Henz SR, Pape UJ, Demar M, Vingron M, Scholkopf B, Weigel D, Lohmann JU
(2005) A gene expression map of Arabidopsis thaliana development. Nat Genet 37: 501-506
Schmit AC (2002) Acentrosomal microtubule nucleation in higher plants. Int Rev Cytol 220: 257-289
Scholl RL, May ST, Ware DH (2000) Seed and molecular resources for Arabidopsis. Plant Physiol 124:
1477-1480
Shimizu T, Ho LL, Lai ZC (2008) The mob as tumor suppressor gene is essential for early development
and regulates tissue growth in Drosophila. Genetics 178: 957-965
Shou W, Seol JH, Shevchenko A, Baskerville C, Moazed D, Chen ZW, Jang J, Shevchenko A,
Charbonneau H, Deshaies RJ (1999) Exit from mitosis is triggered by Tem1-dependent release of
the protein phosphatase Cdc14 from nucleolar RENT complex. Cell 97: 233-244
Stegmeier F, Amon A (2004) Closing mitosis: the functions of the Cdc14 phosphatase and its regulation.
Annu Rev Genet 38: 203-232
Chapter IV
107
Tapon N, Harvey KF, Bell DW, Wahrer DC, Schiripo TA, Haber DA, Hariharan IK (2002) salvador
Promotes both cell cycle exit and apoptosis in Drosophila and is mutated in human cancer cell
lines. Cell 110: 467-478
Verma DP (2001) Cytokinesis and Building of the Cell Plate in Plants. Annu Rev Plant Physiol Plant Mol
Biol 52: 751-784
Visintin R, Craig K, Hwang ES, Prinz S, Tyers M, Amon A (1998) The phosphatase Cdc14 triggers
mitotic exit by reversal of Cdk-dependent phosphorylation. Mol Cell 2: 709-718
Vitulo N, Vezzi A, Galla G, Citterio S, Marino G, Ruperti B, Zermiani M, Albertini E, Valle G, Barcaccia
G (2007) Characterization and evolution of the cell cycle-associated mob domain-containing
proteins in eukaryotes. Evol Bioinform Online 3: 121-158
Willemsen V, Scheres B (2004) Mechanisms of pattern formation in plant embryogenesis. Annu Rev
Genet 38: 587-614
Yoshida S, Toh-e A (2001) Regulation of the localization of Dbf2 and mob1 during cell division of
saccharomyces cerevisiae. Genes Genet Syst 76: 141-147
Acknowledgements
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Acknowledgements
I would like to first thank the people that initially gave me the opportunity to
accomplish this PhD. My sincere gratitude thus goes to my thesis supervisor Prof. Dr.
Klaus Palme for welcoming me into his group and for allowing me to form my
scientific expertise through the participation in high level research projects.
I am also very grateful to Dr. Benedetto Ruperti, who first sponsored me when I was a
freshly graduated student and kept encouraging me all the way through my PhD. I
could not have achieved this without his support and friendship.
I would like to extend all my thanks to the experienced researchers who critically
supervised my work in the different projects in which I have been involved. My
gratitude goes first to Dr. Alexander Dovzhenko, who was for me a constant source of
new ideas and a partner for stimulating discussions. I am grateful to Dr. Olaf Tietz for
introducing me to several lab techniques initially new to me and inspiring me with
provocative points of view. Many thanks again to Dr. Benedetto Ruperti for the
opportunity to work on a new exciting research topic.
I want to express my heartfelt gratitude to those of my colleagues that during these
years have become real friends and shared with me good and bad moments: thanks to
Claude, Nicola, Filipa, Violante, Ines, Oscar, Yamuna, Alessandro and Sara. Many
thanks in particular to Claude, who represented for me a brilliant example of
perseverance and was always a source of helpful suggestions.
I am grateful to: all my fellow PhD students for their support and collaboration; Dr.
Franck Ditengou, Dr. Cristina Dal Bosco, Dr. Filipa Santos Schröter, Dr. Christina Neu
and Dr. Taras Pasternak for assistance and stimulating discussions; Katja Rapp for
making life in the lab much easier.
Many thanks go to Dr. Alexander Dovzhenko, Dr. Filipa Santos Schröter, Dr. Benedetto
Ruperti and Claude Becker for critical reading of this thesis and helpful comments.
I wish to thank my father and Luciana, because letting an Italian boy leave home to go
abroad is not easy at all. I am also grateful to Fabio and Mattia for their friendship and
constant encouragement.
Acknowledgements
110
Last but not least, I would like to thank Karina and Filippo for being my main
motivation in what I have done and the family I have always longed for.