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

PIN polarity and cell cycle regulation in root

development

7

Chapter II Discrete distribution of AtPIN2 in plasma membrane

domains

35

Chapter III Effects of sphingolipid biosynthesis inhibition on PIN2

polarity and root development in Arabidopsis

57

Chapter IV Analysis of AtMOB1 function in plant development 81

Acknowledgements 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 sphingolipid-

enriched 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) Sterol-

dependent 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).

PIN1PIN3PIN7

PIN1PIN4

PIN3PIN7

PIN2PIN

2PIN

2 PIN2

PIN2 PI

N2

PIN2PIN

2PIN

2 PIN2

Figure 2. Schematic representation of auxin

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.

PIN1PIN3PIN7

PIN1PIN4

PIN3PIN7

PIN2PIN

2PIN

2 PIN2

PIN2 PI

N2

PIN2PIN

2PIN

2 PIN2

PIN1PIN3PIN7

PIN1PIN4

PIN3PIN7

PIN2PIN

2PIN

2 PIN2

PIN2 PI

N2

PIN2PIN

2PIN

2 PIN2

Figure 2. Schematic representation of auxin

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.

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-to-

basal 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 (Kleine-

Vehn 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 GNOM-

LIKE1 (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 cell-

cycle 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 RETINOBALSTOMA-

RELATED 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.

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- 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 root-

hypocotyl 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-D-

glucose (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 water-

corrected 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 hexose-

proton 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.

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- 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 ABC-

type 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 sphingolipid-

enriched 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 time-

points 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 Alexa-

conjugated 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 Alexa555-

conjugated antibodies. Emission was detected at 500-550 nm for GFP and Alexa488-

conjugated 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-acyl-

sphinganine 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 t-

test, 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 PIN2-

GFP: 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 PM-

localised PIN2 while preventing its de novo synthesis.

Chapter III 71

Figure 5. Inhibition of sphingolipid biosynthesis causes a general reduction of the endocytosis rate. (A-

D) 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 (A-

H).

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 dose-

response 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.

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- 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/D-

TOPO_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

Chapter IV 88

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 anti-

MOB 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.

Primers for RT-PCR analysis

TCCTGCAAAACAAAACCAGACP7

CTAAGAAGAGCGCACCATCAGP8

Primers for T-DNA insertion lines genotyping

57

GCGTGGACCGCTTGCTGCAACTLBb1

ATATTGACCATCATACTCATTGCGK_8409

GGTGCAACCACCTTGATCTTAct2_rev57

TGTTCACCACTACCGCAGAAAct2_for

TGGACAGAGGATTTGGGTTTP1057

CGTGTCTCACTCCGATAAAGCP9

TGAGTCTCTTTGGGTTAGGP650

TGAGTCTCTTTGGGTTAGGP5

GGATTCGTGTGGCTTTCTTCP4

GGATTCGTGTGGCTTTCTTCP3

AATCAACCATGAACCTGAATCCP2

TCTGATATTAACCGCAAACGCP1

Annealing T (°C)SequencePrimer

Primers for RT-PCR analysis

TCCTGCAAAACAAAACCAGACP7

CTAAGAAGAGCGCACCATCAGP8

Primers for T-DNA insertion lines genotyping

57

GCGTGGACCGCTTGCTGCAACTLBb1

ATATTGACCATCATACTCATTGCGK_8409

GGTGCAACCACCTTGATCTTAct2_rev57

TGTTCACCACTACCGCAGAAAct2_for

TGGACAGAGGATTTGGGTTTP1057

CGTGTCTCACTCCGATAAAGCP9

TGAGTCTCTTTGGGTTAGGP650

TGAGTCTCTTTGGGTTAGGP5

GGATTCGTGTGGCTTTCTTCP4

GGATTCGTGTGGCTTTCTTCP3

AATCAACCATGAACCTGAATCCP2

TCTGATATTAACCGCAAACGCP1

Annealing T (°C)SequencePrimer

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 wild-

type 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.

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::GFP-

MOB1A 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.

43.3%33.3%20%26.7%10%Phenotype penetrance

3030303030Observed root tips

1310683Root tips with disordered cell. pattern

mob1A-16E_74G_52F_1wt

43.3%33.3%20%26.7%10%Phenotype penetrance

3030303030Observed root tips

1310683Root tips with disordered cell. pattern

mob1A-16E_74G_52F_1wt

Mob1A RNAi lines

Chapter IV 97

A B C

D

E

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.

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Acknowledgements 109

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.