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The dark side of plant growth:The role of plant hormones in apical hook

development of Arabidopsis thaliana

2012

Petra Žádníková

The dark side of plan

t growth

:The role of plan

t horm

ones in

apical hook developm

ent ofA

rabidopsis thaliana Petra Žádníková

Cover:

3D projection of apical hooks of PIN7::PIN3-YFPxDR5::GFP

Ghent University

Faculty of Sciences

Department of Plant Biotechnology and Bioinformatics

The dark side of plant growth:

The role of plant hormones in apical hook


Petra Ţádníková

Thesis submitted as fulfilment of the requirements for the degree of

PhD in Sciences: Biotechnology

Academic year 2012-2013

Promoters: Prof. Dr. Eva Benková and Prof. Dr. Jiří Friml

VIB - Plant Systems Biology

Technologiepark 927

9000 Gent, Belgium

Research presented in this thesis was performed at the Ghent University, Department of

Plant Biotechnology and Bioinformatics and VIB, Department of Plant Systems Biology,

Gent, Belgium and at Masaryk University, Laboratory of Molecular Plant Physiology,

Department of Functional Genomics and Proteomics and the Institute of Experimental

Biology, Brno, Czech Republic.

The author and promoter give the authorization to consult and copy parts of this work for

personal use only. Every other use is subjected to the copyright laws and permission to

reproduce any material contained in this work should be obtained from the author or

promoter.

Il ne faut appeler Science que l'ensemble des recettes qui réussissent toujours ("Science

means simply the aggregate of all the recipes that are always successful")

Paul Valéry

Exam commission

Promoters

Prof. Dr. Eva Benková (promotor) Email: [email protected]

Department of Plant Biotechnology and Bioinformatics, Ghent University

Department of Plant Systems Biology, VIB

Technologiepark 927

9052 Gent, Belgium

Prof. Dr. Jiří Friml (co-promotor) Email: [email protected]

Department of Plant Biotechnology and Genetics, Ghent University


Technologiepark 927

9052 Gent, Belgium

Chair

Prof. Dr. Ann Depicker Email: [email protected]



Technologiepark 927

9052 Gent, Belgium

Other members of the jury

Prof. Dr. Tom Beeckman Email: [email protected]



Technologiepark 927

9052 Gent, Belgium

Prof. Dr. Gerrit T.S. Beemster Email: [email protected]

Laboratory for Molecular Plant Physiology and Biotechnology

CGB building U

University of Antwerpen 171

2020 Antwerpen, Belgium

Dr. Stephen Depuydt Email: [email protected]



Technologiepark 927

9052 Gent, Belgium

Prof. Dr. Ingo Heilmann Email: [email protected]

Department of Stress and Developmental Biology

Leibniz Institute of Plant Biochemistry

Weinberg 3, D-06120 Halle (Saale), Germany

Prof. Dr. Herman Höfte Email: [email protected]

Institut Jean-Pierre Bourgin

Unité Mixte de Recherche 1318

Institut National de la Recherche Agronomique-AgroParisTech

Route de Saint-Cyr

78026 Versailles, France

Dr. Moritz Nowack Email: [email protected]



Technologiepark 927

9052 Gent, Belgium

Table of contents

Scope 1

Abbrevreviations 3

Chapter 1: Introduction 5

Hormonal regulation of apical hook development

Chapter 2: Role of PIN-mediated auxin efflux in apical hook 27


Chapter 3: Role of the transport-dependent auxin 65

distribution in apical hook development in Arabidopsis

Chapter 4: The role of cytokinins in apical hook development 103

Chapter 5: Conclusion and future perspectives 151

Summary 161

Acknowledgements 163

Curriculum Vitae 165

Scope

1

Scope

n the beginning, there was the Seed. The seed is a small embryo enclosed in a

seed coat, from which an entire plant emerges. During seed germination, by which a new

plant's life begins, the first organ that appears is the little embryonic root, called the

radicle. Through the radicle, the emerging seedling becomes anchored in the ground and

starts absorbing water and important nutrients from the soil. Subsequently, the embryonic

shoot arises from the seed that consists of cotyledons (preliminary leaves), the hypocotyl

(stem of germinating seedlings) and the shoot apical meristem (SAM), the founder cell

population on which the entire future life of the plant depends. The just emerged young

seedlings are very fragile. Because plants germinate in the soil in the dark, their tissues

(especially the SAM) could be damaged when they penetrate the soil to reach the surface

and to get into the light. For that reason, etiolated plants have evolved a strategy to protect

their fragile tissues from damage by bending their hypocotyls, thus, hiding the important

delicate tissues below an arch. This arch, so-called the apical hook, is formed by

differential cell division and elongation on the opposite side of the hypocotyl. To

guarantee a proper apical hook development, this process must be tightly regulated.

Several plant hormones have been identified as the key regulators of cell division and

elongation, resulting in differential growth. The individual hormonal action is governed

principally interactions with other hormones and their signaling pathways. Although the

principles of most plant hormonal interactions have been examined extensively, the

molecular basis of the hormonal crosstalk remains largely unknown.

The apical hook is a suitable model system to study the hormonal crosstalk that

controls differential growth because it is a well described developmental process that can

be followed easily in time and is regulated by several hormones, including auxin,

cytokinin, and ethylene. Therefore, the aim of this thesis was to investigate the hormonal

regulation of differential growth in the apical hook, to characterize the hormonal

interaction during this process, and to gain insights into the molecular mechanisms of

these interactions.

At the start, experimental methods had to be established to study the apical hook

development. First, real-time phenotypes were analyzed by means of the continuous

Scope

2

monitoring of seedling growth in the dark. By using a camera Canon Rebel T2i-550DH,

the whole process of the apical hook development could be recorded. Likewise, the

determination of the parameters for the confocal microscopy and the quantitative analysis

of green fluorescent protein (GFP) reporter signals for the apical hooks was also a

prerequisite. Lastly, a method based on transversal sections of banded hooks with

paraformaldehyde fixed GFP-expressing material was instituted.

The introductory chapter 1 gives an overview of the current knowledge on the

apical hook development, with focus on the role of individual hormones and their

crosstalk. Chapter 2 assesses the role of auxin in apical hook development, specifically

the contribution of auxin efflux carriers to regulate the asymmetric auxin distribution.

Moreover, in chapter 2, the involvement of ethylene during apical hook development is

presented. Chapter 3 combines computational and experimental approaches to determine

(i) which are the requirements for spatial auxin redistribution during the formation of

apical hook, (ii) how does ethylene impact on the auxin redistribution, and (iii) what is

the role of cell division during the apical hook development. Subsequently, chapter 4

explores the involvement of cytokinins in the regulation of differential growth in apical

hook development. In this chapter 4, the cytokinin and ethylene crosstalk is investigated

and their impact on the asymmetric auxin distribution as well. Concluding remarks and

future perspectives are presented in chapter 5, with a final summary highlighting the

most relevant findings.

Abbreviations

3

Abbreviations

ABA Abscisic acid

ABI5 ABA INSENSITIVE 5

ACC 1-Aminocyclopropane-1-carboxylic acid

ACO ACC OXIDASE 1

ACS ACC SYNTHASE 5

AHK ARABIDOPSIS HISTIDINE PROTEIN KINASE

AHP ARABIDOPSIS HISTIDINE PHOSPHOTRANSFER PROTEIN

ALF1 ABERRANT LATERAL ROOT FORMATION 1

ARF AUXIN RESPONSE FACTOR

ARR ARABIDOPSIS RESPONSE REGULATOR

ASA1 ANTHRANILATE SYNTHASE ALPHA SUBUNIT 1

ASB1 ANTHRANILATE SYNTHASE BETA SUBUNIT 1

Aux/IAA Auxin/indole-3-acetic acid

AUX1 AUXIN RESISTANT 1

AVG Aminoethoxy-vinylglycine

AXR AUXIN RESISTANT

BAP 6-Benzylaminopurine

BDL BODENLOS

BIN2 BRASSINOSTEROID-INSENSITIVE 2

BR Brassinosteroids

CCB1 CABBAGE/DWARF1

CK Cytokinin

CKX CYTOKININ OXIDASE/DEHYDROGENASE

COP1 CONSTITUTIVE PHOTOMORPHOGENIC 1

CPD CONSTITUTIVE PHOTOMORPHOGENESIS AND DWARF

CRE1/AHK4 CYTOKININ RESPONSE 1/ARABIDOPSIS HISTIDINE KINASE 4

CTR1 CONSTITUTIVE TRIPLE RESPONSE 1

CYCB1 CYCLIN B1

DET2 DE-ETIOLATED2,

EIL2 ETHYLENE INSENSITIVE3-LIKE 2

EIN ETHYLENE INSENSITIVE 2

ER Endoplasmic reticulum

ERS ETHYLENE RESPONSE SENSOR

ETO ETHYLENE OVERPRODUCER

ETR ETHYLENE RESPONSE

Fig. Figures

GA Gibberellic acid

GFP Green Fluorescent Protein

GUS β-Glucuronidase

HAG Hours after germination

HLS HOOKLESS

IAA Indole-3-acetic acid

IAA1 INDOLE-3-ACETIC ACID INDUCIBLE 1

IAA12/BDL INDOLE-3-ACETIC ACID INDUCIBLE 12/BODENLOS

IAA14/SLR INDOLE-3-ACETIC ACID INDUCIBLE 14/ SOLITARY ROOT

IAA3/SHY2 INDOLE-3-ACETIC ACID INDUCIBLE 3/SHORT HYPOCOTYL 2

IPT ISOPENTENYLTRANSFERASE

Abbreviations

4

KN KNOLLE

LAX LIKE-AUX1

MDR/PGP Multidrug resistance/P-glycoprotein

MS Murashige and Skoog mediaum

MTSB Microtubule-stabilising buffer

NAO 1-Napthoxyacetic acid

NPA N-1-Naphthylphthalamic acid

NPH NONPHOTOTROPIC HYPOCOTYL

PAT Polar auxin transport

PHY PHYTOCHROME

PID PINOID

PIF PHYTOCHROME-INTERACTING FACTOR

PILS PIN-LIKES

PIN PIN-FORMED

RCN1 ROOTS CURL IN NPA

RT-PCR Real-time polymerase chain reaction

SAM Shoot apical meristem

SUR1 SUPER ROOT 1

TAR2 TRYPTOPHAN AMINOTRANSFERASE RELATED 2

TCS Two-component system

TIR1 TRANSPORT INHIBITOR RESPONSE 1

WT Wild type

YUC1 YUCCA1

Abbreviations

5

Chapter 1

Introduction Hormonal regulation of apical hook

development

6

Adapted from:

Ţádníková, P., Cesarino I., and Benková E. Hormonal regulation of apical hook

development (manuscript in preparation)

Author contributions:

P.Ţ. prepared the figures; P.Ţ. and I.C. wrote the manuscript.

Picture:

Apical hooks of wild-type Col-0 seedlings, 48 hours after germination

Introduction

7

Hormonal regulation of apical hook development

Ţádníková, P1., Cesarino I

1. and Benková E.

1*

1Department of Plant Systems Biology, Flanders Institute for Biotechnology (VIB) and

Department of Plant Biotechnology and Bioinformatics, Ghent University, 9052 Gent,

Belgium.

* Author for correspondence ([email protected])

Chapter 1

8

ABSTRACT

Dark-grown dicotyledonous seedlings form a hook-like structure at the top of the

hypocotyl. To ensure correct apical hook development, this process has to be tightly

coordinated. Plant hormones have been identified as important endogenous regulators of

cell elongation and division, resulting in differential growth in several developmental

processes, including apical hook establishment. In the last decades, many studies have

unraveled the plant hormones auxin and ethylene as keys regulators of hook development.

Here, we review the current knowledge on the involvement of plant hormones, such as

auxin, ethylene, gibberellins, brassinosteroids, and cytokinins in apical hook

development. We also focus on the interaction between hormones and their impact on

proper hook establishment.

Introduction

9

In the beginning, there was a seed: early seedling development

In the life cycle of higher plants, the seed is the first stage and consists of the

embryo that is sealed in a seed coat (testa). Seed development is typically divided into

two major phases: embryo development and seed maturation (Bentsink and Koornneef,

2008). The embryogenesis, which is a morphogenetic phase, generates a relatively simple

core structure, the seedling, whereas the entire adult plant morphology is acquired in a

postgermination phase by the activities of the apical meristems (Capron et al., 2009).

Seedling development starts with seed germination when the first organ, the radicle,

emerges through the seed coat after disrupting the testa and the micropylar endosperm.

After emergence of the radicle, the embryonic shoot arises that consists of preliminary

leaves (the cotyledons) and the germinating seedling stem (the hypocotyl) carrying the

shoot apical meristem (SAM), from which the entire future plant develops (Bewley and

Black, 1994; Koornneef and Karssen, 1994). Seedling development is largely determined

by light availability. When grown in the light, seedlings develop long roots, short

hypocotyls and open cotyledons, in a process known as photomorphogenesis. However,

seedlings germinate mostlly in the dark, underneath the soil surface. In the absence of

light, the seedlings use primarily stored seed reserves for etiolated growth, so-called

skotomorphogenesis, in which they develop short roots, long hypocotyls, and closed

cotyledons (Arsovski et al., 2012). Exposure to light leads rapidly to changes in the

developmental program and provokes a shift from skotomorphogenesis to

photomorphogenesis. As dark-grown seedlings are made of very delicate tissues, they

have to shield their tender shoot apical meristem and cotyledons from damage when they

grow through the soil toward the surface to get to the light. Dicotyledonous plants have

evolved a strategy to protect this fragile shoot meristematic tissue: after embryonic shoot

emergence from the seed coat, the apical end of the hypocotyl bends over to form a hook-

like structure, the so-called apical hook (Fig. 1A and 3A) (Raz and Ecker, 1999). The

delicate tissues, cotyledons and SAM, are hidden below the apical hook and, thus, are

preserved from harm during growth through the soil. In contrast, monocotyledonous

plants form a spear-like structure, called coleoptile (Fig. 1B), that covers the cotyledon of

the embryo as well as its SAM. The coleoptile is pushed up until it reaches the soil

surface, whereafter elongation stops and the first leaves emerge (Siegel et al., 1962).

Chapter 1

10

A B

Figure 1. Mechanism protecting the shoot apical meristem

(A) Apical hook – dicotyledonous plants. (B) Coleoptile – monocotyledonous plants (pictures adapted from

http://www.ppdl.purdue.edu and unpublished data from Ţádníková)

The apical hook undergoes three distinct stages

Apical hook development follows three distinct phases: formation, maintenance,

and opening (Raz and Ecker, 1999; Vandenbussche et al., 2010; Ţádníková et al., 2010).

The formation phase starts soon after germination and is completed when the hook

curvature reaches approximately 180º, a process that takes approximately 24 hours in

Arabidopsis thaliana. Subsequently, the apical hook remains closed while the hypocotyl

undergoes elongation. Hook maintenance can be disturbed or interrupted by light that

triggers the full opening of the hook, which is typically completed in approximately 6

hours (Liscum and Hangarter, 1993; Wu et al., 2010). In dark-grown seedlings, the

maintenance phase is also followed by the hook opening, which happens normally 70–90

hours later (Vandenbussche et al., 2010; Ţádníková et al., 2010) and is over when the

hook has gradually opened to reach an angle of 0º (Fig. 2).

Figure 2. Three phases of apical hook development

(pictures adapted from Ţádníková et al., 2010. supplementary material)

Introduction

11

Stages of apical hook

The apical hook formation is orchestrated by differential growth along the apical-

basal axis of the hypocotyl, with distinct contributions of cell elongation and cell division

on the opposite sides of the structure. A few, but significant, cell division events are

localized in the apical inner side of the hook, while cell elongation is reduced and the arc

length is shorter. In contrast, cell elongation is an active process in the outer side of the

hook that permits the arc length to increase. At the basal side, the cell elongation rate is

equal between outer and inner sides due to similar cell densities and, therefore, without

curvature. Thus, the differential relative contribution of the cell elongation and cell

division mechanisms is essential for proper hook development (Raz and Koornneef,

2001).

Kinematic analysis of the hypocotyl curvature showed that the elongation rate of

apex cells on the outer side of the hook, where curvature is increasing, exceeded that of

the inner side, where curvature is decreasing. During seedling growth, the hypocotyl hook

is maintained as a morphological structure while the cells that comprise it are persistently

replaced. During the hook maintenance, convective curvature changes associated with

movement to a region with different curvature, is dominant. Once cells were displaced

past the midpoint of the hook, the rate of expansion of cells on the inner surface of the

hypocotyl exceeded the outer and caused the hypocotyl to straighten. Suggesting

importance of coordinated regulation of the rate of cell elongation is required to achieve

differential growth and formation of the apical hook (Silk and Erickson, 1978).

Regulation of apical hook development

To ensure the correct coordination of differential growth and, consequently,

proper apical hook development, a tight regulation of this process is essential. Besides the

photomorphogenic control, hook development depends on the activity of a number of

plant hormones, including auxin, ethylene, gibberellins, and brassinosteroids

(Vandenbussche et al., 2010). Additionally, the output of one hormonal pathway is

typically modulated by interactions with other hormones, thus creating a hormonal

network that consists in an important developmental regulatory system (Vanstraelen and

Benková, 2012). Nevertheless, the specific nature of the regulatory mechanism and

interactions among these regulators remains poorly understood. In the following sections,

Chapter 1

12

an overview is given on the current knowledge regarding the role of each plant hormone

involved in apical hook development as well as on the physiological and developmental

consequences of their interactions.

The role of plant hormones in apical hook development

AUXIN

Importance of auxin for apical hook development

Apical hook development has been mainly connected to auxin, because mutations

or pharmacological treatments affecting auxin accumulation, perception, transport, or

signaling impair hook development (Gallego-Bartolome et al., 2011). For example,

mutants in the auxin perception genes, such as TRANSPORT INHIBITOR RESPONSE

(TIR1) and Auxin F-box (AFB) members (Dharmasiri et al., 2005), and in the auxin

signaling pathway, such as auxin resistant1 (axr1) and nonphototropic hypocotyl4 (nph4)

(Leyser et al., 1993) present a defective hook development. Furthermore, a proper auxin

balance is crucial for a correct hook formation, because defects in hook development

occur in auxin biosynthesis mutants that overproduce auxin, such as superroot1 (sur1)

and yucca1-D (yuc1-D) (Boerjan et al., 1995; Zhao et al., 2001). In addition, treatments

with polar auxin transport (PAT) inhibitors or exogenous auxin result in a hookless

phenotype, suggesting that the establishment of an auxin gradient is essential (Lehman et

al., 1996). Auxin accumulates at the apical hook side with a slower growth rate, i.e. the

concave side (Fig. 3D) (Raz and Ecker, 1999). Therefore, differential growth and

consequent hook formation have been suggested to result from an auxin gradient formed

in the apical region of elongating hypocotyls. Moreover, whereas auxin accumulation at

the concave side is critical for formation and maintenance of the hook structure, hook

opening is correlated with the subsequent auxin diffusion from the hook zone

(Vanstraelen and Benková, 2012).

Introduction

13

Auxin transport

Polar auxin transport has been proposed to control differential growth in the apical

hook formation. Diffusion or carrier-mediated uptake regulates the auxin influx in the

cell, wheras the hormone leaves the cell through the action of efflux carriers (Kramer and

Bennett, 2006). Auxin uptake is regulated by the AUXIN/LIKE-AUXIN (AUX/LAX)

family of influx carriers (Yang et al., 2006) and the efflux carriers include the PIN-

FORMED (PIN) family (Petrášek et al., 2006) and a number of multi-drug resistant/P-

glycoprotein (MDR/PGP) ABC transporters (Blakeslee et al., 2007). Dark-grown

seedlings of Arabidopsis treated with the auxin influx inhibitor 1-napthtoxyacetic acid

reduced the hook formation rate and bending, revealing the importance of the auxin influx

during the early phases of hook formation (Vandenbussche et al., 2010). In addition,

apical hook development depends on the auxin influx carriers: developmental hook

establishment is mainly assisted by LAX3, but AUX1 seems to play a prominent role in

ethylene-regulated hook exaggeration (Vandenbussche et al., 2010). Similarly, early

treatment with the auxin efflux inhibitor 1-naphthylphthalamic acid (NPA) abolished the

asymmetric auxin distribution and hook formation, whereas transient NPA application at

the moment of maximal hook bending led to an amplified opening, highlighting the

importance of the auxin efflux in the control of particular phases of apical hook

development (Ţádníková et al., 2010). Furthermore, loss-of-function mutants revealed

that the spatiotemporal regulation of the PIN gene expression is important for apical hook

development, in which PIN3 is the key regulator of the formation and maintenance

phases, assisted by PIN4 and PIN7, whereas PIN1, PIN4 and PIN7 appear to be involved

in the regulation of the transition between the maintenance and opening phases

(Ţádníková et al., 2010).

Involvement of auxin in differential growth

A similar mechanism of auxin-dependent differential growth controls other plant

adaptive responses, such as phototropism (Fig. 3B) and gravitropism (Fig. 3C), in which

plants grow toward the light and in response to gravity, respectively. In the dark,

expression and activity of PINOID (PID), a serine-threonine protein kinase that

negatively regulates auxin signaling (Christensen et al., 2000), correlate with the apolar

targeting of PIN3 at all cell sides. Light represses the PID transcription and polarizes the

cellular localization of PIN3 in endodermis cells of hypocotyls on the shaded side,

Chapter 1

14

resulting in changes in the lateral auxin transport toward that side of the shoot, where it

promotes cellular elongation and, consequently, differential growth (Ding et al., 2011). In

response to gravisstimulation, PIN3 is polarized at the bottom side of gravity-sensing root

cells, where it will redirect the auxin flux toward the bottom and trigger gravitropic

bending (Kleine-Vehn et al., 2010). Interestingly, although the auxin-dependent tropic

growth in response to gravity results in similar developmental outputs (i.e. differential

growth resulting in bending), the effect of auxin accumulation in the target tissue is

opposite, when responses in hypocotyls and roots are compared. In hypocotyls, auxin is

redirected to the shaded side (Fig. 3E), where it promotes growth, but, in roots, auxin is

transported downward, where it inhibits the growth at the bottom side (Fig. 3F) (Friml et

al., 2003; Kleine-Vehn et al., 2010). Moreover, PIN3 asymmetry has also been described

in apical hook development, with an increased signal in cortical cells at the convex side,

presumably causing the differential transport of auxin to the concave side of the apical

hook (Ţádníková et al., 2010). Similarly to gravistimulated root, auxin asymmetric

accumulation at the concave side of apical hook inhibits cell elongation (Fig. 3D).

Distinct auxin responses might be caused by different absolute levels of auxin among

single cells, or due to differential responsiveness of specific plant tissues to auxin.

Nevertheless, the underlying mechanisms modulating differential growth caused by auxin

remain unclear.

Introduction

15

Figure 3. Auxin response during differential growth

(A) Dark grown Arabidopsis plant. (B) Arabidopsis plant displaying phototropism after irradiation by

unilateral light. (C) Arabidopsis plant displaying gravitropism after gravistimulation. (D) DR5::GUS

expression of a dark-grown seedling 36 hours after germination. (E) DR5::GUS expression in a hypocotyl

upon stimulation by unilateral light. (F) DR5::GUS expression in a root upon gravistimulation (Adapted

from Friml et al. 2002; unpublished data from P. Ţádníková and H. Rakusová) (Drawings of the angles on

hypocotyl and root axis caused by asymmetric growth illustrate differential growth).

ETHYLENE

Auxin is not the only plant hormone implicated in the regulation of differential

growth in apical hooks. Ethylene is known to prolong the hook formation phase, which

results in an exaggerated apical hook (Vanstraelen and Benková, 2012). Indeed,

exogenous ethylene (Guzman and Ecker, 1990) or ethylene overproduction mutants, such

as the ethylene overproducer (eto) (Woeste et al., 1999), resulted in an amplified hook

curvature, whereaa ethylene-insensitive mutants were hookless (Guzman and Ecker,

Chapter 1

16

1990). Hook exaggeration is part of the typical triple-response phenotype in ethylene-

treated dark-grown Arabidopsis seedlings that has been used for the isolation of ethylene

mutants and the elucidation of ethylene biosynthesis and signaling (Guzman and Ecker,

1990; Vandenbussche et al., 2010).

Auxin and ethylene crosstalk

Several lines of evidence support the existence of a crosstalk between auxin and

ethylene in apical hook development. In fact, it has become clear that hormones do not

act only in a linear pathway, but rather form a complex network of interactions and

feedback circuits that determines the final output of individual hormone activities (Fig. 3)

(Vanstraelen and Benková, 2012). Numerous studies indicate that ethylene influences

auxin transport, biosynthesis, and signaling to control differential growth in the apical

hook (Muday et al., 2012). Ethylene regulates local auxin biosynthesis in apical hook

development by stimulating the expression of the TRYPTOPHAN AMINOTRANSFERASE

RELATED2 (TAR2) gene that increases the auxin level in cotyledons and hypocotyls

(Stepanova et al., 2008; Vandenbussche et al., 2010). Similarly, ethylene enhances the

expression of the auxin influx carrier AUX1 at the concave side of the hook, thus,

contributing to the formation of an auxin maximum (Vandenbussche et al., 2010;

Vandenbussche et al., 2012). In addition, ethylene also modulates the activity of the PIN-

dependent auxin efflux machinery by transcriptional and posttranscriptional mechanisms

(Ţádníková et al., 2010).

Several components of the auxin signaling pathway are asymmetrically expressed

in apical hooks and are essential for their development, such as INDOLE-3-ACETIC

ACID3/SHORT HYPOCOTYL2 (IAA3/SHY2), IAA12, and IAA13 (Ţádníková et al., 2010).

The expression of all three genes was strongly enhanced at the concave side of the hook

by application of ethylene (Ţádníková et al., 2010; Muday et al., 2012). In Arabidopsis,

IAA3/SHY2 is a probable candidate to play a major role in the control of deetiolation

processes, including hook opening (Vandenbussche et al., 2012). Ethylene-dependent

hook exaggeration is also a consequence of influence of ethylene on molecular

components of the auxin signaling pathway. Ethylene activates the transcription of

HOOKLESS1 (HLS1), an ethylene-responsive gene that encodes an N-acetyltransferase

inhibiting the AUXIN RESPONSE FACTOR2 (ARF2) repressor and, thus, controls local

auxin concentrations (Lehman et al., 1996). Apical hook formation is completely

Introduction

17

prevented in the loss-of-function mutant hls1 (Li et al., 2004), whereas overexpression of

HLS1 causes apical hook exaggeration (Lehman et al., 1996). Therefore, the auxin signal

has been suggested to act downstream of the ethylene signaling (Vandenbussche et al.,

2010) and HLS1 to be a key molecular integrator of auxin and ethylene signaling in the

regulation of apical hook development (An et al., 2012; Vanstraelen and Benková, 2012).

Interestingly, a similar interplay between ethylene and auxin has been observed during

root elongation, in which ethylene affects auxin biosynthesis, transport, and signaling by

up-regulation of the ANTHRANILATE SYNTHASE α1 (ASA1), AUX1, and IAA1 genes,

respectively (Stepanova et al., 2005; Swarup et al., 2007). However, in roots, the response

of the PIN gene expression to ethylene differs from that in the apical hook, hinting at a

developmental context-specific character of the ethylene-mediated regulation of the PIN

expression (Vanstraelen and Benková, 2012).

In summary, several studies suggest that ethylene suppresses the mechanisms

involved in the transition from the formation to the maintenance phase by controlling the

auxin activity through (i) interaction with the auxin biosynthetic pathway; (ii) auxin

response factors; (iii) modulation of the PIN gene expression; and (iv) regulation of the

polar localization of PIN proteins (Ţádníková et al., 2010).

Chapter 1

18

Figure 4. Complex hormonal networks underlying light-regulated plant development encompassing

light (photomorphogenesis) (left) and dark-grown seedlings (right)

(A) Arrows and inhibition lines represent positive and negative interactions, respectively. Components of

auxin (green), cytokinin (CK, red), abscisic acid (ABA, blue), gibberellin (GA, purple), brassinosteroids

(BR, brown), and ethylene (gray) are highlighted. Diagrams at the bottom represent schematic drawings of

a germinating seedling (left) and light- and dark-regulated morphogenesis (right) with the impact of the

different hormones summarized. Abbreviations: IAA, indole acidic acid; SAM, shoot apical meristem

(Adapted from Vanstraelen and Benková 2012)

Introduction

19

GIBBERELLINS

Gibberellins (GAs) have also been reported to regulate proper hook development.

Dark-grown Arabidopsis seedlings with reduced GA contents show characteristics of

light-grown plants, such as inhibition of hypocotyl elongation, cotyledon opening, mis-

regulation of light-controlled genes, and loss of apical hooks (Achard et al., 2003;

Alabadi et al., 2004). In addition, the presence of the GA biosynthesis inhibitor

paclobutrazol completely prevents apical hook formation (Vriezen et al., 2004; Gallego-

Bartolome et al., 2011). These observations suggest two distinct functions for GAs in

seedling development: promotion of skotomorphogenesis, including the formation and

maintenance of apical hooks and suppression of photomorphogenesis in the dark by the

repression of ligh-regulated genes (Alabadi et al., 2004). GA signaling is necessary

during the hook formation phase and the magnitude of the hook curvature largely depends

on this activity during this early phase of hook development (Gallego-Bartolome et al.,

2011; An et al., 2012). GAs play a role in apical hook development through the

transcriptional regulation of several genes of the ethylene and auxin pathways. The

ethylene-independent activity of GAs modulates both the auxin response by directly

activating the HLS1 expression and the auxin transport by sustaining the expression of

PIN3 during the maintenance and opening phases and of PIN7 in early phases of hook

development (Gallego-Bartolome et al., 2011). Therefore, GAs act probably directly in

the regulation of the hook curvature by modulating the asymmetric auxin distribution in

the hook region.

In many cases, GA signaling is mediated by nuclear growth repressors, the

DELLA proteins that are key repressors of GA responses by the negative modulation of

the GA-regulated gene expression (An et al., 2012) and, therefore, restrain plant growth.

GAs derepress the hormone responses by stimulating DELLA polyubiquitination via a

specific Skp-Cullin-F-box (SCF) E3 ubiquitin ligase complex and its rapid degradation

via the 26S proteasome pathway (Achard et al., 2003). GAs also regulate hook formation

by trancriptional modulation of the ethylene pathway through the GA/DELLA activity. In

the absence of GAs, DELLA proteins negatively regulate the expression of the ethylene

biosynthetic genes ACC SYNTHASE8 (ACS8) and ACS5/ETO2, of which the activity

contributes to hook development, whereas GAs promote ethylene biosynthesis by

derepressing the DELLA activity and, thus, cooordinately prevent hook opening

(Gallego-Bartolome et al., 2011). In addition, ETHYLENE INSENSITIVE 3/EIN3-LIKE

Chapter 1

20

1 (EIN3/EIL1) has been shown to be the integration node linking the GA and ethylene

pathways during apical hook development (An et al., 2012). EIN3/EIL1 is the central

transcription factor in the ethylene signaling and its function in apical hook formation is

the direct activation of the HLS1 expression. Besides the direct repression of the HLS1

expression, DELLA proteins also suppress EIN3/EIL1 through protein-protein

interactions. Therefore, GAs rescue the repression of DELLA proteins on EIN3/EIL1,

whereas ethylene stabilizes EIN3/EIL1 (An et al., 2012).

BRASSINOSTEROIDS

Some studies support that brassinosteroids (BRs) interact with the auxin and

ethylene pathways during the regulation of apical hook development. The expression of

the CONSTITUTIVE PHOTOMORPHOGENESIS AND DWARF (CPD) gene, which is

involved in brassinolide synthesis, was found to be enhanced at the outer side of the hook

upon ethylene treatment. In addition, the BR biosynthesis mutants, cabbage1/dwarf1-6

(cbb1) and de-etiolated2 (det2), display the typical triple-response phenotype, except for

the open cotyledons. Dark-grown seedlings of these mutants did not present hook

amplification upon ethylene treatment, whereas the appearance of the wild type seedlings

treated with a BR biosynthesis inhibitor (brassinazole) was comparable with that of the

BR biosynthesis mutants. Moreover, the combined application of ethylene and

brassinazole suppressed the formation of an exaggerated hook, a phenotype observed

after ethylene treatment alone. These observations suggest that BRs play a role in apical

hook development downstream of the ethylene signal (De Grauwe et al., 2005).

Furthermore, exogenous BRs were found to distribute auxin homogeneously, leading to a

strongly reduced hook curvature (De Grauwe et al., 2005). BRs promote the lateral auxin

transport (from the stele to the outer layers), but negatively affect the basipetal auxin

transport (toward the root). A model has been proposed for the interaction between auxin,

ethylene, and BRs, in which all three hormones influence each other. Exogenous ethylene

treatment or endogenous accumulation alters the auxin distribution, either directly or

through BR biosynthesis, whereas the asymmetric auxin distribution might affect

ethylene biosynthesis or sensitivity at the inner side of the hook, inhibiting cell elongation

and, thus, producing an exaggerated apical hook (De Grauwe et al., 2005).

Another study suggests a model in which BRs synergistically increase the seedling

sensitivity to auxin (Vert et al., 2008). BRs might enhance the auxin responsiveness by

Introduction

21

mitigating the repression on auxin target genes. Phosphorylation by the BR-regulated

BRASSINOSTEROID-INSENSITIVE2 (BIN2) kinase results in loss of the ARF2 DNA-

binding and repression activities. Consequently, BRs would modify the gene expression

state of the co-regulated genes, whereas auxin would increase the expression through the

ARF activators, leading to synergistic transcription increases (Vert et al., 2008).

Interestingly, as mentioned above, the auxin response factor ARF2 is also a target for

HLS1 that, together with EIN3/EIL1, is the integration node linking the GA, auxin, and

ethylene pathways during apical hook development. Indeed, BRs partially rescue the

defects in hypocotyl elongation in the hls1 mutant (De Grauwe et al., 2005). Therefore,

BR-dependent ARF2 phosphorylation might be a parallel pathway, representing another

regulation layer in the complex regulatory network controlling apical hook development.

CYTOKININS

Cytokinins are important regulators of plant cell proliferation and differentiation

and also control various physiological and developmental processes in plant growth and

development, including shoot meristem formation and activities (Zhao, 2008), control of

shoot/root balance (Werner et al., 2001), lateral root morphogenesis (Laplaze et al.,

2007), leaf and cotyledon expansion (Downes and Crowell, 1998), and delay of

senescence (Riefler et al., 2006). Although cytokinins have been proposed to be involved

in the regulation of apical hook development, their exact role during this process remains

largely unknown.

Even though cytokinins were discovered half a century ago, the basic molecular

mechanisms of cytokinin biosynthesis and signal transduction have been elucidated only

recently. For example, an essential part of the cytokinin-related regulation of

development implicates the interaction with other hormones, especially auxin. Cytokinins

regulate the auxin pathway mainly by affecting the expression of their signaling

components (Vanstraelen and Benková, 2012). The cytokinin signal is perceived by the

ARABIDOPSIS HISTIDINE KINASE 3 (AHK3) receptor and transduced by the primary

cytokinin-response transcription factors ARABIDOPSIS RESPONSE REGULATOR1

(ARR1) and ARR2 that activate the IAA3/SHY2 gene, an auxin signaling repressor that

negatively regulates the auxin responses and reduces the expression of the PIN auxin

efflux carriers; thus, cytokinins redistribute auxin, stimulating cell differentiation (Dello

Ioio et al., 2008). In contrast, auxin mediates the degradation of the SHY2 protein,

Chapter 1

22

maintaining PIN activities and cell division. Therefore, the consequence of the auxin and

cytokinin interaction that regulates the root meristem size by balancing cell differentiation

and cell division results in root growth (Dello Ioio et al., 2008). In addition, the auxin

distribution and activity can also be controlled by cytokinin in a transcription-independent

manner. In this case, the cytokinin signal is transduced through the CRE1/AHK4 receptor

that modulates the endocytic trafficking and the redirection of the membrane protein

PIN1 for lytic degradation into the vacuoles. However, the molecular mechanism by

which cytokinins drive the endocytotic trafficking of PIN1 remains elusive (Marhavý et

al., 2011; Vanstraelen and Benková, 2012).

Crosstalk between biosynthesis pathways is another mechanism through which

cytokinin and auxin modulate each other‟s activities. Application or induced ectopic

biosynthesis of cytokinins rapidly increased the auxin biosynthesis in young roots and

shoot tissues of Arabidopsis. Additionally, a reduced endogenous cytokinin content as a

consequence of either the inactivation of one or more cytokinin biosynthetic

ISOPENTENYLTRANSFERASE genes or the induction of the cytokinin degradation gene

CYTOKININ OXIDASE (CKX) resulted in a decrease in auxin biosynthesis (Jones et al.,

2010). Cytokinins have been shown to directly trigger the auxin biosynthesis by

modifying the transcript abundance of several auxin biosynthetic genes. Auxin feedback

occurs by inducing the CKX-mediated degradation of cytokinins. Therefore, a regulatory

loop has been proposed for auxin-cytokinin interactions in the root apex in which

cytokinin acts as a positive regulator of the auxin biosynthesis, but auxin as a negative

regulator of the cytokinin biosynthesis (Jones et al., 2010).

Besides the crosstalk with auxin, cytokinin also interacts with ethylene to

modulate plant developmental processes. Both auxin and cytokinin alone stimulate

ethylene biosynthesis in etiolated Arabidopsis seedlings and the simultaneous treatment

with both hormones has a synergistic effect (Woeste et al., 1999). Ethylene biosynthesis

is mediated by two major steps: the conversion of S-adenosyl-methionine to 1-

aminocyclopropane-1-carboxylic acid (ACC) by ACC synthase (ACS) and the production

of ethylene from ACC by ACC oxidase (Vandenbussche et al., 2012). Some studies

suggest that auxin and cytokinin enhance the ACS function through distinct mechanisms:

auxin increases the ACS transcript levels and cytokinin stabilizes the ACS proteins

produced from these transcripts, thus accounting for the synergistic interaction (Rashotte

et al., 2005). In this process, ACS5 is the primary aim of cytokinin and its C-terminal

domain targets the protein for rapid degradation by the 26S proteasome. Cytokinin

Introduction

23

presumably blocks this targeting by promoting the phosphorylation of the C-terminal

domain by a calcium-dependent protein kinase, avoiding protein degradation (Vogel et

al., 1998; Chae et al., 2003). Cytokinin has been reported to intensify the apical hook

curvature, an effect also explained by the enhanced ethylene biosynthesis through the

abovementioned mechanism (Vogel et al, 1998, Woeste et al., 1999; Chae and Kieber,

2005, Hansen et al., 2009). Finally, cytokinins have also been shown to act synergistically

with BRs in the ethylene biosynthesis of etiolated Arabidopsis seedlings; in this case too,

these hormones probably act by distinct mechanisms (Woeste et al., 1999).

Although the effect of cytokinins on apical hook development has never been

studied in details, this regulator might also play an important role in such developmental

processes, because cytokinins interact with mostly all the hormones previously identified

to be involved in hook formation.

CONCLUSION

Several reports have demonstrated the individual hormones have overlapping

functions in specific physiological processes. Therefore, the final developmental output

results from a complex network with several layers of interactions among the different

hormones. Despite years of extensive research, still many questions remain unsolved

regarding the specific nature of the regulatory mechanism and the interaction among the

hormonal regulators during apical hook development. Future efforts could benefit from

genetic, genomic, biochemical and computational tools to gain more insight into the

molecular processes that modulate the development of this fascinating evolutionary

acquisition, the apical hook.

Chapter 1

24

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27

Chapter 2 Role of PIN-mediated auxin efflux in apical hook development of Arabidopsis thaliana

28

Adapted from:

Ţádníková et al. (2010). Role of PIN-mediated auxin efflux in apical hook development

of Arabidopsis thaliana. Development 137, 607 - 617


E.B. and P.Ţ. conceived and designed the experiments; P.Ţ. performed most of the

experiments; P.Ţ. prepared the figures, and E.B. wrote the manuscript.

Picture:

Apical hooks of ACC-treated PIN3::GFP seedlings, 48 hours after germination

Role of PIN-mediated auxin efflux in apical hook development of Arabidopsis thaliana

29

Role of PIN-mediated auxin efflux in apical hook development of Arabidopsis

thaliana

Petra Ţádníková1,2

, Jan Petrášek3,4

, Peter Marhavý1, Vered Raz

5, Filip Vandenbussche

4,

Zhaojun Ding1, Kateřina Schwarzerová

6, Miyo T. Morita

7, Masao Tasaka

7, Jan Hejátko

2,

Dominique Van Der Straeten4, Jiří Friml

1,2 and Eva Benková

1*

1Department of Plant Systems Biology, Flanders Institute for Biotechnology (VIB) and

Department of Plant Biotechnology and Bioinformatics, Ghent University, 9052 Gent,

Belgium.

2Laboratory of Molecular Plant Physiology, Department of Functional Genomics and

Proteomics, Institute of Experimental Biology, Masaryk University, 62500 Brno, Czech

Republic.

3Institute of Experimental Botany, 16502 Prague, Czech Republic.

4Unit Plant Hormone

Signaling & Bio-imaging, Department of Physiology, Ghent University, 9000 Gent,

Belgium. 5Department of Human Genetics, Leiden University Medical Center, 2300 RC

Leiden, The Netherlands. 6Department of Plant Physiology, Charles University, 12844

Prague, Czech Republic. 7

Graduate School of Biological Sciences, Nara Institute of

Science and Technology, 630-0101 Ikoma, Japan.

*Author for correspondence ([email protected])

Chapter 2

30

ABSTRACT

The apical hook of dark-grown seedlings is a simple structure developing soon

after germination to protect the meristem tissues during emergence through the soil and

that opens upon exposure to light. The differential growth at the apical hook proceeds in

three sequential steps, regulated by multiple hormones mainly auxin and ethylene. We

show that the progress of the apical hook through developmental phases depends on the

dynamic, asymmetric distribution of auxin, which is regulated by auxin efflux carriers of

the PIN family. Several PIN proteins exhibited specific, partially overlapping spatial and

temporal expression patterns of which the subcellular localization suggested auxin fluxes

during hook development. Genetic manipulation of individual PIN activities interfered

with different stages of hook development, implying that specific combinations of PIN

genes are required for the progress of the apical hook through developmental phases.

Furthermore, ethylene might modulate the apical hook development by prolongation of

the formation and strong suppression of the maintenance phase. This ethylene effect is, in

part, mediated by regulation of PIN-dependent auxin efflux and auxin signaling.


31

INTRODUCTION

In dicotyledonous plants, the apical hook protects the delicate shoot meristem as

the seedling makes its way through the soil towards the surface (Darwin and Darwin,

1881). The apical hook develops soon after germination, is maintained until seedling

emergence from the soil, and opens upon exposure to light. The formation of the apical

hook is coordinated by the localized cell division and differential cell elongation on each

side of the hook (Raz and Koornneef, 2001). Maintenance phase depends on the adjusted

differential elongation rates (Silk and Erickson, 1978) retaining the curvature fixed (Raz

and Ecker, 1999), and gradual equilibration of cell lengths at both the concave and

convex side of the hook results in its opening.

To ensure proper differential growth coordination, this process must be tightly

regulated. Plant hormones have been identified as important regulatory substances that

control cell division and differentiation, including cell elongation. Typically, the output of

one hormonal pathway is modulated by interactions with other hormones, thus creating a

hormonal network that represents an important developmental regulatory system (Coenen

and Lomax, 1997; Pitts et al., 1998; Collett et al., 2000; Vandenbussche and Van Der

Straeten, 2004). Plant hormones, ethylene and auxin, have been implicated as regulators

of differential growth in the apical hook (Lehman et al., 1996; reviewed in Lomax, 1997).

Ethylene enhances hook curvature (reviewed in Ecker, 1995) as observed in the

constitutive triple response1 (ctr1) (Kieber et al., 1993) and the ethylene overproducer

(eto) (Woeste et al., 1999) mutants or by exogenous ethylene application (Guzmán and

Ecker, 1990). However, the mechanism by which ethylene triggers differential growth is

unclear.

During apical hook development, differential auxin responses have been reported

(Friml et al., 2002b; Li et al., 2004). Mutants in the auxin perception genes TRANSPORT

INHIBITOR RESPONSE (TIR1/AFB) (Dharmasiri et al., 2005a), and in the auxin-

signalling pathway, such as auxin resistant1 (axr1) (Leyser et al., 1993), nonphototropic

hypocotyl4 (nph4) (Harper et al., 2000), or superroot1 (sur1/alf1/hls3) accumulating high

levels of free auxin (Boerjan et al., 1995; Lehman et al., 1996) exhibit reduced hook

curvature. Exogenous application of auxin and polar auxin transport (PAT) inhibitors

affected the hook curvature (Schwark and Schierle, 1992; Lehman et al., 1996),

suggesting that PAT is required for proper differential growth in the apical hook. The

PAT is regulated by the AUX/LAX family of auxin influx (Bennett et al. 1996; Yang et al.,

Chapter 2

32

2006), the PIN family of auxin efflux (Luschnig et al. 1998; Petrášek at al., 2006) carriers

and the multiple drug resistance/P-glycoprotein (MDR/PGP) family of ABC transporters

(Noh et al., 2001, Blakeslee et al., 2007). Mutants in genes regulating PAT, such as roots

curl in NPA (rcn1) (Garbers et al., 1996) and pin3 (Friml et al., 2002b) have impaired the

apical hook development and expression of MDR1/ABCB19 transporter was shown to be

increased in etiolated hypocotyls (Blakeslee et al., 2007).

As both auxin and ethylene are involved in the regulation of the apical hook

development their activities are mutually coordinated. Ethylene has been shown to

enhance apical hook bending by activating transcription of HOOKLESS1 (HLS1) that

inhibits the AUXIN RESPONSE FACTOR2 (ARF2) (Lehman et al., 1996; Li et al., 2004).

A stimulatory effect of ethylene on the auxin biosynthetic pathway ((Swarup et al.,

2007)), points to another mode of interaction on the hormone metabolism level. Several

genes of the auxin biosynthesis pathways are under the transcriptional control of ethylene,

such as ANTHRANILATE SYNTHASE α1 (ASA1) and ANTHRANILATE SYNTHASE β1

(ASB1) (Stepanova et al., 2005), or a small family of tryptophan aminotransferase-

encoding genes, TAA1 and TAR2, regulating the indole-3-pyruvic acid branch of the auxin

biosynthetic pathway (Stepanova et al., 2008; Tao et al., 2008).

Here, we show that the apical hook development depends on the auxin transport

that controls the temporal and spatial auxin distribution. Several auxin efflux carrier

genes of the PIN family exhibited partially overlapping expression patterns in the apical

hook and elimination of their activities by mutations interfered with particular phases of

the hook development. Ethylene-induced enhancement of the apical hook formation and

subsequent hook curvature exaggeration was accompanied by changes in auxin

distribution and expression of several PIN genes. This suggests that part of the ethylene

effect might be mediated by auxin and regulation of the auxin efflux.


33

RESULTS

Real-time analysis of the apical hook development

To follow the apical hook development in real time, we performed continuous

monitoring of seedling growth in the dark (for details see Material and Methods). The

hook growth was recorded continuously from the moment of germination and the angle of

curvature was measured as a parameter reflecting the momentary stage of the apical hook

development (Fig. S1A in the supplementary material). Typically, the hook underwent

three developmental phases: (i) formation, the period starting from seed germination until

the hook angle reaches 180°; (ii) maintenance, the hook remains in the fully closed stage;

and (iii) opening, when it gradually opens until an angle of 0° (Fig. 1A and Fig. S1B in

the supplementary material), (Raz and Ecker, 1999). The real-time analysis enabled us to

precisely monitor duration and kinetics through particular phases. Under our experimental

conditions, the formation phase lasted 26 h (from germination until 95% of the seedlings

exhibited a closed hook angle of 177.75°±2.50); the maintenance was typically 25 h (at

least 95% of the seedlings had a fully closed apical hook); and the opening phase took

approximately 88 h (from the start of the opening, when less than 95% of the seedlings

had a closed hook angle of 177.75°±2.25 until 95% of the seedlings exhibited an open

hook angle of 8°±5.77), confirming previous reports (Raz and Ecker, 1999).

The advantage of our approach when compared to previous analyses is that the

analysis in real-time minimizes inaccuracies caused by variability in germination time

and enables an accurate monitoring of all growth phases. This phenotype analysis is a

powerful approach to identify the developmental effects of specific mutations.

Dynamics of auxin distribution and response during apical hook development

To examine the auxin distribution during the apical hook development, we

monitored the expression of the auxin response reporters DR5::GUS and DR5rev::GFP

(Ulmasov et al., 1997; Sabatini et al., 1999; Friml et al., 2003b); their activity has been

shown to correlate well with the auxin distribution in embryogenesis and root

development (Benková et al., 2003; Friml et al., 2003b). Shortly upon germination the

DR5 reporter activity was detected in cotyledons and the boundary zone between the

cotyledons and hypocotyl; subsequently, during the hook formation, an increased activity

Chapter 2

34

of DR5 was localized in the cortex and epidermal cells at the concave (inner) side of the

hook (Fig. 1C). This pattern of the DR5::GUS activity was typical throughout the

maintenance phase. The hook opening was accompanied with a more diffused DR5 signal

in the zone underneath the hook (Fig. 1C; Vandenbussche et al. 2010).

Developmental responses to auxin are mediated by the TIR/AFB auxin receptors

of the F-box protein family (Dharmasiri et al., 2005a) directing the negative regulators of

the Aux/IAA family for degradation (Overvoorde et al., 2005) in an auxin-sensitive

manner. ARF transcription factors (Okushima et al., 2005) are released from inhibition by

the Aux/IAAs and activate the expression of auxin response genes (Dharmasiri et al.,

2005a) To determine how asymmetry in the auxin distribution is transmitted by

downstream acting factors, expression and functional analysis of genes of the auxin

signaling pathway were examined. The IAA3, IAA12, and IAA13 exhibited an

asymmetrical expression pattern similar to that of DR5, suggesting that the auxin

signaling pathway was differentially active on the concave and convex sides of the apical

hook (Fig. 1H-J). Genetic inhibition of the auxin response by expression of the stabilized

repressors in shy2-2, bdl, or iaa13 mutants prevented the hook formation (Fig. 1B and

Fig. S3A, B in the supplementary material), and strongly reduced DR5 expression in the

hook of the shy2-2 mutant (Fig. 1G). Mutations in genes coding for the ARF response

regulators (nph4/arf7 and arf19) resulted in defective apical hook development. Mild

defects in the hook formation and maintenance were revealed in the single arf19 mutant

in contrast to severe defects in the nph4-1 mutant (Fig. 1B) (Harper et al., 2000). Lack of

both NPH4 and ARF19 functions led to an additive apical hook phenotype (Fig. 1B).

ARF7::GUS expression was detected in the apical hook zone, but, did not exhibit a

pronounced asymmetry (Fig. S3C in the supplementary material).

In summary, the auxin distribution during the apical hook development is very

dynamic. An auxin maximum at the concave side of the hook curvature is gradually

established during the formation and retained during the maintenance phase, while it is

lost by the opening of the hook. Asymmetry in the auxin distribution is interpreted by the

downstream auxin signaling pathway, involving the function of Aux/IAA and ARF genes.


35

Fig. 1. Auxin distribution and response during the apical hook development

(A) Kinetics of the apical hook development in Col seedlings on MS, upon NPA, and ethylene treatment.

(B) Apical hook development in shy2-2, nph4-1, arf19 single and nph4-1, arf19 double mutants. (C, D)

DR5::GUS expression through apical hook development in MS (C), and ethylene treated seedlings (D). (E,

F) DR5::GUS expression upon NPA (E) and NPA and ethylene (F) treatment. (G) DR5::GUS expression in

the apical hook of shy2-2 mutants (H-J) IAA3::GUS (H), IAA12::GUS (I) and IAA13::GUS (J) expression

in untreated, ethylene- and NPA-treated apical hooks. ACC used as precursor of ethylene biosynthesis, MS,

Murashige and Skoog medium only, Col, control Columbia seedlings. Error bars represent standard errors

(s.e.m.).

Auxin efflux carriers are differentially expressed during apical hook development

The differential DR5 expression pattern during the apical hook development might

result from either asymmetry in the local auxin biosynthesis, or activity of the PAT. For

the TRYPTOPHAN AMINOTRASFERASE1 (TAA1/WEI8), its close family member TAR2,

and the flavin monooxygenase YUC1, a tissue-specific expression in the apical hook was

demonstrated (Stepanova et al., 2008; Vandenbussche et al., 2010). The importance of the

auxin efflux for the apical hook development was tested with the auxin efflux inhibitor

NPA (Morris, 2000). NPA abolished the apical hook formation (Fig. 1A), the asymmetric

expression of DR5::GUS (Fig. 1E), and IAA13::GUS (Fig. 1J) in the “hook zone”, and

modest increase in the ARF7::GUS expression was observed (Fig. S3D in the

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DR5::GUS MS

DR5::GUS ACC

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shy2-2x

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pSHY2::GUS MS ACC

pIAA12::GUS MS ACC

pIAA12::GUS MS ACC NPA

Chapter 2

36

supplementary material). Transient NPA applications at the moment of maximal hook

bending resulted in enhanced hook opening, suggesting that the auxin efflux is required

for the hook opening process as well (Fig. S2 in the supplementary material).

To study the spatial and temporal expression patterns of the auxin efflux

components during the apical hook development PIN::GUS reporter lines were used. We

focused on the PIN1, PIN2, PIN3, PIN4, and PIN7 subfamily of plasma membrane-

localized PIN proteins (Mravec et al., 2009).

During the hook formation, PIN1, PIN3 and PIN4 were expressed. While PIN1

expression was restricted to the central cylinder (Fig. 2A), PIN3 and PIN4 were

additionally expressed in the epidermis and cortex along the hypocotyls. The expression

zones of PIN3 and PIN4 partially overlapped, but the zone of maximal curvature was

almost free of the PIN4::GUS signal (Fig. 2C and 2D). During the maintenance phase, the

PIN4 expression changed and an increased signal was detected in the hook, preferentially

at its concave side (Fig. 2D).

In the opening phase a general reduction in the expression of most of the PIN

genes occurred (Fig. 2A, C, D), except PIN7. PIN7 expression was eliminated from the

hook and restricted to the hypocotyl zone beneath during the formation and maintenance

phases and spread during the opening upwards towards the cotyledons (Fig. 2E). PIN2

was not expressed in the apical hook (Fig. 2B).

In conclusion, during the apical hook development, a complex spatio-temporal

regulation of the PIN gene expression occurs, suggesting that specific combinations of

PIN genes might control the progress of the apical hook through particular developmental

phases.


37

Fig. 2. Differentially expressed auxin efflux carriers in control and ethylene treated apical hooks

(A-J) Expression pattern of PIN1::GUS (A,F), PIN2::GUS (B,G), PIN3::GUS (C,H), PIN4::GUS (D,I)

PIN7:GUS (E,J) during formation, maintenance, and early opening phases of the apical hook development

in MS treated seedlings (A-E), and upon ethylene treatment (F-J). ACC was used as precursor of ethylene

biosynthesis. The formation, maintenance, and opening phases corresponded to 12 h, 48 h, and 72 h after

germination, respectively.

Auxin efflux carriers are involved in regulation of apical hook development

To examine the function of individual auxin efflux carriers, hook development in

pin1, pin3, pin4, and pin7 loss-of-function mutants was analyzed in real time. The most

severe defects were found in the pin3 mutant: apical hooks of pin3 never reached the fully

closed stage and were unable to maintain the bended structure. At the moment of having

reached the maximal angle of curvature, pin3 hooks gradually progressed through the

opening phase (Fig. 3A). The kinetics of the opening phase in pin3 mutant did not

significantly differ from the control. The pin3 mutant exhibited strongly reduced DR5


MS ACC

PIN1::GUS

PIN2::GUS

PIN3::GUS

PIN4::GUS

PIN7::GUS

PIN2::GUS

PIN3::GUS

PIN4::GUS

PIN7::GUS

PIN1::GUS


B

A

C

D

E

F

G

H

I

J

Chapter 2

38

reporter expression in the bending zone, which was under the detection limit throughout

the whole process of the hook development (Fig. S4A in supplementary material). In

contrast, an enhanced DR5 signal was detected in pin3 cotyledons. This implies that

cotyledons might play a role as a source of auxin in the apical hook development,

similarly to the results of Vandenbussche et al. 2010. Additional inhibition of the auxin

efflux by NPA in pin3 mutant enhanced the DR5 signal in the cotyledons, but also at the

most apical part of hypocotyls (Fig. S4B in supplementary material). Induction of the

DR5::GUS expression upon NPA treatment, which was under the detection limit in the

MS-treated pin3 hooks, points to the existence of additional, NPA-sensitive transporters

in these tissues.

When compared to pin3, the pin4 mutant exhibited less severe defects. The pin4

apical hooks reached only slightly less than 180° (Fig. 3A) and were able to remain in the

closed stage, although the maintenance duration was shorter than that of the control hooks

(Fig. 3A). No dramatic changes were observed in the kinetics of the hook opening.

Analysis of the pin4 mutant suggests that PIN4 might be primarily involved in the control

of the maintenance or transition between maintenance and opening phases. Because of the

partially overlapping expression we tested the apical hook development in pin3, pin4

double mutants. The pin3, pin4 showed an additive phenotype and both the hook

formation and maintenance were more severely affected than in either single mutants

(compare Fig. 3C to 3A). The development in the pin7 mutant was comparable to that of

the pin4, suggesting that PIN7 might act together with PIN4 in the hook maintenance-

opening (Fig. 3A). To test their functional redundancy, we analyzed the pin4, pin7 double

mutant. Lack of both PIN4 and PIN7 affected the apical hook development more severely

than the single mutants. The pin4, pin7 were unable to form closed hooks, maintenance

phase was reduced, and opening of their hooks was delayed (Fig. 3C). The pin1 mutant

exhibited a phenotype, similar to that of pin4 and pin7, characterized by reduced

maintenance and premature transition to the opening phase (Fig. 3B).

Altogether our analysis of the pin mutants revealed that several auxin efflux

carriers participate in the regulation of the apical hook development. We could identify

PIN3 as the key regulator of the formation and maintenance phases assisted by PIN4 and

PIN7. In addition, PIN1, PIN4, and PIN7 seem to be involved in the regulation of the

transition between maintenance and opening phases.


39

Fig. 3. Apical hook development in auxin efflux carrier mutants

(A-C) Monitoring of apical hook development in pin3, pin4, and pin7 (A), pin1 (B) single mutants and

pin3, pin4 and pin4, pin7 double mutants (C) in comparison with the wild type. MS, Murashige and Skoog

medium only; Col, control Columbia seedlings. Error bars represent standard errors (s.e.m.).

Membrane localization of the PIN3-GFP suggests transport routes of auxin during

the apical hook development

Expression and mutant phenotype analyses denote PIN3 as the major auxin efflux

carrier during the hook formation and maintenance. As direction of the PAT was shown

to be determined by membrane localization of auxin efflux carriers (Wiśniewska et al.,

2006), we analyzed the membrane localization of the PIN3 protein with the PIN3::PIN3-

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Chapter 2

40

GFP and PIN3::PIN3-YFP reporter lines. Functionality of both constructs was tested

using real-time analysis. We observed that PIN3::PIN3-GFP rescued the pin3 phenotype,

but in both the pin3/PIN3::PIN3-GFP and Columbia wild-type wt/PIN3::PIN3-GFP

backgrounds, formation of the hook was delayed, possibly caused by the slight

overexpression of PIN3::PIN3-GFP (Fig. S5A in the supplementary material). Indeed, no

effect on the apical hook development was detected in the Columbia wild-type, when the

weakly expressing PIN3::PIN3-YFP line was assayed (Fig. S5B in the supplementary

material).

The PIN3 membrane localization in whole-mount and transversal sections of fixed

apical hooks was analyzed with a combination of Z stacks and three-dimensional

projection. Both PIN3::PIN3-GFP and PIN3::PIN3-YFP lines exhibited the same

expression pattern and membrane localization. On longitudinal confocal sections, the

presence of PIN3 in the cortex and epidermis was confirmed as demonstrated by the

PIN3::GUS (Fig. 4A; Fig. S7 in the supplementary material). Additionally, transversal

sections revealed the PIN3-YFP signal in endodermis (Fig. 4D¸ Fig. S8 in the

supplementary material). However, while PIN3::GUS expression between the concave

and convex sides of the hook was not obviously different, the PIN3-GFP signal was

stronger on the membranes at the convex side. Fluorescence quantification on the

transversal membranes showed a reduction in the PIN3-GFP signal by 21% (n=10) and

similarly, by 28% (n=10) on the lateral membranes at the concave in comparison to the

convex side (Fig. S6A in the supplementary material).

Membrane localization of PIN3 varied depending on tissue type and cell position

along the bending of apical hook. In endodermis, the PIN3-YFP signal was often found at

the outer and transversal plasma membranes (Fig. 4D; Fig. S8 in the supplementary

material). In the cortex, PIN3-GFP was localized on the lateral membranes pointing

towards the outer epidermal tissues and on the transversal membranes preferentially at the

basal side of cells (Fig. 4A; Fig. S7 in the supplementary material). Along the apical-

basal axis, in the epidermal cells closer to hypocotyl-root junction, PIN3-GFP was

detected predominantly on the basal, while near the cotyledons on the apical side of cells

(Fig. 4A; Fig. S7 in the supplementary material).

Next, we analyzed the impact of inhibited auxin efflux on the PIN3 expression

and protein localization. Quantification of the PIN3-GFP signal revealed that upon NPA

treatment PIN3 did not differ significantly between the convex and concave sides neither

on transversal nor on lateral membranes of cortex cells (Fig. 4C, and S6A and B in the


41

supplementary material). Furthermore, proportionally more PIN3-GFP was localized to

transversal than to lateral membranes (Fig. S6A in the supplementary material).

Our studies suggest that PIN3 participates on auxin distribution during the apical

hook development by transporting auxin in a lateral direction through the endodermis,

cortex and epidermis and basipetally towards roots. In the epidermal tissues, PIN3 seems

to regulate the auxin redistribution in both apical (towards shoot) and basal (towards root)

directions. Asymmetry of PIN3 with reduced signal on the concave side of the hook

might cause differential transport rates on two sides of the bending hypocotyl during

formation, resulting in enhanced accumulation of auxin at the concave side of the apical

hook. Loss of PIN3-GFP asymmetry, and reduction of lateral in favor of transversal

membrane localization upon NPA hint to an additional involvement of posttranscriptional

regulatory mechanisms.

Chapter 2

42

Fig. 4. Analysis of PIN3 membrane localization

(A-C) Subcellular localization of PIN3-GFP on MS (A), upon ethylene (B), and NPA (C) treatment. (D)

Transversal section through apical hook expressing PIN3::PIN3-YFP. PIN3-GFP signal (green). ACC was

used as a precursor of ethylene biosynthesis. Longitudinal confocal sections in vivo (A-C), and transversal

sections of fixed (D) apical hooks. (Arrows point to the membranes with prevalent localization of PIN3-

GFP, e-endodermis, c-cortex, ep-epidermis).

PIN1, PIN4, and PIN7 participate in auxin redistribution during hook development

Phenotype analysis of pin1, pin4, and pin7 single and multiple loss-of-function

mutants suggested involvement of respective PIN proteins in the apical hook

ec

c

ep

A B

C D

12 hours MS 12 hours ACC

36 hours MS 36 hours ACC

24 hours NPA 24 hours MS


43

development. To reveal the character of the auxin redistribution directed by these PIN

family members, their localization was studied via translational GFP fusion reporter lines.

In agreement with the PIN4::GUS, only a weak PIN4:PIN4-GFP expression was

detected during the early hook formation in the cortex and epidermal cells at the base of

hypocotyls (data not shown). With progress of the formation and in the maintenance

phase, the PIN4-GFP signal appeared in the hook curvature. However, in contrast to

PIN4::GUS expression enhanced at concave side, PIN4-GFP signal was stronger at

convex side of the hook (Fig. 2D compared to Fig. 5A; and Fig. S9, Fig. S10 in the

supplementary material). PIN4-GFP was predominantly located at the basal and partially

also at the lateral membranes heading towards the epidermis. Notably, the PIN4-GFP

signal was stronger in cortex than that in epidermis, suggesting its prevalent role in the

auxin distribution through cortex tissue. Inhibition of hook development by NPA resulted

in symmetric PIN4-GFP signal in the cortex and epidermis and enhanced localization at

the basal sides of cells (Fig. 5A).

A weak PIN7-GFP signal was detected during the formation phase in cortex and

epidermal cells of the hook curvature, which gradually increased in the zone below,

similarly to PIN7:GUS. Later developmental stages were accompanied with gradual

enhancement of the PIN7-GFP signal in the apical hooks (Fig. 5B, Fig. S11 in the

supplementary material). No signs of preferential cell membrane polarity or asymmetrical

expression between the convex and concave sides were observed. In contrast to PIN4-

GFP, the PIN7-GFP signal was significantly stronger in epidermis than in cortex. In

hypocotyls treated with NPA, PIN7-GFP was localized in a non-polar manner in

epidermal cells and a weaker signal was detected in the cortex cells (Fig. 5B).

During hook formation PIN1::PIN1-GFP was expressed neither in cortex nor in

epidermis. Only randomly, a few isolated cells in the apical hooks were found to express

PIN1-GFP (data no shown). At first, during the maintenance phase, PIN1-GFP appeared

in the epidermal cells, exclusively at the concave side of the hook (Fig. 5C, Fig. S12 in

the supplementary material), as similarly reported (Blakeslee et al., 2007). This localized

expression of PIN1 was not revealed by PIN1::GUS, probably because of the detection

limit. In NPA-treated seedlings PIN1-GFP was detected in all epidermal cells of the

“hook zone” preferentially at their transversal membranes (Fig. 5C).

In summary, auxin redistribution during the apical hook development involves the

coordinated action of several auxin efflux carriers. PIN4 along with PIN3, seems to

facilitate transport of auxin in cortex and epidermal cells at the convex side of the hook.

Chapter 2

44

PIN7 appeared to be an important regulator of the auxin efflux in the epidermis and PIN1,

besides its role in the central cylinder, seemed to be involved in the control of auxin

levels directly at the position of the auxin maxima during maintenance and opening

phases.

PIN4::PIN4-GFP MS NPA

C

B

A



Figure 5. Analysis of PIN1, PIN4 and PIN7 membrane localization

(A-C) Subcellular localization of PIN4-GFP (A), PIN7-GFP (B) and PIN1-GFP (C) on MS and NPA

medium. Localization of PIN-GFP signal (green) was examined by longitudinal confocal sections through

the whole-mount apical hooks.


45

Ethylene postpones transition between formation and maintenance phases by

interaction with the PAT machinery

Besides auxin, another major regulator of the apical hook development is the

phytohormone ethylene. To investigate the possible mechanisms involved in the ethylene-

regulated apical hook development, we analyzed (i) kinetics of the hook development and

(ii) the presumable interaction of ethylene with the auxin transport machinery.

To increase the ethylene levels, we used the precursor of the ethylene biosynthesis

ACC (Yang and Hoffman, 1984). Ethylene did not affect the hook formation kinetics,

although the final hook angle significantly exceeded 180°, but seemed primarily to

prolong the formation phase that leads to an exaggeration of the hook curvature (Fig. 1A).

The exaggerated apical hooks did not tend to remain in the stage of the maximal bending,

but after having reached the peak to gradually progress through the opening phase. The

opening kinetics was not markedly affected by ethylene (Fig. 1A, Vandenbussche et al.,

2010).

To correlate changes in hook development with auxin distribution, DR5::GUS

expression was monitored. Typically, at the beginning of the hook formation phase, the

DR5 expression pattern did not significantly differ in ACC-treated and untreated hooks

(Fig. 1D). The most apparent difference was found at the stage of maximal curvature. In

exaggerated apical hooks, DR5::GUS expression was restricted to the few cells on their

concave site (Fig. 1D; Vandenbussche et al., 2010), suggesting that the mechanisms

regulating the asymmetric auxin accumulation in cells at the concave side remain active

longer. During the opening, the asymmetric DR5::GUS expression gradually disappeared

in a manner similar to that in control seedlings (Fig. 1D). Accordingly, the enhanced

asymmetry in the expression of regulators of the auxin signaling pathway (IAA3::GUS,

IAA12::GUS, and IAA13::GUS) was observed upon ethylene treatment (Fig. 1H-J). No

obvious changes in expression of ARF7 upon ACC treatment were detected when

compared to control seedlings (Fig. S3E in the supplementary material).

Modulation of the DR5::GUS expression by ethylene might point to the

interaction of ethylene with the PAT. Our observation that NPA completely blocked the

ethylene-induced formation of exaggerated apical hooks in seedlings simultaneously

treated with ACC and NPA, supports the hypothesis that mechanisms regulating auxin

distribution might act downstream of the ethylene (Fig. 1F; Vandenbussche et al., 2010).

Chapter 2

46

Analysis of several PIN genes in ethylene-treated apical hooks revealed that their

expression was modulated. Ethylene down regulated PIN1::GUS and PIN4::GUS

transcription (Fig. 2F, I compared to Fig. 2A, D). PIN3::GUS expression sustained higher

at the time when in the untreated hooks reduced PIN3 signal was observed (Fig. 2H

compared to Fig. 2C). The expression of PIN7:GUS was omitted from the hook at the

time when in untreated seedlings it progressed to the hook zone (Fig. 2J compare to Fig.

2E). PIN2::GUS expression in the apical hook was not changed by ethylene treatment

(Fig. 2B compare to Fig. 2G).

Because of its dominating role, we closely examined hook development in pin3

mutant under increased ethylene levels. pin3 seedlings exhibited reduced sensitivity to

ethylene and never formed exaggerated apical hook (Fig. S4D). Accordingly, DR5

expression was drastically reduced in the apical hooks of pin3 seedlings, but was

enhanced in cotyledons (Fig. S4C in the supplementary material).

Next, the ethylene effects on PIN3::PIN3-GFP expression and membrane

localization were studied. Quantification of the membrane PIN3-GFP signal revealed an

increased PIN3-GFP accumulation on the cell membranes at the convex sides of the

apical hooks, similarly to control seedlings (Fig. 4B and Fig. S6A in the supplementary

material). However, in contrast to control seedlings, the asymmetry between the convex

and concave sides was enhanced for the PIN3-GFP at the lateral, but reduced at the

transversal membranes (Fig. S6 in the supplementary material). The apical localization

of PIN3-GFP in the epidermal cells in the proximity of the cotyledons was in ethylene

treated hooks more pronounced (Fig. 4B).

In conclusion, our data show that ethylene prolongs formation phase, resulting in

exaggeration of the hook curvature. Part of the ethylene-regulated apical hook

development might involve modulation of the activity of the PIN-dependent auxin efflux

machinery by transcriptional and posttranscriptional mechanisms.


47

DISCUSSION

Dynamic auxin distribution during apical hook development

Development of the apical hook depends on tightly regulated differential growth

of cells. Formation of the hook curvature is a consequence of differential growth,

resulting in shorter cells on the concave than on the convex side of the hook (Raz and

Ecker, 1999; Vandenbussche and Van Der Straeten, 2004, Vriezen et al., 2004). This

asymmetry must be maintained during the maintenance phase, while during the opening,

mechanisms gradually equilibrating cell sizes on both sides of the hook have to be

activated. Although mechanistically the process of the apical hook development is well

described, the knowledge on the regulatory molecular mechanisms involved is limited.

The plant hormone auxin has been shown to play an important role in the

regulation of differential growth that determines gravitropic or phototropic responses of

plant organs. In all cases, differential growth of cells correlated with asymmetric auxin

distribution (Friml et al., 2002b; Abas et al., 2006). Analysis of the auxin response

distribution using the DR5 reporter (Ulmasov et al., 1997) during the apical hook

development revealed a dynamic modulation of auxin levels through particular

developmental phases. Shortly after germination no asymmetric auxin distribution in

hypocotyls can be observed. At first, during the hook formation auxin maxima are

gradually established in cortex and epidermal cells at the concave side of the hook. Later,

during transition to the opening phase, this differential auxin distribution is gradually

loosened. Hence, the apical hook progresses through developmental phases that might

require mechanisms for (1) the establishment of asymmetrical auxin levels, consequently

leading to differential growth rates of cells; (2) the regulation of the formation-to-

maintenance transition and stabilization of the asymmetrical auxin distribution; (3) the

control of the maintenance-to-opening phase transition; and (4) management of the

opening process accompanied with gradual loosening of auxin maxima.

PIN-dependent auxin efflux is involved in regulation of auxin distribution during

apical hook development

Dynamic modulation of auxin levels during hook development might result from

the coordinated action of local, tissue-specific auxin metabolism, and auxin transport.

Chapter 2

48

Recently, genes involved in auxin biosynthesis have been identified (Stepanova et al.,

2008; Zhao et al., 2001) and their role in the apical hook development demonstrated

(Stepanova et al., 2008; Vandenbussche et al., 2010). Here we show that PIN-mediated

auxin efflux (Petrášek et al., 2006) is an important mechanism participating in the

dynamic distribution of auxin during the apical hook development. Inhibition of the auxin

efflux by NPA interferes with the asymmetric auxin distribution and prevents formation

of the apical hook structure. Several auxin efflux carrier genes of the PIN family are

differentially expressed during apical hook development. PIN1, PIN3, and PIN4 are

expressed primarily during the hook formation and maintenance, while PIN1 and PIN7

might additionally modulate the auxin transport during the transition from maintenance to

opening phase.

Elimination of PIN genes led to specific defects in the apical hook development.

The phenotype of the pin3 mutant supports the central role of PIN3 in the formation and

maintenance phases, while PIN1, PIN4 and PIN7 seemed to control the maintenance or

the maintenance-to-opening transition. The analysis of multiple pin3, pin4, and pin4, pin7

mutants confirmed that PIN3 is assisted by PIN4 and PIN7 during the hook formation.

Establishment of auxin maxima during hook formation

The increased accumulation of auxin in a few cells at the concave side of the hook

suggests that the formation of the maximum might require redistribution in different

directions: lateral, from the central cylinder towards cortex and epidermal cells, but also

between cortex and epidermal cells; and longitudinal, along the hypocotyl axis.

Analysis of membrane localizations of PIN3 and other PIN proteins suggests that

PIN3 in endodermis, cortex, and epidermal cells might coordinate auxin redistribution

laterally from endodermis through the cortex and epidermis layers. Along the main axis in

cortex and epidermis, auxin seems to be transported primarily in basipetal direction

towards the root. The auxin transport in the cortex cells appears to be regulated by PIN3,

along with PIN4, while in the epidermis mainly by the combined action of PIN3 and

PIN7. In addition, auxin might be transported partly in acropetal direction, as suggested

by the apical location of PIN3 in epidermal cells in the proximity of the cotyledons.

An important question is how an asymmetry can be achieved in the auxin

distribution between convex and concave sides. As shown by the quantification of the

PIN3-GFP signal, asymmetry in favor of cells at the convex side can be detected already


49

very early during the hook formation phase. Slightly later, the PIN4 protein is also

observed at the convex, but not concave, side of the hook. We hypothesize that higher

levels of PIN3 and PIN4 might increase the auxin transport rate at the outer side of the

hook and contribute to the asymmetry in distribution, resulting in auxin accumulation at

the inner side of the hook.

While in the early phases, establishment and maintenance of the auxin maxima are

crucial, later, loosening of the auxin maxima takes place to coordinate hook opening. The

asymmetric character of YUC1 expression with a maximum at the convex side of the

apical hook, as shown by Vandenbussche et al., 2010, brings to the attention importance

of the local auxin biosynthesis in this developmental phase. This probably illustrates a

mechanism of equilibration of auxin levels on the two opposite sides of the hook

curvature. On the other hand as also shown by Vandenbussche et al.,

differentiation/maturing of the vasculature in hypocotyls does not seem to be involved in

timing of the hook opening. Transient inhibition of auxin transport by NPA induces apical

hook opening, demonstrating that PAT needs to be functional also at this developmental

phase. The PIN1-GFP signal observed at the concave side of the hook, as also shown by

Blakeslee et al. (2007), suggests that PIN1 might be an important co-regulator of auxin

levels directly in the auxin maxima. The pin1 as well as pin4 and pin7 loss-of-function

mutants exhibit shortening of the maintenance phase, but a relatively weak phenotype

defect of single mutant insinuates redundancy of PIN genes in the regulation of the hook

maintenance and transition to opening.

Although the auxin metabolism and transport represent important machineries

mediating apical hook opening, the crucial question remains how exogenous stimuli; such

as light, are integrated into the initiation and regulation of the opening process.

Ethylene interaction with the PAT machinery during apical hook development

Although ethylene-induced exaggeration of the apical hook is a well-known

phenomenon (Raz and Ecker, 1999), the knowledge on the mechanisms behind is limited.

Our real-time analysis revealed that ethylene affects primarily transition from the

formation to the maintenance phase. Ethylene-treated hooks continue to bend at the time

when control hooks already passed to the maintenance phase. This leads to the absence of

the maintenance phase and direct transition of formation to opening phase.

Chapter 2

50

Ethylene stimulates the expression of the TAR2 gene involved in auxin

biosynthesis and increases auxin level in cotyledons and hypocotyl tissues (Stepanova et

al., 2008; Vandenbussche et al., 2010). These observations are pointing to an important

role of ethylene-regulated local auxin biosynthesis in the apical hook development,

similarly as has been shown for the ethylene regulated root elongation (Růţička et al.,

2007; Stepanova et al., 2007; Swarup et al., 2007). Moreover, rescue of the apical hook

phenotype in the ethylene perception mutants by exogenous auxin hints at auxin acting

downstream of the ethylene pathway (Vandenbussche et al., 2010). NPA as well as lack

of PIN3 activity counteract the ethylene effect on hook development, revealing that

modulations in the auxin levels in responsive tissues are not only mediated through an

ethylene regulated local auxin biosynthesis, but also involves an auxin transport.

Ethylene restricted expression of DR5 reporter to several cells at the concave side

of the hook (Vandenbussche et al., 2010), and enhanced expression of Aux/IAA regulators

IAA3, IAA12 and IAA13, as recently demonstrated also in tomato apical hook (Chaabouni

et al., 2009). The ethylene modulated expression of PIN genes in apical hooks shows

specificities when compared to roots. In the root meristem PIN1, PIN2, and PIN4 were up

regulated (Růţička et al., 2007), and this stimulatory effect of ethylene could also be

confirmed in the roots of etiolated seedlings (data not shown). In contrary, during the

apical hook development PIN1 and PIN4 seem to be rather down regulated while PIN2

expression is not affected in the presence of ethylene.

On the protein level, ethylene enhanced asymmetry in the PIN3-GFP signal in

favor of the lateral localization of PIN3 in the cortex cell membranes at the convex side.

In summary, our data suggest that ethylene suppresses mechanisms involved in

transition from formation to maintenance phase. This might include control over auxin

activity through (i) interaction with the auxin biosynthesis, (ii) auxin response (iii)

regulation of the PIN gene expression, and (iv) modulation of the polar localization of the

PIN proteins.

Model of auxin distribution during apical hook formation

In summary, we propose the presumable model for the auxin distribution during

apical hook development in the absence of an enhanced ethylene signal (Fig. 6). Auxin,

synthesized in the cotyledons, the shoot apical meristem, and the apical zone of the

hypocotyl, is transported basipetally towards the root (Vandenbussche et al., 2010). This


51

movement is regulated by the coordinated activities of the auxin influx carrier AUX1 that

controls the auxin flow from the apical meristem and cotyledons towards the hypocotyl

(Vandenbussche et al., 2010) and efflux carriers (PIN1 and PIN3) acting in the central

cylinder. Lateral redistribution of auxin from the endodermis towards the cortex is

promoted by PIN3 and further through the cortex and epidermis by AUX1

(Vandenbussche et al., 2010), PIN3 and PIN4. In both cortex and epidermal cells, auxin

might be basipetally directed by PIN3 and PIN4 acting predominantly in the cortex and

PIN3, PIN7, AUX1 and LAX3 (Vandenbussche et al., 2010) in the epidermis. As

suggested by the membrane localization of PIN3 in the epidermis, part of auxin might be

redirected acropetally towards the cotyledons.

The increased localization of PIN3 and PIN4 on the convex side of the hook might

contribute to the lateral asymmetry in the auxin distribution, probably by the enhanced

auxin transport rate and, consequently, in the more efficient auxin draining in the outer

(epidermal and cortex) hook layers. Later, transition to the opening phase is accompanied

by PIN1 expression at the concave side of the hook and the simultaneous progress of

PIN7 into the unfolding hook.

In addition, the enhanced asymmetrical expression of the Aux/IAA genes

(IAA3/SHY2, IAA12/BDL and IAA13) at the concave side of the hook points to the

existence of a negative feedback regulatory loop. Increased accumulation of the

IAA3/SHY2 and other homologues might contribute to the regulation of the PIN

expression (Vieten et al., 2005; Dello Ioio et al., 2008), and PIN polarity (Sauer et al.,

2006), thus reinforcing the imbalance between auxin levels at the convex and concave

sides of the apical hook. Recently, proteolytic turnover of PIN proteins was reported as an

important mechanism in the regulation of differential PIN-mediated PAT (Abas et al.,

2006; Kleine-Vehn et al., 2008). Although there is no direct evidence, discrepancy

between PIN3::GUS and PIN3::PIN3-GFP or PIN4::GUS and PIN4::PIN4-GFP

expression, respectively, might hint to this type of regulation during the apical hook

development. Ultimate conclusion on this question requires more detailed examination.

Chapter 2

52

Fig. 6. Presumable model of auxin distribution during apical hook formation

Auxin synthesized in cotyledons, shoot apical meristem and apical zone of the hypocotyl is transported in

basipetal direction towards the root (reviewed in Zhao 2010). This basipetal movement is enhanced by the

auxin influx (AUX1) that controls the auxin flow from the cotyledons and shoot apical meristem

(Vandenbussche et al. 2010) and by the efflux (PIN1 and PIN3) carriers that act in the central cylinder.

Lateral redistribution of auxin through endodermis is promoted by PIN3 and farther through the cortex and

the epidermis by AUX1, PIN3 and PIN4. In cortex, auxin is basipetaly directed by PIN3, PIN4 and in

epidermis by PIN3, PIN7, AUX1 and LAX3. In the proximity of cotyledons, part of the auxin might be

transported in apical direction by PIN3 at apical membranes in epidermis. Increased PIN3 and PIN4

localization at the cell membranes on the convex side, might enhance the auxin transport rate, resulting in a

more efficient draining of auxin in the outer cortex and epidermal layers of the hook. The asymmetric

expression of Aux/IAA genes with maxima at the concave side might be involved in a regulatory feedback

loop that reinforces the asymmetrical auxin redistribution. (Abbreviations: e-endodermis, c-cortex, ep-

epidermis).


53

MATERIALS AND METHODS

Plant material

The transgenic Arabidopsis thaliana (L.) Heynh. lines have been described

elsewhere: DR5::GUS (Sabatini et al., 1999); PIN1,2,3,4::GUS; PIN1::PIN1-GFP

(Benková et al., 2003); PIN7:GUS (Vieten et al., 2005); PIN4::PIN4-GFP; PIN7::PIN7-

GFP (Blilou et al., 2005); nph4-1; arf19-1; nph4-1, arf19-1 double mutant; ARF7::GUS

(Okushima et al., 2005); pSHY2::GUS; pIAA12::GUS; pIAA13::GUS; pBDL::bdl;

pSHY2::shy2-2; pIAA13::iaa13; pSHY2::shy2/DR5::GUS (Weijers et al., 2005); pin1-3

(Okada et al., 1991); and pin4-1 (Friml et al., 2002a); the mutant lines pin7-2, pin3-4, and

pin3-5 with sequence-indexed insertions were identified in the Signal Insertion Mutant

Library (http://signal.salk.edu/cgi-bin/tdnaexpress/). The double mutants pin4, pin7 and

pin3, pin4 (Friml et al., 2003b). The PIN3::PIN3-GFP and PIN3::PIN3-YFP plants were

obtained as follow: a 12.5-kb PIN3 genomic fragment generated from BAC F15H11 and

digested with NgoM IV, was cloned into the XmaI site of the pBIN19 binary vector. The

GFP or YFP fragment was inserted at the end of the first exon of PIN3 using the Counter-

Selection BAC Modification Kit (Gene Bridges). pin3-4 were crossed with DR5::GUS

and PIN3::PIN3-GFP lines. pin1 phenotype was analyzed on the heterozygous

population. The apical hook development was recorded, seedlings were numbered and the

pin1 homozygotes were identified based on pin-like shoot phenotypes in adult stage.

Growth conditions

Seeds were plated on half-strength Murashige and Skoog (MS) medium with 1%

sucrose, 1% agar (pH 5.7), or supplemented either with 5 µM 1-aminocyclopropane-1-

carboxylate (ACC; Sigma-Aldrich) or with 5 µM N-(1-naphtyl)phtalamic acid (NPA;

Duchefa), vernalized for 2 days at 4°C, exposed to light for 3 h at 18°C, and cultivated in

the dark at 20°C. Seedlings were processed 12, 24, 36, 48, 60, and 72 h after germination

or used for real-time phenotype analysis.

Chapter 2

54

Real-time analysis and statistics of the apical hook development

Development of seedlings was recorded at 1-h intervals for 5 days at 24°C with an

infrared light source (880 nm LED; Velleman, Belgium) by a spectrum-enhanced camera

(EOS035 Canon Rebel Xti; 400DH) with built-in clear wideband-multicoated filter and

standard accessories (Canon) and operated by the EOS utility software. Angles between

hypocotyl axis and cotyledons were measured by ImageJ (National Institutes of Health;

http://rsb.info.nih.gov/ij). Fifteen seedlings with synchronized germination start

processed.

GUS analysis

Histochemical GUS staining was done as published (Friml et al., 2003a) for

24 hours at 37°C. Seedlings mounted in chloral hydrate (Fluka) were analyzed with a

automatic virtual slide scanner dotSlide BX microscope frame (BX51 microscope;

Olympus) and 10x UPLSAPO objective equipped with a digital CCD camera (2/3” CCD

camera, 6.45 µm x 6.45 µm pixel size, high sensitivity, high resolution, Peltier cooled,

dynamic range of 3 x 12 bit). Images were processed in Adobe Photoshop.

Localization of PIN::PIN-GFP reporter proteins and PIN3-GFP quantitative

analysis

Confocal laser scanning micrographs were obtained with a Leica TCS SP2 AOBS

with a 488-nm argon laser line for excitation of the GFP fluorescence. Emissions were

detected between 505-580 nm. Using a 20x water immersion objective (NA = 1,25,

pinhole) confocal scans were done with pinhole at 1 Airy unit.

The PIN3-GFP localization was examined by confocal Z sectioning and 3D-

reconstruction. Each image represents either a single focal plan or a projection of

individual images taken as a Z series. Z-stacking was done at a 1024-scan format,

collecting at least 10 images through cortex and epidermal layers. Full Z-stack confocal

images were 3D- projected with the Leica confocal software.

The fluorescence intensity of the membrane PIN3-GFP signal was quantified with

ImageJ. At least 10 seedlings per treatment were analyzed. Images were processed in

Adobe Ilustrator.


55

Localization of PIN3-YFP on transversal sections

24 hours old PIN3::PIN3-YFP seedlings were fixed 2 hours in 3.7%

paraformaldehyde (Serva) in MTSB (50 mM PIPES; 5 mM EGTA; 1 mM MgSO4,

pH=6.8) and immobilized in 5% (w/v) water solution of low-melting agarose (Sigma).

Agarose blocks were mounted with agarose onto the stage of Motorized Advance

Vibroslice (World Precision Instruments, UK), 25 m transversal sections through the

apical hook were observed with Zeiss LSM 510 confocal microscope (Carl Zeiss Jena,

Germany).

ACKNOWLEDGMENTS

We thank Herman Höfte, Gerrit S.T. Beemster, and Karel Spruyt for help with the

establishment of the real-time analysis, Stéphanie Robert and Jürgen Kleine-Vehn for

helpful comments, and Martine De Cock for help in preparing the manuscript. This work

was supported by grants from the European Research Council (Starting Independent

Research Grant ERC-2007-Stg-207362-HCPO to E.B), the Ministry of Education, Youth

and Sports of the Czech Republic (LC06034 and MSM0021622415 to P.Z., J.H, and J.F.),

The Grant Agency of the Academy of Sciences of the Czech Republic (IAA601630703 to

P.Z., J.H. and J.F.), the Ghent University (“Bijzonder Onderzoeksfonds”, Bilateral

Cooperation” BILA-06), and the Interuniversity Attraction Poles Programme (IUAP

VI/33), initiated by the Belgian State, Science Policy Office. F.V. is a Postdoctoral fellow

of the Research Foundation-Flanders.

Chapter 2

56

SUPPLEMENTARY MATERIAL

020406080

100120140160180200

0 20 40 60 80 100 120

angl

es [°]


Col


A B

C

180° 135 ° 0 °

Figure S1. Real-time analysis of the apical hook development

(A) Angle of curvature meassured as angle between hypocotyl axis and cotyledons. (B) Apical hook

development through three distinct developmental phases: formation, maintenance, and opening. Error bars

are s.e.m. (n=10)

0

20

40

60

80

100

120

140

160

180

0 10 20 30 40 50 60 70 80 90 100 110 120

an

gle

s [°

]


MS

NPA

A

Figure S2. Enhancement of the opening phase of the hook development by inhibition of the polar

auxin transport

(A) Acceleration of the opening of the apical hook by the transient application of NPA on a closed apical

hook. Error bars are s.e.m. (n=10).


57

050

100150200

Col iaa13 Col iaa13

48 72


angl

es [°]

0

50

100

150

200

Col bdl Col bdl

48 72


angl

es [°

]

B


ARF7::GUS MS

D

C

E

ARF7::GUS NPA

ARF7::GUS ACC

A

bdl bdl

iaa13 iaa13

Figure S3. Aux/IAAs and ARFs participate on regulation of the apical hook development

(A-B) Accumulation of iaa12/bdl and iaa13 negative regulators in pBDL::bdl and IAA13::iaa13 seedlings,

respectively, intereferes with hook development. (C-E) Expression of AFR7::GUS is detected in the apical

hook during hook formation, maintenance and opening (C), modest increase upon NPA (D) and no change

upon ethylene (E) treatments. ACC used as precursor of ethylene biosynthesis. The formation phase

corresponds to 12h after germination; maintenance 48h and opening phase 72h after germination.

Chapter 2

58

0

50

100

150

200

250

48 72 92

angl

es [°]


Colpin3


DR5::GUS NPA

B

pin3xDR5::GUS NPA


DR5::GUS MS

A

pin3xDR5::GUS MS


DR5::GUS ACC

DC

pin3xDR5::GUS ACC

Figure S4. Defects in the apical hook development of pin3 mutant correlate with changed DR5

reporter expression

(A-C) DR5::GUS expression is below the detection limit in pin3mutant apical hooks on MS (A) and at

increased ethylene level seedlings throughout the whole development (C). (B) NPA treatment enhances

expression of DR5::GUS in the hook zone of hypocotyls and cotyledons. (D) The pin3 mutant exhibits

resistance to ethylene and does not form an exaggerated apical hook. ACC was used as a precursor of

ethylene biosynthesis. The formation, maintenance and opening phases correspond to 12, 48 and 72 hours

after germination, respectively.


59

020406080

100120140160180200220

0 20 40 60 80 100 120

an

gle

s [°

]


ColPIN3::PIN3-YFP/Col

020406080

100120140160180200220

0 20 40 60 80 100 120

an

gle

s [°

]


Colpin3PIN3::PIN3-GFP/ColPIN3::PIN3-GFP/pin3

B

A

Figure S5. Complementation of pin3 by PIN3:GFP during apical hook development

(A) Partial rescue of pin3 defects in apical hook development by PIN3::PIN3-GFP. Note the limited

interference of PIN3-GFP with the hook development at the maintenance phase in both wild-type (Col) and

pin3 backgrounds. (B) In contrast to PIN3::PIN3-GFP, PIN3::PIN3-YFP does not interfere with the apical

hook development in the wild-type background. WT, control Columbia seedlings. Real-time phenotype was

analyzed on 10-15 seedlings exhibiting a synchronized germination start. Error bars represent standard

errors (s.e.m.).

0

20

40

60

80

100

120

MS ACC NPA

rela

tive

flu

ores

cenc

e [%

]

BA 1 1 2 1 2 2

Figure S6. Quantification of PIN3-GFP in the apical hook

(A) Increased accumulation of the PIN3-GFP signal in the lateral and transversal cortex cell membranes at

the convex side of the control and ethylene-treated hooks. NPA treatment reduces asymmetry of the PIN3

signal between the convex and concave sides of the apical hook. (B) Quantification of the GFP signal on

lateral and transveral membranes of five cortex cells at the convex and concave sides of the hook.

Yellow lines (perpendicular to the respective membrane) indicate the position of measurements at lateral

and transversal membranes to quantify intensity of GFP signal. Apical hooks of 10 seedlings at the early

maintenance phase (24 h) were used for measurements. Error bars represent standard errors (s.e.m.).

1 - Student‟s t test P < 0.05; 2 - Student‟s t test P > 0.5.

Chapter 2

60

ADDITIONAL SUPPLEMENTARY MATERIAL

Supplementary material containing movies is available at:

http://dev.biologists.org/lookup/suppl/doi:10.1242/dev.041277/-/DC1

Figure S7. PIN3::PIN3-GFP expression in the apical hook

PIN3-GFP membrane localization analysis using Z sectioning and three-dimensional

reconstruction. Seedling in formation phase.

Figure S8. PIN3::PIN3-YFP expression in the apical hook

PIN3-YFP membrane localization analysis on transversal section using Z sectioning and

three-dimensional reconstruction. Seedling in formation phase.



reconstruction. Seedling in formation phase.



reconstruction. Seedling in maintenance phase.




Figure S12. PIN1::PIN1-YFP expression in the apical hook

PIN1-YFP membrane localization analysis using Z sectioning and three-dimensional



61

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Role of the transport-dependent auxin distribution in apical hook development in Arabidopsis

65

Chapter 3


66

Adapted from:

Ţádníková, P., Wabnik, K., Friml, J., Prusinkiewicz, P. and Benková, E:. Role of the

transport-dependent auxin distribution in apical hook development in Arabidopsis

(manuscript in preparation)


E.B. and P.Ţ. conceived and designed the experiments; P.Ţ. performed all experiments;

P.Ţ. prepared the figures; and P.Ţ., K.W. and E.B. wrote the manuscript.

Picture:

Apical hooks of wild-type Col-0 seedlings stained with SCRI Renaissance 2200, 48 hours

after germination


67

Role of the transport-dependent auxin distribution in apical hook development in

Arabidopsis

Ţádníková, P.1,2,3*

, Wabnik, K.1,2*

, Friml, J.1,2,3

, Prusinkiewicz, P.3 and Benková, E.

1,2

1Department of Plant System Biology, VIB, 9052 Gent, Belgium

2Department of Plant Biotechnology and Bioinformatics, Ghent University, 9052 Gent,

Belgium

2Laboratory of Molecular Plant Physiology, Department of Functional Genomics and

Proteomics, Institute of Experimental Biology, Masaryk University, Brno, Czech

Republic

3Department of Computer Science, University of Calgary, Calgary, Alberta, T2N 1N4,

Canada

*These authors contributed equally to this work

Chapter 3

68

ABSTRACT

Auxin is a major plant growth regulator of which the asymmetric distribution

controls differential growth during the plant's lifetime. A fascinating example of

differential growth in plants is the formation of apical hooks that protect the fragile shoot

apical meristems when they reach the soil surface during seed germination. The

asymmetric auxin distribution or auxin gradient relies on the concerted action of PIN-

FORMED (PIN) auxin efflux carriers and is crucial for the developmental processes, such

as the apical hook formation. However, the detailed mechanisms underlying the

developmental interpretation of these auxin gradients into the differential cellular growth

are still unclear. Here, we combined in silico and in vivo approaches to examine the

minimal requirements for the auxin gradient-guided asymmetrical growth during the

formation of apical hooks in Arabidopsis thaliana. We demonstrate that the asymmetric

expression of PIN genes at the concave (inner) versus convex (outer) sides of the hook is

a driving force for the auxin gradient establishment during apical hook development.

Predictions of the apical hooks from a computer model were validated through

experimental analyses of the auxin response and PIN dynamics in both wild-type and

mutant plants. Our model presents how the auxin gradient could result in the anisotropic

growth of the apical hook and how the hook curvature is primarily established through

interplay between the PIN-dependent polar auxin transport and the auxin-mediated

growth dynamics. Finally,using computer modelling, we demonstrate that the plant

hormone ethylene influences the PIN protein activity and the cell proliferation pattern

during apical hook development, thereby modulating the apical hook curvature.


69

INTRODUCTION

Germination, the process by which seedling emerge from seed coats, followed by

the beginning of growth, occurs in the dark underneath the ground. Plants have evolved

mechanisms to protect the fragile shoot apical meristem when breaking through the soil

surface; namely, shortly after germination, the hypocotyls create hook-shaped structures

that shield the apex meristem. The apical hook is formed by differential growth through

the combined activities of cell elongation and cell division on the opposite sides of the

hypocotyl and is maintained as long as the hypocotyl elongates in the dark. Upon

irradiation of the seedling, the apical hook straightens out, a process accompanied by

expansion and greening of the young cotyledons (Raz and Ecker, 1999; Raz and

Koornneef, 2001; Ţádníková et al., 2010).

The plant hormones auxin and ethylene have been recognized as ones of the major

regulators of apical hook development (Ecker, 1995; Lehman et al., 1996). The apical

hook is formed by the establishment of an auxin maximum at the concave side of the

apical hook and alterations of this auxin maximum affect apical hook development; e.g., a

reduced curvature is displayed in the auxin biosynthesis mutant yucca1 (yuc1), the auxin

overproduction mutant superroot1 (sur1) and the auxin signaling mutant auxin resistant1

(axr1) and non-phototrophic hypocotyl (nph4) (Leyser et al., 1993; Boerjan et al., 1995;

Lehman et al., 1996; Tian and Reed, 1999; Zhao et al., 2001). Similarly, genetic or

chemical interference with the auxin transport results in a defective apical hook

phenotype (Schwark and Schierle, 1992; Lehman et al., 1996), indicating that the polar

auxin transport is required for proper apical hook bending. Thus, the precise auxin

distribution and its response are crucial for the asymmetric growth and proper

development of the hypocotyl. However, the presumable mechanisms remain unclear that

translate transport-dependent auxin gradients into this differential growth.

Ethylene-treated etiolated Arabidopsis seedlings display a „triple response‟ that

includes thickening and shortening of the hypocotyl with an exaggerated apical hook. The

identical response is obtained with the mutants constitutive triple response1 (ctr1) and

etylene overproducer (eto), whereas ethylene-insensitive mutants are hookless or exhibit

severe defects in apical hook development (Guzman and Ecker, 1990; Kieber et al., 1993;

Raz and Ecker, 1999). Ethylene and auxin might partially act independently on the

downstream targets genes; however, they tend to crossregulate their biosynthesis

pathways and, thus, crosstalk with their signalling pathways (Li et al., 2004; Stepanova et

Chapter 3

70

al., 2005; Stepanova et al., 2007). Ethylene promotes the expression of TRYPTOPHAN

AMINOTRANSFERASE2 (TAR2) gene and, accordingly, the auxin levels increase in

ethylene-treated apical hooks (Vandenbussche et al., 2010). Additionally, the ethylene

interaction with auxin on the metabolic level also impacts on the auxin transport

presumably by the modulation of the expression of several PIN genes (Ţádníková et al.

2010). Another convergence point of the ethylene and auxin pathways is represented by

the HOOKLESS 1 (HLS1) expression. Ethylene activates the transcription of HLS1, a key

hook bending-promoting transcription factor promoting that consequently inhibits the

AUXIN RESPONSE FACTOR 2 (ARF2) gene (Lehman et al., 1996; Li et al., 2004), a

negative auxin response regulator.

Apical hooks are dynamic structures of the development depends on tightly

controlled and coordinated cell divisions and differential growth occurring at the most

upper part of growing hypocotyls. Although the role of auxin and its differential auxin

distribution is well established, the mechanism is still unknown by which auxin integrates

division and differential growth programs during apical hook development.

Here, we combined experimental and computational approaches to assess the

minimal requirements for the precise spatiotemporally controlled auxin redistribution

during the apical hook formation. First, we demonstrated the vital importance of the axial

asymmetry of the PIN expression in hypocotyl tissue layers, including the cortex and

epidermis, for generating spatially focused auxin maxima in the epidermis at the concave

side of the hook. Second, by integrating cell growth mechanics in our computer model,

we identified prominent mechanisms that couple polar auxin distribution to cell division

and differential growth to regulate the apical hook curvature. Finally, these computer

model predictions were validated by experimental observations of seedlings under control

conditions as well under conditions perturbing the normal apical hook formation process,

such as exposure to high ethylene levels. Based on both experimental data and model

predictions, we propose that hook curvature is primarily controlled through the hormonal

crosstalk that links the PIN-dependent polar auxin transport to the auxin-modulated cell

growth dynamics.


71

RESULTS

The auxin response maximum is centred in the epidermal cells at the concave side of

the apical hook

The auxin response gradient in the apical hook seems to participate in the

establishment of the hook curvature (Li et al., 2004; Vandenbusche et al., 2010;

Ţádníková et al., 2010) because an interference with this gradient results, in extreme

cases, in impaired hook bending (Vandenbusche et al., 2010; Ţádníková et al., 2010; ).

However, the precise positioning and the detailed mechanism of the auxin maxima

formation are not fully understood (Vandenbussche et al., 2010; Ţádníková et al., 2010).

Previously, based on detailed analyses of the expression of the PIN auxin efflux

carriers in the apical hook, we had proposed that PIN3 directed auxin from the central

cylinder through the endodermis toward the cortex and epidermal tissue layers. Auxin

transport in the cortex cells appears to be regulated by PIN3, along with PIN4, whereas in

the epidermis it is mainly controlled by the combined action of PIN3 and PIN7. As

revealed by the quantification of the PIN3-green fluorescent protein (GFP) signal,

asymmetry in favor of cells on the convex side was detected very early during the hook

formation phase. Slightly later, the PIN4 protein also occurred at the convex, but not

concave, side of the hook. Based on these results, we hypothesized that increased levels

of PIN3 and PIN4 might enhance the auxin transport rate at the outer side of the hook,

thereby contributing to the asymmetry in the distribution that results in auxin

accumulation at the inner side (Ţádníková et al., 2010). To validate our hypothesis, we

utilized a computer model approach that integrated experimentally derived data on the

PIN polar localizations associated with apical hook formation (Ţádníková et al., 2010).

The cellular template for the model simulations was derived from confocal microscopy

images with digital segmentation algorithms (MorphoGraphX, template slide) (see

supplementary methods for more details) (http://www.morphographx.org/) superimposed

on transversal cross sections of the apical hook.

As the exact mechanisms are unknown on how the PIN expression is controlled at

the concave and convex sides of the hook, we conceived a simple formula translating the

cell position within the whole hook crosssection into the strength of the PIN expression in

this cell (Model A) (See supplementary methods) as inferred from the experiments

(Ţádníková et al., 2010). Therefore, the orientation of each individual cell with respect to

Chapter 3

72

the position on the concave side of the apical hook was determined by the relative level of

the PIN protein activity; accordingly, the farther the cell from the concave side of the

hook, the higher the PIN expression (see supplementary methods; Fig. S1A in the

supplementary material). Subsequently, to supply auxin to our system, we assumed that

auxin was provided from the vascular tissues (Fig. 1A; green asterisk marking the auxin

source) from where it was subsequently transported to the outer cell layers (Fig. S1C-S1F

in the supplementary material). Based on the experimental observations, we initially

made the assumption that PIN3 and PIN4 were differentially regulated in the cortex

tissues with the highest expression at the convex hook side (Ţádníková et al., 2010).

Hence, model A predicted the establishment of the auxin maximum in the cortex on the

concave side of the apical hook (Fig. S1A, in the supplementary material). Interestingly, a

virtual introduction of differential PIN expression patterns in both the cortex and

epidermis (Model B) led to the highest auxin accumulation in the epidermis on the

concave side of the apical hook (Fig. 1A; Fig. S1B in the supplementary material).

To dissect which of the mathematical predictions corresponded best with the auxin

distribution pattern in the apical hook, we visualized the auxin response with the synthetic

auxin-responsive promoter DR5 fused to GFP. Surprisingly, the transverse sections

through the apical hook revealed that auxin presumably accumulated in the epidermis

with only a weak activity in a few adjacent cortex cells on the concave side of the hook

(Fig. 1B), similarly as predicted by Model B (Fig. S1B and S1D in the supplementary

material). Thus, the asymmetric PIN expression in both the cortex and epidermis tissues,

as predicted by the model, seems to be required to center the maximal auxin responses in

a few epidermis cells on the concave side of the hook (Fig. 1A; Fig S1B and S1F in the

supplementary material).


73

Figure 1. Auxin distribution pattern in the apical hook predicted by the computational model

(A) Based on PIN expression pattern as extrapolated from longitudinal confocal sections in the apical hook

(Ţádníková et al., 2010), the computational model predicts the establishment of the auxin maxima in the

epidermal cells at the concave side of an apical hook. PIN1 is expressed in the vascular tissue marked by a

green star (auxin source). PIN3 is expressed in the endodermis, cortex, and partially in the epidermis. PIN4

and PIN7 are expressed in the epidermis. The auxin content of the cells is presented in green; the

cumulative PIN protein activity (PIN3, PIN4, and PIN7) is presented in red/yellow. (B) Transverse section

of the apical hook, confirming the DR5::GFP auxin response maxima in the epidermal cells at the concave

site of the apical hook.

PIN4 and PIN7 are expressed asymmetricallyin the epidermal cells

According to the computer model, the experimentally observed auxin

accumulation pattern in epidermal cells at the concave side of the apical hook requires the

asymmetric expression of PIN genes both in the cortex and epidermis (Fig. S1B in the

supplementary material). To identify which of the auxin afflux carriers are involved in the

redistribution of auxin through the epidermis, we analyzed the expression of PIN3, PIN4,

and PIN7 homologs. In contrast to the enhanced expression of PIN3 observed in the

cortex cells at the convex versus the concave side of the hook structure (Ţádníková et al.,

2010), the PIN3 expression in the epidermal cells did not differ significantly between the

concave and convex sides (Fig. 2A, 2B). Subsequently, we tested other PIN proteins, such

as PIN4 and PIN7 that are active during apical hook development (Ţádníková et al.,

2010). Notably, the PIN4-GFP membrane signal at the convex hook side was visibly

enhanced in contrast to the weak signal in both cortex and epidermis at the concave side

(Fig. 2D, 2E). Similarly to the expression of PIN4, the expression of PIN7-GFP in the

epidermis was significantly stronger than that in the cortex and higher in the epidermal

cells at the convex than at the concave side (Fig. 2G, 2H). Taken together, our findings

Chapter 3

74

identified PIN4 and PIN7 as suitable candidates that could promote and coordinate the

asymmetric auxin distribution within the epidermis during apical hook formation.

PIN3::PIN3-GFP MS

PIN3::PIN3-GFP ACC

PIN4::PIN4-GFP MS

PIN4::PIN4-GFP ACC

PIN7::PIN7-GFP MS

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Figure 2. PIN3-GFP, PIN4-GFP, and PIN7-GFP in epidermal cells of the apical hook

(A, D, and G) Quantification of the PIN3-GFP, PIN4-GFP, and PIN7-GFP signals, respectively, on

transverse membranes of five epidermal cells at the concave and convex sides of untreated (MS) and

ethylene (ACC)-treated apical hooks. (B, E, and H) Untreated and (C, F, and I) ethylene-treated apical

hooks expressing PIN3-GFP, PIN4-GFP, and PIN7-GFP, respectively. VSNI GenStat 14th was used for

the statistical analysis (p-value for t-test 0.05). Stars indicate statistically significant differences between

columns. Apical hooks of 10 seedlings at the early maintenance phase (26 hours) were used for

quantification of the PIN-GFP membrane signal. Error bars represent standard errors (A, D). PIN3-GFP,

PIN4-GFP, and PIN7-GFP signals were quantified on longitudinal confocal sections through apical hooks.

Pictures were reconstructed from z-stacks (B, C, E, and F).


75

PIN4 and PIN7 coordinate the auxin flux through the epidermis and promote the

formation of auxin maxima

Using computer simulation, we tested to what extent the spatial differences in the

PIN protein activity from the convex to the concave sides would affect the asymmetric

distribution of auxin in the apical hook. Based on the model predictions, a compromised

asymmetry in the PIN proteins through the epidermis would result presumably in a

scattered auxin accumulation pattern (Fig. S2 in the supplementary material). To verify

these predictions (Fig. 1A, 3A; Fig. S2 in the supplementary material), we investigated

whether the loss of individual PIN protein activities affected the auxin distribution. The

number of cells with the DR5 signal on the transverse sections of at least 15 Arabidopsis

hooks was scored and the proportion was calculated of the cells with the DR5 signal

relative to the total number of cells in the hook epidermis (Fig. 3M). Quantitative

evaluation of the DR5-GFP signal demonstrated that approximately 44% of cells had a

detectable DR5-GFP signal in the wild-type plants (Fig. 3B, 3N). In case of the pin3

mutant, no significant decrease or increase in the number of epidermal cells with a DR5-

GFP signal was observed (42.09%) (Fig 3E, 3N), despite the overall decrease in the DR5

signal detected previously in pin3 hooks (Ţádníková et al., 2010) (Fig. S4A, S4C in the

supplementary material). In contrast, the pin4 and pin7 mutants displayed an increased

proportion of the cells containing the DR5-GFP signal: a significant increase by 7.7%

(49.8% of cells with an observable DR5 signal) (Fig. 3H, 3N) and by 11.91% (54% of

cells with an observable DR5 signal) (Fig. 3K, 3N) for pin4 and pin7, respectively. These

results were comparable to those observed in the computer model simulations (Fig 3A,

3D, 3G, 3J, 3N). Additionally, the computer model of the pin4 and pin7 mutants that

incorporated 1/4 of the default parameter values of the PIN4 and PIN7 transporter rates

predicted a slightly higher percentage of auxin-containing cells (58%) than that of each of

the single mutants, hinting at a possible functional redundancy between the PIN4 and

PIN7 proteins (Fig 2N). Taken together, both experimental data and model predictions

suggest that the establishment of the confined auxin maximum in the apical hook requires

the asymmetric PIN protein production in both the cortex and the epidermis.

Chapter 3

76

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pin3xDR5::GFP MS pin3xDR5::GFP ACC




77

Figure 3. PIN-dependent auxin distribution in the epidermis of an apical hook

(A, D, G, and J) Computational model prediction of the auxin distribution in untreated apical hooks of wild-

type (A), pin3 (D), pin4 (G), and pin7 (J) mutant plants. (B, E, H, and K) DR5::GFP expression monitored

on transverse sections of untreated apical hooks in wild-type (B), pin3 (E), pin4 (H), and pin7 (K)

backgrounds. (C, F, I, and L) DR5::GFP expression monitored on transverse sections of ethylene (ACC)-

treated apical hooks in wild-type (B), pin3 (E), pin4 (H), and pin7 (K) backgrounds. (M) Scheme

illustrating the scoring of the DR5-positive cells in the epidermis - the proportion of the cells with a DR5

signal was relative to the total cell number in the epidermal layer of apical hook. (N) In vivo versus in silico

quantifications of the proportion of the DR5-positive epidermal cells in untreated and ethylene-treated wild-

type, pin3, pin4, and pin7 mutant plants, respectively. Seedlings were fixed in 3.7% paraformaldehyde and

sectioned 26 hours after germination. The heating map for the auxin and PIN levels in the computer model

is as in Fig. 1A and represents the DR5::GFP expression (green) and the autofluorencence (red). VSNI

GenStat 14th was used for the statistical analysis (p-value for t-test 0.05). Stars indicate statistically

significant differences between columns. Apical hooks of 10 seedlings at the early maintenance phase (26

hours) were used for measurements. Error bars represent standard errors.

Auxin gradient in the hook is modulated by ethylene

Besides auxin, the plant hormone ethylene in known to play a role in the

regulation of the apical hook development. Defects in the ethylene signaling prevent hook

formation, whereas increased ethylene levels prolong the formation phase of the apical

hooks and lead to exaggerated hooks (Vandenbussche et al., 2010; Ţádníková et al.,

2010). Ethylene contributes to the regulation of the apical hook development through

different mechanisms, involving (i) promotion of auxin biosynthesis by the upregulation

of the TAR2 expression (Vandenbussche et al., 2010) and (ii) modulation of the

asymmetric auxin distribution, eventually affecting establishment and positioning of the

auxin maxima during apical hook development (Vandenbussche et al., 2010; Ţádníková

et al., 2010) and as demonstrated (Vandebusche et al., 2010), auxin maxima in ethylene-

treated apical hooks are restricted to fewer cells along the hook curvature than in

untreated hooks.

To determine the impact of ethylene on the auxin gradient across the apical hook,

we analyzed DR5-GFP-expressing seedlings that were treated with the ethylene precursor

1-aminocyclopropane-1-carboxylic acid (ACC). The proportion of cells with the DR5-

GFP signal in the epidermis was significantly higher (6.6%; n=15) in ethylene-treated

than in untreated apical hooks (Fig. 3C). Notably, these diffused auxin maxima in

ethylene-treated seedlings were reminiscent of those observed in the pin4 and pin7

mutants (Fig 3H, 3K).

Detailed examination of the ethylene effect on the PIN3, PIN4, and PIN7

expression in the epidermis revealed a dramatic reduction in the PIN4-GFP signal on the

Chapter 3

78

convex and concave sides in ethylene-treated apical hooks (Fig. 2D, 2F). As the PIN3

expression had significantly increased at the convex side only, ethylene might enhance

the PIN3 asymmetry between the convex and concave sides (Fig. 2A, 2C), whereas the

expression of PIN7-GFP increased as well, but, in contrast to PIN3, on both sides of the

apical hook. Thus, largely in agreement with the previous report demonstrating the

ethylene interference with the auxin transport (Ţádníková et al., 2010), ethylene might

interfere with the asymmetric PIN expression between convex and concave sides of the

hook (Fig. 2G, 2I). PIN4, together with PIN7, appeared to be the central auxin efflux

carrier controlling auxin transport and auxin accumulation in the epidermis. Thereby, the

overall downregulation of PIN4, along with the asymmetrical disruption in the PIN7

expression by ethylene, might contribute to the expansion of the auxin maxima in

ethylene-treated hooks. Indeed, treatment of the pin4 and pin7 mutants by ethylene did

not widen the auxin maxima towards convex side when compared to untreated hooks

(Fig. 3I, 3L, 3N), whereas pin3 displayed amplified auxin maxima, similarly to its control

(Fig. 3F, 3N). This observation implies that the ethylene-mediated control of the PIN4

and PIN7 expression might be an important mechanism to regulate the polar auxin flux

within the epidermis.

The dynamic computer model predicts cell elongation and cell division patterns

during apical hook formation

The apical hook curvature is established in the most upper parts of the hypocotyls

immediately after germination. Although auxin appears to be one of the central

developmental cues regulating hook curvature formation, the mechanistic information

remains scarece on this dynamic process that depends on tightly coordinated cell division

and differential cell elongation. To tackle the mechanisms of the hook structure

establishment, we developed a dynamic computer model to test the minimal requirements

for the formation of the hook structure (Fig. 1A, Fig. 4). We used a simplified

longitudinal representation of the apical hook starting from a few cells (represented by

boxes) to mimic a seedling stage (Fig. 4). Experimentally observed PIN localizations

were incorporated into the hook model (Vandenbussche et al., 2010; Ţádníková et al.,

2010) to control the auxin flow through different vascular and endodermal cell layers

(Fig. 4A). Next, we defined a flexible „hook zone‟ at a proximal distance from the

cotyledons, where the hook bending occurs in planta (Fig. 4A). This ‟hook zone‟


79

represents a developmental window (window width is defined by two distance thresholds;

see supplementary material) that exhibits asymmetric PIN protein activities and is

dynamically adapted as the hypocotyl grows and divides. To couple auxin distributions to

cell growth, we assumed that high concentrations of auxin inhibit cell elongation (see

supplementary methods).

Finally, our model integrates the experimentally observed cell proliferation pattern

that was inferred from the expression of the B-type cyclin 1 (CYCB1;1) reporter that

marks cells in the S2-to-M transition phase (Raz and Koornneef, 2001) (Fig. 5) and the

KNOLLE (KN) reporter expressed exclusively during cytokinesis and shows a cell-plate

localization (Reichardt et al., 2007) (Fig. S5 in the supplementary material). Similarly to

previous report (Raz and Koornneef, 2001), we observed that only cells that were at a

close distance from the cotyledons had an enhanced proliferation activity (Fig. S5 in the

supplementary material). For full model details, we refer to the supplementary material.

The model simulations of growing and dividing hypocotyls predicted the gradual

formation of auxin concentration maxima on the concave or inner side of the apical hook

(Fig. 4B), as observed in planta (Ţádníková et al, 2010). Interestingly, in our model, the

establishment was expected of an auxin concentration gradient spanning across the

transversal direction, hinting at a possible auxin-mediated anisotropy of cell growth

(auxin-deficient cells elongated rapidly) that would result in the mechanical bending of

the apical hook (Fig. 4B).

Next, we tested in silico whether the suppressed PIN-dependent auxin

redistribution affected the hook curvature formation. We simulated the multiple

pin3xpin4xpin7 mutant by reducing the overall PIN transport rates to 1/4 of their default

values (Fig. 4C). The model predicted, in agreement with our experimental observations

(Ţádníková et al., 2010), the reassembling of the diffused auxin maxima through both the

convex and concave sides of the hook, with a less pronounced cell growth anisotropy and

impaired apical hook bending as a consequence (Fig. 4C). Additionally, we found

through the model simulations that the strength of the auxin-mediated cell growth

anisotropy (see supplementary material) determined the degree of apical hook curvature

(Fig. 4E, 4F). Therefore, the model indicates that the coupling of the PIN-dependent

asymmetric auxin distribution to the auxin-modulated growth anisotropy could suffice to

establish the apical hook curvature that is observed in wild-type plants (Fig. 4B).

To substantiate the model predictions, we tested whether the feedback between the

polar auxin transport and the auxin-modulated cell growth could lead at any of the

Chapter 3

80

conditions to the exaggerated hook curvature, as observed in ethylene-treated seedlings.

We hypothesized that the amplification of the hook might be supported by (i) a more

confined and possibly stronger auxin maximum due to a presumably ethylene-mediated

misregulation of the PIN3, PIN4, and PIN7 efflux carriers (Fig. 2) (Vandebusche et al.,

2010); (ii) an extended cell proliferation caused by ethylene hyperactivity (Fig. 5F, 5G;

Fig. S5A, S5C in the supplementary material), preferentially on the concave side of the

hook (Fig. S5F, S5G in the supplementary material); and (iii) an ethylene-mediated

increase of the auxin levels (Vandenbusche et al., 2010) that might feedback on the cell

division pattern.

To distinguish between these different possibilities, we first tested whether more

spatially confined auxin maxima in our model (Fig. 4H) would enhance growth

anisotropy and, thus, the mechanical hook exaggeration (Ţádníková et al., 2010).

Interestingly, our simulations indicated that this scenario is not enough to explain the

ethylene effect on the hook bending (Fig. 4H). We found that the implementation of

experimentally observed (Fig. 5F, 5G) scattered cell proliferation patterns in ethylene-

treated hooks (Fig. 4D) was sufficient to reproduce the exaggerated hook phenotype (Fig.

4I). These findings indicate that, besides the effect of ethylene on the auxin transport,

other factors, such as regulated cell divisions, might be critical to obtain an amplified

apical hook structure.

We attempted to test whether linking of high auxin concentrations to a high cell

proliferation frequency could similarly reproduce the ethylene effect on the hook

curvature. Under this condition, our model reassembled a “wild-type” like pattern of the

apical hook (Fig. 4I) similar to that predicted with the model lacking the positive effect of

auxin on cell divisions (Fig. 4B). Nevertheless, our model could not reproduce

exagerrated hook phenotypes reminiscent of ethylene effects. Notably, an introduction of

ethylene effect on auxin biosynthesis (Vandenbusche et al., 2010) that was virtually

mimicked in our model by increasing overall auxin content in the hook (Fig 4J), was

sufficient to reassemble hyperbending of the apical hook and thus their exaggerated

phenotypes (Fig 4J). Moreover, these model predictions were not affected by virtual

restriction of auxin maxima (Fig 4K).

Taken together, our experimental and modelling results suggest that hook

curvature is primarily controlled through feedback between PIN-dependent polar auxin

transport and auxin-mediated cell growth and cell proliferation. Our findings suggest that

ethylene, might interfere with both polar auxin transport and auxin-mediated cell growth


81

and cell proliferation . Our findings suggest that ethylene appears to interffer with both

polar auxin transport by regulating PIN protein expression and cell division patterning,

presumably through modulation of auxin biosynthesis, thereby altering the hook

formation.

Chapter 3

82

Figure 4. Dynamic computer model of apical hook formation

(A) Computer-simulated formation of apical hook from an early developmental phase, represented by an

initial block of cells (seedling stage). The developed apical hook was divided in three developmental zones

corresponding to the hypocotyl, apical hook, and cotyledons. Cells associated with the apical hook zone

display differential PIN protein expression pattern that reflects dynamics in the regulation of the PIN

expression in this zone. Mean auxin concentrations in adjacent cells was used to estimate local growth rate

based on nonlinear logistic function (see supplementary methods). Experimentaly-deriven polar subcellular

localizations of PIN proteins in the hypocotyl was integrated in the model. Auxin concentrations and PIN

protein activities are shown in green and red, respectively. (B-F) Model simulations of „wild-type-

like‟hooks (B),a virtual simulation of pin4pin7pin3 mutant (C), model with extended zone of rapidly

dividing cells (mimics ethylene treatment) (D), weak anisotropy of axial growth (E), enhanced anisotropy

of axial growth (F). (G, H) Simulations of model with virtual restriction of auxin maxima without (G) and

with (H) extended cell proliferation pattern. (I) Coupling simulation of auxin feedback on both cell

elongation and cell division. (J) Simulation of ethylene effect on elevated auxin levels in the coupled

feedback model (I). (K) Virtually; the same as (I), but with a confined auxin maxima.


83

CycB::GUS

CycB::GUS


Figure 5. Cell division during apical hook development in Arabidopsis

(A-D) CycB::GUS expression during formation (A), maintenance (B, C), and opening (D) phases of apical

hook in unterated seedlings (MS). (E-H) CycB::GUS expression during formation (E), maintenance (F, G),

and opening (H) phases of apical hook in ethylene-treated seedlings.

Figure 6. Model of auxin redistribution during apical hook formation

(A, B) Model of redistribution of auxin in untreated apical hook (MS) (A) and treated by ethylene (ACC)

(B). Auxin synthesized in the cotyledons and the shoot apical meristem is transported in the direction of the

root (Vandenbussche et al., 2010). This flow takes place in the central cylinder and is coordinated by

activities of PIN1 along with PIN3. The auxin transport through the endodermis toward the cortex and

epidermis is driven mainly by the PIN3 and PIN4 carriers (according to previous model) (Ţádníková et al.,

2010). Under normal conditions, the PIN4 and PIN7 proteins, of which the activity is enhanced on the

convex side of the apical hook , promote the auxin redistribution toward the concave side in epidermal cells

(A). At increased ethylene levels (B), the decreased PIN4 expression and reduced asymmetry of PIN7

between the convex and concave sides might weaken the auxin flow toward the concave side in epidermis,

resulting in broadening the auxin maxima.

Chapter 3

84

DISCUSSION

Asymmetric PIN expression in the cortex and epidermis regulates the establishment

of auxin gradients in the apical hook

The formation of the apical hook in the upper part of hypocotyls is central for the

protection of early seedling development and depends on tightly controlled differential

cell growth on opposite sides of the hypocotyl (Lehman et al., 1996; Vandenbussche et

al., 2010; Ţádníková et al. 2010). The regulation of cell elongation in hypocotyls has been

proposed to depend on differential auxin distribution and downstream auxin responses.

Auxin responses at the concave side of the apical hook are in excess of the optimal level

(Evans et al., 1994; Vandenbussche et al., 2003), implying an inhibition of the cell

elongation, possibly through the ethylene signaling pathway (Abel et al., 1995;

Tsuchisaka and Theologis, 2004). In contrast, low auxin responses at the convex side of

the hook presumably promote cell elongation.

The strong repressing effect of the auxin transport inhibitor N-1-

naphthylphthalamic acid (NPA) on the apical hook development demonstrated that the

polar auxin transport is necessary to establish auxin gradients during apical hook

development. Here, we focused on the central components of the polar auxin transport

machinery, the auxin efflux carriers of the PIN family that determine the polar auxin

transport in the apical hook zone (Vandenbussche et al., 2010; Ţádníková et al., 2010).

Previously, we have proposed that the asymmetric expression of PIN3 and PIN4 in the

cortex cells might promote the auxin accumulation at the concave side of the apical hook

(Ţádníková et al., 2010). Nevertheless, the precise cellular location of the auxin maxima

the hooks was unknown until now.

To assess the requirements for the transport-mediated auxin distribution during the

apical hook formation, we combined computer modeling and experimental validation of

the model predictions. Initially, the model predictions based on our experimental data

(Ţádníková et al., 2010) hinted at the establishment of auxin response maxima in the

cortex cells at the concave side of the apical hook. However, the implementation of an

alternative model that accounts for the asymmetric PIN expression in both cortex and

adjacent epidermis led to a new prediction of the maximal auxin responses, primarily in

the few epidermal cells at the concave side of the apical hook, as further validated with

the DR5 auxin response reporter in planta. Furthermore, computational simulations of


85

various auxin transport mutants predicted that the manipulation of the PIN expression

probably affected the auxin maxima and apical hook bending. These predictions were

validated by in vivo genetic studies, highlighting the important role of the precise auxin

distribution in the regulation of the apical hook curvature. We revealed that the auxin

carriers PIN4 and PIN7 display asymmetric expression patterns in the epidermis and,

thus, probably participate in the creation of an auxin gradient in the apical hook.

Moreover, the pin4 and pin7 mutants had impaired and scattered auxin distribution

patterns, similar to those predicted by our computer model simulations.

Taken together, our experimental and theoretical results demonstrate how the

directional PIN-driven polar auxin flow is translated into auxin gradients with

morphogenic properties that guide anisotropic cell growth, thereby determining the apical

hook curvature. The central component of this mechanism is the PIN-mediated auxin

accumulation in the epidermal cells at the concave side of the apical hook that is

presumably governed by the combined action of the auxin transporters PIN3, PIN4,

and/or PIN7.

Ethylene modulates the auxin distribution by regulating the PIN protein activity

Alterations in ethylene responses have a strong impact on the apical hook

formation. An increase in ethylene leads to the typical apical hook exaggeration

phenotype, whereas ethylene response mutants display defects in the apical hook

formation (Raz and Ecker, 1999: Vandenbussche et al., 2010). Ethylene might control

the apical hook formation through combine the modulation of auxin synthesis and polar

auxin transport (Vandenbussche et al. 2010, Zadnikova et al., 2010).

Notably, besides its function in the regulation of biosynthesis and the auxin efflux

transporters, we found that ethylene dramatically affects cell proliferation patterns.

Localization studies revealed that the cell division markers CYCB1;1::GUS (Raz and

Koornneef, 2001) and KNOLLE::GFP (Reichard et al. 2007) are enriched along the

primary growth axis in ethylene-treated plants. This ethylene effect, which was unaltered

by the inhibition of auxin transport is of critical importance for formation of the

exaggerated apical hook structure.

Taken together, through computer model simulations, we have identified the

minimal requirements for obtaining wild-type-like and exaggerated hook phenotypes.

Chapter 3

86

This minimal framework links an auxin-mediated anisotropic cell growth to ethylene-

promoted cell proliferation.

We propose that plants utilize hormonal crosstalk mechanisms to modulate the

polar auxin transport and, thus, the auxin gradients that would effectively feedback on the

cell growth anisotropy and, hence, the whole organ curvature. In addition, this process

probably requires the simultaneous regulation of the cell division patterning to assure

enough cell material for such a flexible mechanical tissue bending during the

developmental patterning.


87

MATERIAL AND METHODS

Plant Material

The transgenic Arabidopsis thaliana (L.) Heynh. lines have been described

previously: DR5rev::GFP (Blilou et al., 2005); PIN3::PIN3-GFP (Ţádníková et al.,

2010); PIN4::PIN4-GFP; PIN7::PIN7-GFP (Blilou et al., 2005), CYC::GUS (Ferreira et

al., 1994), DR5::GUS (Sabatini et al., 1999), KN::KN-GFP (Reichardt et al., 2007) and

pin3-4, pin4-3, pin7-1 (Ţádníková et al., 2010) as well as the double mutants pin4 pin7

and pin3 pin4 (Friml et al., 2003) and pin3 pin7 (Blilou et al., 2005). The mutants pin3-4,

pin4-3, and pin7-1 were crossed with the lines DR5::GFP (Ulmasov et al., 1997).

Growth conditions

Seeds were surface sterilized with ethanol, plated on half-strength Murashige and

Skoog medium (Duchefa) with 1% sucrose, 0.8% agar, pH 5.7; 5 µM 1-

aminocyclopropane-1-carboxylic acid (ACC; Sigma-Aldrich) or 5 µM N-(1-

naphtyl)phtalamic acid (NPA; Duchefa) was added to the media. Plates were vernalized

for 2 days at 4°C, exposed to light for 12 hours at 18°C to synchronize the germination

start, and cultivated in the dark at 18°C. Seedlings were imaged 24 hours after

germination by confocal microscopy, stained with GUS, and used either for real-time

phenotype analysis or for transverse sectioning.

Confocal Microscopy

For confocal microscopy images, Zeiss LSM 710 with Zeiss C-Apochromat

63x/1.20 water immersion objective and Leica TCS SP2 AOBS with HC PL APO

20x/0.70 water immersion objective were used. The GFP signal after a 488-nm argon

laser line excitation was detected in the spectral range from 500 nm to 590 nm for the

Zeiss and from 505 nm to 580 nm for the Leica system. The yellow fluorescent protein

(YFP) signal after 514-nm argon laser line excitation was detected in the spectral range

from 525 nm to 600 nm for the Zeiss system.

Chapter 3

88

Quantitative analysis

The maximum projection confocal-based pictures were reconstructed with the

Zeiss ZEN 2009 software from full z-stack of longitudinal whole-mount, 24-hour-old

etiolated seedlings. These pictures were used for quantitative analysis. Optical sections

were taken through the cortex and epidermal layers. The fluorescence intensity of the

PIN3-GFP and PIN7-GFP signals was quantified on transverse membranes at the convex

and concave sides of the apical hooks with ImageJ (NIH; http://rsb.info.nih.gov/ij). At

least 10 seedlings were analyzed per treatment. VSNI GenStat 14th was used for the

statistical analysis (p-value for t-test was 0.05). Error bars represent standard errors.

Transverse sections

DR5::DR5-GFP (24-hour-old) seedlings or mutant seedlings crossed with

DR5::DR5-GFP were processed as described previously (Ţádníková et al., 2010). The

DR5-GFP signal was quantified; the number of cells with the DR5 signal was divided by

the total number of cells in the apical hook; for example, 14 cells with a signal out of 29

is equal to 48% (Fig. 4A). VSNI GenStat 14th was used for the statistical analysis (p-

value for t-test was 0.05). Error bars represent standard errors.

Real-time analysis of apical hook development

Seedling development was recorded at 1-hour intervals for 7 days at 18°C with an


(Canon Rebel T2i, 550DH, EF-S 18-55 mm, IS lens kit with built-in clear filter

wideband-multicoated and standard Canon accessories) and operated by the EOS utility

software. Angles between the hypocotyl axes and cotyledons were measured by ImageJ

(NIH; http://rsb.info.nih.gov/ij). Fifteen seedlings with synchronized germination start

were processed. Error bars represent standard errors.

Histochemical analysis of GUS activity

Histochemical β-glucuronidase (GUS) staining was done as described previously

(Friml et al., 2003; Ţádníková et al., 2010). The staining reaction was incubated at 37ºC

http://rsb.info.nih.gov/ij


89

in the dark for 8 hours. Seedlings mounted in chloral hydrate (Fluka) were analyzed with

an automatic virtual slide-scanner microscope frame (dotSlide BX51 microscope;

Olympus) and 10_UPLSAPO objective equipped with a digital CCD camera (2/3_ CCD

camera, 6.45-6.45 µm pixel size, high sensitivity, high resolution, Peltier cooled, dynamic

range of 3-12 bit). Images were processed with Adobe Photoshop.

ACKNOWLEDGMENTS

We thank Martine De Cock for help in preparing the manuscript. This work was

supported by grants from the European Research Council (Starting Independent Research

Grant ERC-2007-Stg-207362-HCPO to E.B).

Chapter 3

90

SUPPLEMENTARY FIGURES

Figure S1. Dynamic changes of the PIN expression in response to gravity

(A) Dynamic PIN3 gene expression in the cortex tissues of apical hooks led to a preferential accumulation

of auxin in the cortex at the concave side of the apical hook. A function was used to map the relative

position of each cell center vector with respect to the gravity vector: the angle between the cell center vector

and the gravity vector and dot product of these two vectors (MODEL A). (B) Simulation of the dynamics of

the PIN3 expression in both the cortex and epidermis, leading to the preferential auxin accumulation in the

epidermis (MODEL B). Color codes for the auxin concentrations and PIN expression levels are as in Fig.

1. (C and D) Time evolution of simulated with model B that incorporates the expression dynamics of PIN3,

PIN4, and PIN7.


91

0

2

4

6

8

10

12

14

10089796858483727176

Au

xin

(a.

u)

Percentage of auxin containing cells

symmetric

MS

weak

A

Figure S2. Role of differential PIN4 and PIN7 expression in the epidermis

(A) Percentage of auxin-containing cells depending on the strength of the differential PIN protein activity in

the epidermis. (B-D) Positive regulation of the auxin distribution by the differential PIN protein activity in

the epidermis.

Chapter 3

92

Figure S3. Dynamical computer model of the apical hook formation

Simulated formation of the apical hook from the seedling stage, represented by an initial block of cells.

Cells associated with the apical hook zone are sensitive to gravity, as reflected in the dynamical regulation

of the PIN expression in this zone. Auxin concentrations in cells were mapped to a given growth rate by

means of a logistic function (See supplementary methods). Auxin concentrations and the PIN protein

activity are shown in green and red, respectively. (A) Dynamic time-lapse model simulations of „wild-type-

like‟ (B) and of pin4pin7pin3 mutant (C) with broad zone of rapidly dividing cells (ethylene treatment). (D)

Dynamic time-lapse model simulations coupling auxin feedback on cell elongation and cell division. (E)

Dynamic time-lapse model simulations of the ethylene effect on elevated auxin levels in the coupled

feedback model.


93

Figure S4. Quantification of the strength of DR5-GFP expression at the concave side of the hook

(A) Quantification of the strength of the DR5::GFP expression in the middle cell at the concave side of the

hook in wild-type and pin3, pin4, and pin7 mutant backgrounds. (B-E) DR5-GFP expression during the

formation phase of the apical hook in untreated control seedlings (B) and pin3 (C), pin4 (D), and pin7 (E)

mutants. (F-I) DR5-GFP expression during the formation phase of the apical hook in ethylene-treated

control seedlings (F) and pin3 (G), pin4 (H), and pin7 (I) mutants.

Chapter 3

94

Figure S5. Cell division during apical hook development in Arabidopsis

(A) Length of the cell division zone of apical hooks of seedlings untreated (MS) and treated with ethylene

(ACC), NPA (NPA), and ethylene and NPA (NPA+ACC) in 48 hours after germination. (B-E)

KNOLLE::GFP expression in apical hooks of seedlings untreated (MS) and treated with ethylene (ACC),

NPA (NPA), and ethylene and NPA (NPA+ACC) in 48 hours after germination. (F and G) Number of

dividing cells at the concave and convex sides in apical hooks of seedlings untreated (F) or treated with

ethylene (ACC) 24 hours and 48 hours after germination.


95

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309.


97

SUPPORTING COMPUTATIONAL METHODS

Numerical and simulation methods

The cellular grid tissue template for the model was created with the VV (Vertex-

Vertex) programming language (Smith et al. 2003, 2006) and in the L-system-based

modeling software L-studio (Karwowski and Prusinkiewicz, 2004; Prusinkiewicz, 2004)

(http://algorithmicbotany.org/lstudio). The simulations were done by numerical

computations of coupled ODE systems, with an adaptive fifth-order Runge-Kutta method.

All figures were processed in Adobe Illustrator (CS).

Digitalized apical hook cross sections

Confocal image stacks of Arabidopsis hypocotyls (with the PIN3-GFP reporter)

were loaded into MorphoGraphX (http://www.sybit.net/software/MorphoGraphX/) for

processing. The MorphGraphX software is specialized in the extraction of curved

surfaces from 3D data. Confocal section templates can be problematic, because it is

frequently difficult to slice nicely in one single plane through all the cells.

MorphoGraphX overcomes this problem by allowing the user to define a Bezier cutting

surface that can be slightly bent or twisted interactively by moving control points. After

adjustment to obtain a good image of all the cells, the Bezier cutting surface was

converted into a triangular surface mesh and subdivided into high-resolution

(approximately 200,000 points) images. MorphoGraphX uses the VV-based (Smith et al.,

2003; Smith et al., 2006) graph rotation system software for surface mesh subdivision,

storage, and other mesh processing operations. After the surface had finely been

subdivided, a band of light from the confocal stack (approximately 1 µm on both sides)

was projected onto the surface giving a clear outline of the cells. The surface was seeded

and segmented into cells by means of a watershed technique adapted for graphical use.

After segmentation, cells were simplified to reduce the number of model elements. The

resulting cell geometry was projected onto a plane and used to simulate the computer

model with L-studio environment (Karwowski and Prusinkiewicz, 2004; Prusinkiewicz,

2004).

http://algorithmicbotany.org/lstudio

http://www.sybit.net/software/MorphoGraphX/

Chapter 3

98

Mathematical description of the apical hook model

In the computer model, each cell consists of two compartments, namely the

cytoplasm and the plasma membrane (Fig. 4A). The plasma membrane is divided into

several fragments, each associated with one side of the cell facing an adjacent cell

neighbor. Nonpolar cell-to-cell auxin transport is modeled as follows:

iijNj

jiIAAH

i

i AUXINAUXINAUXINlPVdt

dAUXIN

i

)(1

(1)

where AUXINi and AUXINj are mean auxin concentrations in the cytoplasm of the i-th

cell. Vi and li->j are the i-th cell volume and crossing areas between the i-th cell and

adjacent j-th cells, respectively. The parameter PIAAH (10 µm-1

) represents the plasma

membrane permeability for nonpolar auxin transport (Goldsmith et al., 1981). Φ is the

auxin source term (vascular tissue; green stars in Fig. 1A) and most apical cell layer

(cotyledons; Fig. 4A) and µ is the auxin sink (auxin exiting via hypocotyl boundaries;

Fig. 4A).

The PIN-dependent polar auxin transport is modeled as follows:

)11

(1

i

i

i

j

j

jNj

jiPIN

i

i

AUXIN

AUXINP

AUXIN

AUXINPlP

Vdt

dAUXIN

i

(2)

where PPIN controls the PIN-dependent auxin transport across the plasma membrane. PINi

and PINj correspond to the PIN levels in i-th and adjacent j-th cells. We assumed similar

transport rates for PIN1, PIN3, PIN4, and PIN7 by setting PPIN to its default value of 20

(µm-1

) (Goldsmith et al., 1981; Grieneisen et al., 2007; Swarup et al., 2007; Kramer et al.,

2011) only for cell sides (i->j) that correlate with the experimentally derived PIN

polarities (Ţádníková et al., 2010); otherwise, the background PIN permeability of 1 µm-1

was used. To simulate the pin mutants, parameter PPIN has been set to 1/4 of its default

value.


99

Dynamic regulation of PIN protein activity

The differential expression of PIN proteins in the cortex and epidermis has been

modeled by means of two geometric functions: (a) the angle between the cell mass center

(vc) and reference vector (vg) (concave side of the hook) (see supplementary Fig. S1A):

/)cos(gc

vva (3)

and (b) the dot product of cell mass center (vc) and reference vector (vg) (concave side of

the hook) (see supplementary Fig. S1B):

)1(5.0gc

vv (4)

These functions were negligible (Φ=0) for the expression of PIN1 that was restricted to

vascular tissues and endodermis (Fig. 6A). We found that function (4) provides the best

fit to the experimental observations.

Next, we assumed that the PIN protein activity is differentially regulated on the

concave and convex sides of the hook. To assess this issue, we developed the following

formula:

iPIN

iM

PINi PINdPINK

a

dt

dPIN

1

)1(

(5)

where PINi is the PIN level in the i-th cell. Parameters aPIN and dPIN correspond to the

basal rates of the PIN production and degradation, respectively. KM is the saturation rate

of the PIN protein level and η defines the strength of the differential PIN protein activity.

In the case of a model of a growing apical hook, equation (5) was used to calculate the

PIN protein level in the individual cells with the gravity response function Φ > 0 in the

cells corresponding to the apical hook zone (Fig. 4A). The default parameter settings for

Fig. 1 and Fig. 3 were: aPIN = 1 (s-1

); dPIN = 0.03 (s-1

); KM = 100 (µM); η = 1. The setting

was η =1 (see supplementary Fig. S2B), η =0.5 (see supplementary Fig. S2C), and η =0.1

(see supplementary Fig. S2D).

Physically based model of the apical hook

The mechanics of cell growth in the apical hook model (Fig. 4) are based on the

soft body dynamics and mass spring models (Smith, 2011). Each vertex v is connected to

its neighbor vertex u by elastic springs with a given resting length (Lv->u). The force

Chapter 3

100

acting on the point mass of a vertex v located at position pv due to springs Fv is based on

Hooke‟s law and is calculated as:

vNuvu

vu

uv

vu

xvpp

pp

L

ppkF )1(

(6)

where Nv describes the neighbors of vertex v; pu is the position of a adjacent vertex u, kx is

the stiffness of the spring per unit length and was set to 0.7. The norm symbol indicates

the Euclidean distance between the points. For a given difference from the rest length, the

magnitude of the force is reduced as the rest length of the spring increases. In addition to

the forces on a vertex due to springs, a uniform force representing the internal turgor

pressure Fp is included in the model that acts in the direction of the surface normal at each

vertex. Finally, to balance the tissue mesh integrity, we used bending springs that respond

to changes in angle of three adjacent vertices (pv, pu, and pk). A single bending spring

exerts a moment proportional to displacements of the actual angle θv->u,v-> k from a initial

rest angle θ0:

kvuv

kvuv

kvuvpppp

pppp )()(cos 1

,

(7)

)()()(

)()()(()(

0,

,

kvuvuvuv

kvuvuv

Nkukvuvbb

pppppppp

ppppppkFv

))()()(

)()()(

uvkvkvkv

uvkvkv

pppppppp

pppppp

(8)

where kb is the stiffness of the bending spring per unit length and was set to 1.

By combining the basic force components, the balanced force acting on each of the

vertices is given as:

bpvtFFFF

(9)

To calculate the displacement of each vertex i, we used the second Newton‟s Law to

update its velocity (vi) and position (pi) over time:


101

i

i

it

i

vdt

dp

vFdt

dv)(

(10)

where νi is a damping constant (0.2).

In each time step (dt = 0.01), the coupled ODE system (10) was solved by means of an

explicit Euler scheme.

To model the local, longitudinal tissue growth, we assumed that the resting length

of each linear spring that increases over time is governed by following formula:

)(vbuv

uv AUXINzgLdt

dL

(11)

where gb is a isotropic growth rate (0.001 s-1

) and )(v

AUXINz is the anisotropic

growth rate (longitudinal growth) that depends on the mean auxin concentration of two

adjacent cells that share the common spring interface. The anisotropic growth rate was

calculated by inverse mapping of the auxin concentrations to the growth degree:

vAUXINcv

b

aAUXINz

exp1)(

(12)

Other parameters were a=1 and b=0.01 (Fig. 4B-4D and 4G-4K, green line; Fig. 4E, blue

line; and Fig. 4F, red line).

Chapter 3

102

To allow for progressive growth, both the resting lengths of the linear spring and resting

angles of the bending spring were recalculated with a quasi-stationary state of balanced

forces and updated prior to the next algorithm iteration.

Zone definition in the apical hook model

To approximate a 3D representation of the hook by a simpler and computationally

inexpensive 2D model, both ends of the hypocotyl epidermis have been connected, so that

periodic boundary conditions were fulfilled. Cell association to three different zones of

the model (Fig. 4A) was obtained by calculating the relative distance of each cell from the

cotyledons (uppermost cell layer; Fig. 4A). These distances were computed via

summation of linear spring lengths anchored to the most apical cell and afterwards scaled

to given values in the 0-1 range. Next, two cut-off values for these distances were

defined; one associated with the beginning of the hypocotyl region (DHyp) and one with

the end of the cotyledon zone (Dcot). Default values for these cut-off thresholds were:

DHyp = 0.35 and Dcot =0.05 and DHyp = 0.2 and Dcot =0.05 (Fig. 4G, 4H, 4K). To define the

zone of rapid cell division, the default distance threshold Ddiv was introduced and set to

0.2 in all simulations, except Ddiv = 0.4 (Fig. 4G, 4H, 4K). The auxin concentration

threshold for rapid cell division used was set at 0.1 (Fig. 4I, 4J).


103

ADDITIONAL REFERENCES

Goldsmith, M.H.M., Goldsmith, T.H., and Martin, M.H. (1981). Mathematical-Analysis of the

Chemosmotic Polar Diffusion of Auxin through Plant-Tissues. P Natl Acad Sci-Biol 78, 976-980.

Grieneisen, V.A., Xu, J., Maree, A.F.M., Hogeweg, P., and Scheres, B. (2007). Auxin transport is

sufficient to generate a maximum and gradient guiding root growth. Nature 449, 1008-1013.

Karwowski, R., and Prusinkiewicz, P. (2004). The L-system-based plant-modeling environment L-

studion 4.0. In Proceedings of the 4th International Workshop on Functional-Structural Plant

Models, 403-405.

Kramer, E.M., Rutschow, H.L., and Mabie, S.S. (2011). AuxV: a database of auxin transport velocities.


Prusinkiewicz, P. (2004). Art and science for life: Designing and growing virtual plants with L-systems.

Acta Hortic, 15-28.

Smith, C., Prusinkiewicz, P., and Samavati, F. (2003). Local specification of surface subdivision

algorithms. Applications of Graph Transformations with Industrial Relevance 3062, 313-327.

Smith, R.S. (2011). Smith, RS. (2011) Modeling Plant Morphogenesis and Growth. In “New Trends in the

Physics and Mechanics of Biological Systems: Lecture Notes of the Les Houches Summer School:

Volume 92, July 2009, Oxford University Press, 2011.

Smith, R.S., Guyomarc'h, S., Mandel, T., Reinhardt, D., Kuhlemeier, C., and Prusinkiewicz, P. (2006). A plausible model of phyllotaxis. P Natl Acad Sci USA 103, 1301-1306.

Swarup, R., Perry, P., Hagenbeek, D., Van Der Straeten, D., Beemster, G.T., Sandberg, G., Bhalerao,

R., Ljung, K., and Bennett, M.J. (2007). Ethylene upregulates auxin biosynthesis in Arabidopsis

seedlings to enhance inhibition of root cell elongation. Plant Cell 19, 2186-2196.




The role of cytokinin in apical hook development

105

Chapter 4 The role of cytokinins in apical hook development

106

Adapted from:

Ţádníková, P., Vandenbussche, F., Van Der Straeten, D., Friml, J. and Benková E. The

role of cytokinins in apical hook development (manuscript in preparation)


E.B. and P.Ţ. conceived and designed the experiments; P.Ţ. performed all the

experiments; P.Ţ. prepared the figures; and P.Ţ. and E.B. wrote the manuscript.

Picture:

Apical hooks of wild-type Col-0 seedlings stained with propidium iodide, 48 hours after

germination

The role of cytokinins in apical hook developments

107

The role of cytokinins in apical hook development

Petra Ţádníková1,2

, Filip Vandenbussche3, Dominique Van Der Straeten

3, Jiří Friml

1,2 and

Eva Benková1,2*

1Department of Plant Systems Biology, VIB, 9052 Gent, Belgium

2Department of Plant Biotechnology and Bioinformatics, Ghent University, 9052 Gent,

Belgium

3Unit Plant Hormone Signaling & Bio-imaging, Department of Physiology, Ghent

University, 9000 Gent, Belgium

* Author for correspondence ([email protected])

Chapter 4

108

ABSTRACT

The apical hook plays an important role in early seedling development by

protecting the delicate shoot apical meristem and cotyledons during soil penetration. The

plant hormones, including auxin, ethylene, and gibberellins, are known to regulate apical

hook development. Although cytokinin has been proposed to be involved in the control of

apical hook development, its exact role remains largely unknown. By combining genetic

approaches and detailed developmental analyses, we demonstrated that cytokinin

modulates different phases of apical hook development and we showed that cytokinin

activity is coupled with ethylene at the early stages of apical hook development, but

might operate the opening phase independently of ethylene. We determined the distinct

roles of cytokinin receptors in the control of apical hook development: the ARABIDOPSIS

HISTIDINE KINASE2 (AHK2) gene functioned as a negative regulator of the formation

phase and as a positive regulator of the opening phase, whereas AHK3 and CRE1/AHK4

acted as positive regulators of the maintenance phase and as negative regulators of the

opening phase. Our data indicate that the specific function of the cytokinin receptors in

the regulation of apical hook development might result from differences in their

interaction with the ethylene pathway. AHK2 might suppress ethylene biosynthesis and

signaling, but AHK3, together with CRE1/AHK4, might have the opposite function.

Finally, we found that cytokinin altered the auxin distribution during hook development

by affecting the polar auxin transport via interference with the expression level of

particular PIN-FORMED genes.


109

INTRODUCTION

Dicotyledonous plants have evolved a strategy to protect their delicate tissues

from damage during seedling emergence through the soil; soon after germination in the

dark, they form an apical hook in the upper part of the hypocotyl to protect meristematic

primordia and cotyledons (Darwin and Darwin, 1881; Guzmán and Ecker, 1990). The

apical hook is formed by differential growth on its opposite side (Silk and Erickson,

1978; Ecker 1995; Gendreaus et al., 1997) and cell division localized mainly at the

concave hook side, close to the cotyledons (Raz and Koornneef, 2001).

To ensure proper apical hook development, the differential growth progression

has to be regulated precisely. Plant hormones have been identified to be important

endogenous regulators of cell elongation and division, resulting in differential growth.

The plant hormones auxin and ethylene are keys regulators of this developmental process

and their role in hook development has been described (Guzmán and Ecker, 1990.

Lehman et al., 1996; Vandenbussche et al., 2010; Ţádníková et al. 2010). Asymmetric

auxin accumulation on the opposite side of the hypocotyl is essential to guarantee

differential growth and apical hooks as a result (Lehman et al., 1996; Ţádníková et al.,

2010). Mutation or pharmacological treatments affecting auxin biosynthesis,

accumulation, transport, or signaling provoke defects in apical hook development or even

abolish hook formation (Boerjan et al., 1995; Lehman et al., 1996; Stowe-Evans et al.,

1998; Li et al., 2004; Tatematsu et al., 2004; Stepanova et al., 2008; Ţádníková et al.,

2010). Similarly to auxin, ethylene is an important endogenous regulator of apical hook

development. Ethylene application enhances the apical hook curvature and ethylene-

insensitive mutants, such as ethylene response1 (etr1) and ethylene-insensitive2 (ein2),

exhibit a hookless phenotype (Guzmán and Ecker, 1990). A great example of the ethylene

and auxin interaction is the formation of exaggerated apical hooks that can be caused by

expression variations of several auxin efflux carriers, the PIN-FORMED (PIN) proteins,

leading to changes in the auxin distribution (Vandenbussche et al., 2010; Ţádníková et al.,

2010). In addition, other hormones (gibberellins and brassinosteroids) contribute in hook

establishment and their interaction with auxin and ethylene has been described (Guzmán

and Ecker, 1990; Vriezen et al., 2004; Gallego-Bartolomé et al., 2011). Nevertheless, thus

far, very little is known about the role of cytokinins in the regulation of apical hook

development.

Chapter 4

110

Cytokinins had been discovered in a search for factors stimulating plant cell

division (Caplin et al., 1948; Miller et al., 1955) and, subsequently, have been shown to

be important regulators of many physiological and developmental aspects in plants,

including seed germination, leaf senescence (Riefler et al., 2006), leaf and cotyledon

expansion (Dowens and Crowell, 1998), chloroplast differentiation, mobilization of

nutrients (Mok 1994), apical dominance (Swarup et al., 2002), lateral root organogenesis

(Laplaze et al., 2007), formation and activity of the shoot apical meristem (Zhao et al,

2010), and breaking of bud dormancy and floral development (Mok 1994; Riefler et al.,

2006).

Cytokinin signaling pathways involve a two-component phosphorelay regulatory

system that is similar to the bacterial two-component system and consists of a sensor

protein and response regulator proteins (Popas et al., 1996; Lohrmann and Hater, 2002).

Cytokinin receptors belonging to the ARABIDOPSIS HISTIDINE KINASE (AHK)

family mediate the activation of the ARABIDOPSIS HISTIDINE

PHOSPHOTRANSFER (AHP) proteins that transduce the cytokinin signal to B-type

ARABIDOPSIS RESPONSE REGULATORS (ARRs) in the nucleus (Urao et al., 2000;

Kakimoto, 2003; Grefen and Harter, 2004; Bishopp et al., 2009). In Arabidopsis thaliana,

the response regulators are encoded by a multigene family that fall into two basic classes:

the A-type ARR genes, which consist uniquely of a receiver domain, and the B-type ARR

genes, which contain a transcription factor domain in addition to the receiver domain. A-

type ARR genes ensure a negative feedback loop of the cytokinin pathway, whereas the

B-type ARR genes operate as transcription factors (Bishopp et al., 2009).

An essential part of the cytokinin-related regulation of development implicates the

interaction with other hormones, especially auxin. Although cytokinin has been known to

play an important role in plant development, several studies have shown that its

interaction with auxin is crucial for the mutual control of plant developmental processes,

such as lateral root initiation, specification of vascular pattern in roots, or meristem

development (Muller and Sheen, 2008; Dello Ioio et al., 2008; Moubayidin 2009;

Bishopp et al., 2011).

Besides the interaction between cytokinin and auxin, the interaction between

cytokinin and ethylene is remarkable. Cytokinin has been reported to increase the

posttranscriptionally regulated ethylene biosynthesis (Vogel et al, 1998; Woeste et al.,

1999) by preventing the 1-aminocyclopropane-1-carboxylate synthase (ACS) enzyme,


111

involved in ethylene biosynthesis, to degrade, thus increasing the stability of ACS

proteins (Chae and Kieber, 2005).

Here we focus on the involvement of cytokinins in differential hypocotyl growth

that is important for appropriate hook development. Our results suggest that apical hook

development is controlled by cytokinin, of which the activity is tightly coupled with

ethylene in the formation and maintenance phases of apical hook development. Later, the

apical hook opening phase is controlled by cytokinin, but presumably in a manner

independent of ethylene. By combining gene expression and mutant phenotype analyses,

we observed that individual cytokinin receptors play distinct roles in apical hook

development. The AHK2 gene operates as a negative regulator of the formation phase and

as positive regulator of the opening phase. In contrast, AHK3 and CRE1/AHK4 regulate

positively and negatively the maintenance and opening phases, respectively. Furthermore,

we identified several response regulatory components of the cytokinin signaling that

might mediate downstream responses on individual receptors during apical hook

development. Analyses of ethylene levels and pathway indicated that the specific

regulatory roles of cytokinin receptors in apical hook development might result from

differences in their interaction with the ethylene pathway. AHK2 probably acted as

suppressor of the ethylene biosynthesis and signaling, whereas AHK3, together with

CRE1/AHK4, seemed to function the other way round. We demonstrate that a fully

working polar auxin transport machinery is crucial for the cytokinin impact and show

that, similarly to other developmental processes, cytokinin might trigger apical hook

development partially through the control over the components of the polar auxin

transport machinery, the PIN auxin efflux carriers.

Chapter 4

112

RESULTS

Cytokinin interferes in apical hook development

When seedlings grow in the presence of cytokinins in the dark, they display the

characteristic triple-response phenotype, typical of ethylene reaction. Triple response is

characterized by radial expansion of the hypocotyl, inhibition of hypocotyl and root

elongation, and exaggeration of apical hook curvature (Cary et al., 1995; Vogel et al.,

1998). Usually, apical hook development undergoes three phases: formation, starting

from seed germination until the hook is fully closed at 180°; maintenance, during which

the hook remains closed; and opening, when the hook progressively unfolds (Raz and

Ecker, 1999; Ţádníková et al., 2010).

To gain insight into how cytokinin controls apical hook development over time,

we analyzed real-time phenotypes of dark-grown Arabidopsis seedlings (Ţádníková et al.,

2010). Treatment of Arabidopsis seedlings with low concentrations (0.1 µM) of the

exogenous cytokinin, 6-benzylaminopurine (BAP), caused mild defects in the formation

phase. Kinetics of the formation phase in cytokinin-treated seedlings was slightly delayed

when compared with untreated seedlings, the hook was able to fully close, and 15% of the

seedlings displayed an exaggerated apical hook curvature. Furthermore, cytokinin

extended the maintenance phase by 35 hours in comparison to untreated control

seedlings, namely it lasted approximately 80 hours upon cytokinin treatment versus

approximately 25 hours in untreated control seedlings. Cytokinin also modified the

kinetics of the opening phase: the duration was doubled in cytokinin-treated seedlings in

comparison to untreated control seedlings (Fig. 1A).

Similarly, elevated endogenous levels of cytokinin through overexpression of

ISOPENTENYLTRANSFERASE 3 (IPT3), a key enzyme in the cytokinin biosynthesis

(Galihet et al., 2008), led to defects in hook development similar to those upon cytokinin

treatment and hook formation resembled that of the control. Furthermore, a substantial

increase was observed in the maintenance phase from 25 hours to approximately 54 hours

and the kinetics of the opening phase were comparable to those of untreated control

seedlings, but were faster than in cytokinin-treated seedlings (Fig. 1B).

Conversely, depletion of endogenous cytokinin levels due to overexpression of the

cytokinin degradation enzyme CYTOKININ OXIDASE 3 (CKX3) (Pernisová et al., 2008)

hampered hook development. Severe defects were observed in the formation phase, when


113

the hook was not able to fully close and reached a maximum angle of 155.27° ± 4.50°,

and the kinetics of the hook formation were delayed in comparison to control. The

maintenance phase lasted approximately 18 hours and gradually passed to the opening

phase, of which the kinetics were comparable to those of the control (Fig. 1B). In

conclusion, cytokinin might control different phases of apical hook development,

including the formation phase, during which it appeared to be important for proper hook

establishment, the maintenance and opening phases, during which it seeminlly prevented

premature hook opening.

Cytokinin effects on apical hook development are partially mediated through

ethylene

The effect of cytokinin on root and hypocotyl elongations in Arabidopsis is largely

mediated by the production of ethylene (Cary et al., 1995). Cytokinin intensifies the

apical hook curvature in dark-grown seedlings (Cary et al., 1995) as a consequence of the

cytokinin-mediated increase in the stability of ACS proteins that enhances the ethylene

biosynthesis (Vogel et al, 1998; Woeste et al., 1999; Chae and Kieber, 2005; Hansen et

al., 2009).

To examine whether cytokinin might modulate ethylene responses during apical

hook development, the ethylene response reporter EARLY BOLTING IN SHORT DAYS

fused to β-glucuronidase (EBS::GUS) was treated with exogenous cytokinin and ethylene.

Similarly to ethylene that upregulates the expression of the ethylene-responsive reporter

(Vandenbussche et al., 2010) (Fig. S1A, S1C in the supplementary material), cytokinin

upregulated the expression of EBS::GUS in hypocotyls toward the root, although more

weakly than ethylene (Fig. S1A, S1B, S1C in the supplementary material). These results

suggest that cytokinin, like ethylene; stimulates the ethylene response in hypocotyls

during apical hook development.

Our data confirm ethylene as a possible factor for cytokinin-regulated apical hook

development. To determine which phase is affected by cytokinin and to dissect whether

the cytokinin effect is partly mediated by ethylene, we combined pharmacological and

genetic approaches. Treatment with the ethylene precursor 1-aminocyclopropane-1-

carboxylate (ACC) typically provokes an exaggerated apical hook (Guzmán and Ecker,

1990; Vandenbussche et al., 2010). Upon ethylene treatment, the hook formation was

slower than that in untreated control seedlings. Approximately 50% of the seedlings had

Chapter 4

114

an amplified hook after ethylene treatment and the maintenance phase lasted

approximately 50 hours, whereafter the hook opened gradually, but with delayed kinetics.

In comparison to cytokinin, ethylene caused a slower hook formation that lasted

significantly longer, resulting in the exaggerated hook curvature, followed by a very short

maintenance phase that shifted to the opening phase. In contrast, cytokinin caused a

dramatically longer maintenance phase, followed by a slow hook opening (Fig 1A).

Next, we tested whether cytokinin acts through the ethylene biosynthesis during

apical hook development by means of the ethylene biosynthesis inhibitor 2-

aminoethoxyvinylglycine (AVG). Treatment with AVG did not affect the formation

phase when compared with the untreated control, whereas the maintenance phase

shortened by approximately 5 hours. In addition, the kinetics of the opening phase upon

the AVG treatment were unaffected. When AVG was applied simultaneously with

cytokinin, the formation phase was similar to that of the wild type; the maintenance phase

lasted approximately 20 hours, followed by the opening phase. When the hook opened to

an angle of 130.2° ± 10.6°, the process was halted and followed by an arrest phase, during

which the angle of the hook remained at 130° ± 11.7°. This phase lasted for 84 hours,

whereafter the hook gradually opened and the opening kinetics were comparable with

those of hooks treated with ethylene or cytokinin (Fig. 1A). In summary, AVG reversed

the effect of cytokinin on the maintenance phase, but not on the opening phase.

Furthermore, we tested the impact of cytokinin on the ethylene-insensitive mutant

ethylene response 1-3 (etr1-3) (Zhao et al., 2002). The etr1-3 mutant displays no defects

in the formation and opening phases, whereas the maintenance phase is reduced (Fig. 1C)

(Vandenbussche et al., 2010), as in the AVG-treated control seedlings. The cytokinin-

treated etr1-3 mutant had no defective formation and maintenance phases, when

compared with untreated mutants, whereas the opening phase decelerated suddenly the

opening with an halted phase. A very similar phenotype was caused by cytokinin with a

simultaneous application of AVG (Fig. 1C). In conclusion, our data demonstrate that the

effect of cytokinin on exaggerated hook formation and prolongation of the maintenance

phase was rescued by AVG, hinting at a coupled role for ethylene and cytokinin at the

early stages of apical hook development, whereas cytokinin might govern the opening

phase, independently of ethylene.


115

020406080

100120140160180200220240260

0 40 80 120 160 200 240 280

angl

es [

°]


Col_MSCol_BAPetr1-3_MSetr1-3_BAP

020406080

100120140160180200220240260

0 20 40 60 80 100 120 140 160

an

gle

s [°

]


Col_MSCol_BAP35S::IPT3_MS35S::CKX3_MS

A

B C

020406080

100120140160180200220240

0 40 80 120 160 200 240 280

angl

es [

°]


Col_MSCol_BAPCol_BAP+AVGCol_AVGCol_ACC

Figure 1. Main effects of cytokinin on apical hook development in Arabidopsis

(A) Kinetics of apical hook development in Col-0 seedlings untreated (MS) or upon cytokinin (BAP),

BAP+AVG, and ethylene (ACC) treatment. (B) Kinetics of apical hook development in Col-0, IPT3, and

CKX3 under the control of the cauliflower mosaic virus (35S) promoter (35S::IPT3 and 35S::CKX3). (C)

Kinetics of apical hook development in untreated (MS) or cytokinin (BAP)-treated Col-0, ein2-1, and etr1-3

seedlings. MS, Murashige and Skoog medium only; Col, control Columbia seedlings. Error bars represent

s.e.m.

Cytokinin receptors play a specific role in the regulation of hook development

As our analysis showed that cytokinin affects apical hook development, we

subsequently focused on identifying key components of the cytokinin signaling pathway.

In Arabidopsis, the cytokinin signal is perceived by three sensor histidine kinases, AHK2,

AHK3, and CRE1/AHK4 (Inoue et al., 2001; Suzuki et al., 2001; Ueguchi et al., 2001;

Yamada et al., 2001). The expression of the cytokinin receptors was investigated to

determine which of them are involved in the regulation of hook development. No

AHK2::GUS expression was detected in the hook zone during the formation and

maintenance phases (Fig. 2A). AHK2 was expressed only at later stages, when the hook

switched to the opening phase, during which it was relatively strong (Fig. 2A). The

expression of the AHK3 receptor was strong during hook formation, weak in the

maintenance phase, and absent during the opening phase (Fig. 2A, 2B). The CRE1/AHK4

gene was expressed in the central cylinder, without visible changes in strength during the

Chapter 4

116

progress of apical hook development (Fig. 2C). These observations suggest that AHK3

and CRE1/AHK4 might act at the start of the hook establishment, whereas they are all

active during the maintenance phase. In the opening phase, the expression pattern

indicated the predominant contribution of AHK2 and CRE1/AHK4.

Furthermore, the detailed phenotypic characterization of the cytokinin receptor

loss-of-function mutants and their double mutant combinations was done to get insights

into the role of the cytokinin perception in hook development. The hook formation

kinetics of the ahk2-2 mutant were faster than those of its control (Fig. 2A). The apical

hook of ahk2-2 was exaggerated in 52% of the seedlings, whereas no control seedlings

displayed this specific phenotype. The maintenance phase of ahk2-2 lasted approximately

36 hours, in contrast to approximately 25 hours in the wild type, and the opening phase

was delayed in comparison to the control (Fig. 2D). A different phenotype was observed

in the cre1-12/ahk4 and ahk3-3 single mutants that did not show exaggerated hooks. The

kinetics of their formation phases were comparable to those of the controls, but their

maintenance phase was shorter, lasting approximately 15 hours versus approximately 25

hours in their control. The kinetics of the hook opening was faster than those of the

control, especially in ahk4/cre1-12 mutant. To investigate the genetic interaction between

particular cytokinin receptors, we analyzed their double mutants. The formation phase of

the double mutants ahk2-2xahk3-3 and cre1-12/ahk4xahk2-2 was delayed when

compared to their single mutants. Furthermore, the phenotype of the double mutants with

ahk2-2 resembled that of ahk2-2, namely with exaggerated hook and maintenance phase

lasting approximately 36 hours (Fig. 2E, 2F). Thus, lack of AHK3 or CRE1/AHK4 did not

revert the ahk2-2 phenotype on hook maintenance and opening. The phenotype of the

double mutant cre1-12/ahk4xahk3-3 differed from that of single mutants in (i) the slower

hook formation, (ii) the slightly longer maintenance phase, lasting approximately

19 hours, (iii) the deficient hook closure, reaching only 170.43° ± 4.6° in contrast to 180°

in the control, and (iv) the faster hook opening (Fig. 2G).

Taken together, our analyses unveil AHK2 as a negative regulator of the

formation-to-maintenance phase transition, although it positively regulates the

maintenance-to-opening phase transition and the opening phase of the hook. Further

analyses also revealed that the activities of the cytokinin receptors AHK3 and

CRE1/AHK4 were redundant during the hook formation and early maintenance phases,

operating as positive regulators of the maintenance phase and as negative regulators of


117

the transition between the maintenance and opening phases. Moreover, CRE1/AHK4

might also act as a negative regulator of the opening phase.

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Figure 2. Differential expression of particular cytokinin receptors and apical hook kinetics of

cytokinin receptor mutants

(A) Expression pattern of AHK2::GUS in untreated seedlings (MS) in the formation, maintenance, and

opening phases. (B) Expression pattern of AHK3::GUS in untreated seedlings (MS) in the formation,

maintenance, and opening phases. (C) Expression pattern of CRE1::GUS in untreated seedlings (MS) in the

formation, maintenance, and opening phases. (D) Kinetics of apical hook development in Col-8, ahk2-2,

ahk3-3, and cre1-12 single mutants. (E) Kinetics of apical hook development in Col-8, ahk2-2, ahk3-3

single mutants and ahk2-2xahk3-3 double mutant. (F) Kinetics of apical hook development in Col-8, ahk2-

2, cre1-12 single mutants and cre1-12xahk2-2 double mutant. (G) Kinetics of apical hook development in

Col-8, ahk3-3 and cre1-12 single mutants and cre1-12xahk3-3 double mutant. MS, Murashige and Skoog

medium only; Col, control Columbia seedlings. Error bars represent s.e.m.

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AHK2 represses ethylene biosynthesis, but AHK3 does not

Detailed phenotype analysis of hook development of cytokinin receptor loss-of-

function mutants revealed that the phenotype of the ahk2-2 mutant is reminiscent of that

of enhanced ethylene activity. To further explore whether ethylene is truly involved in the

ahk2-2 phenotype, additional experiments were carried out. Initially, the rescue of the

ahk2-2 phenotype by the ethylene biosynthesis inhibitor AVG was tested. AVG was able

to rescue the phenotype of the ahk2-2 mutant, because, upon the AVG treatment, the

exaggerated hook was suppressed, the formation phase was delayed, the maintenance

phase was shorter, and the kinetics of the opening phase were faster than for the ahk2-2

mutant (Fig. 3A). Thus, the AVG treatment indicated that the phenotype of the ahk2-2

mutant might be mediated by an increase in ethylene biosynthesis or ethylene levels that

cause the typical triple response.

To evaluate whether the cytokinin receptors feed back on the ethylene

biosynthesis, signaling, metabolism, or response, we utilized the quantitative reverse-

transcription (RT)-PCR for expression analyses. The ahk2-2 mutant expression analysis

revealed that the expression was enhanced of most of the genes involved in ethylene

biosynthesis (reviewed in Wang et al., 2002), namely ACS1, ACS2, ACS4, ACS5, ACS6,

ACS7, ACS8, ACS9, and ACC OXIDASE2 (ACO2), of which the strongest expression was

approximately 4-fold and 6-fold higher for ACS9 and ACS7 than that of wild type,

respectively. In addition, genes involved in the ethylene signalling cascade (reviewed in

Wang et al., 2002) were also upregulated (ETR1, ERT2, ERS1, ERS2, ETHYLENE

OVERPRODUCER1 [ETO1], CONSTITUTIVE TRIPLE RESPONSE1 [CTR1], EIN2,

EIN3, EIN4, and EIN5) as well as the ethylene-responsive gene HOOKLESS (HLS).

These data indicate an enhanced ethylene biosynthesis and signaling that, consequently,

results in the triple-response phenotype of ahk2-2.

The ahk3-3 mutant responded differently, with a significant downregulation of

most genes involved in ethylene biosynthesis (ACS1, ACS4, ACS9, and ACS10) and

upregulation by only approximately 2-fold of ACS2 and ACS6. The ethylene-responsive

gene HLS (Lehman et al., 1996) was slightly upregulated, whereas the expression of the

ethylene signaling and metabolism genes did change dramatically.

Finally, the effects of the cre1-12/ahk4 mutant were milder than those of the other

ethylene receptor mutants: the ethylene biosynthesis genes ACS2 and ACS6 were slightly

upregulated and ACS1, ACS4, ACS8, ACS10, ACO2, and ACO4 were somewhat


119

downregulated; the expression of HLS was upregulated, but to a lesser extent than in the

ahk2-2 mutant; and no dramatic changes were observed in the other genes involved in

ethylene signaling and metabolism (Fig. 4A).

Thereafter, to confirm the regulatory role of the cytokinin receptor mutants in

ethylene biosynthesis during hook development, we measured the ethylene production in

real-time (Bijnen et al., 1996). The ethylene production was significantly higher in the

ahk2-2 mutant than that in the control (Fig. 4B), increased from 48 hours after

germination, and remained until 120 hours after germination. In the ahk3-3 and cre1-

12/ahk4 mutants, no remarkable changes in the ethylene production were visible, except

for a slight increase only 76 hours after germination in the latter mutant (Fig. 4B).

Finally, the expression of ethylene receptor genes was investigated with the

transgenic lines ETHYLENE RESPONSE SENSOR1 (ERS1)::GUS and ETR1::GUS to

have an overview of their role in the control of hook development and to compare their

spatial expression pattern with that of the cytokinin receptors. Both genes were

asymmetrically expressed during hook formation in the hook zone and central cylinder

(Fig. S1A, S1B in the supplementary material). The expression pattern overlapped with

CRE1::GUS and partially in AHK2::GUS and AHK3::GUS seedlings (Fig. 2A, 2B, 2C).

Our findings suggest that ethylene and cytokinin receptors might act jointly to control

hook development.

Together, our observations further support specific roles for the cytokinin

receptors in the interaction with the ethylene pathway as part of the crosstalk between

cytokinin and ethylene during apical hook development. AHK2 might act as suppressor of

ethylene biosynthesis and signaling, whereas AHK3 would positively regulate these

processes. Based on the expression data and ethylene measurements, the function of

CRE1/AHK4 in the interaction with ethylene could not be assigned with certainty, despite

the similarity to AHK3.

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Figure 3. Rescue of the ahk2-2 phenotype by the ethylene biosynthesis inhibitor AVG

(A) Kinetics of apical hook development in untreated Col-0, etr1-3 (MS), and AVG-treated seedlings. MS,

Murashige and Skoog medium only; Col, control Columbia seedlings. Error bars represent s.e.m.

Figure 4. Quantitative RT-PCR analysis and ethylene measurements in cytokinin receptor mutants

(A) Expression analysis of ethylene biosynthesis, ethylene signaling, and ethylene-responsive genes in Col-

8 and ahk2-2, ahk3-3, and cre1-12 single-mutant seedlings 48 hours after germination. (B) Ethylene

measurements in 1-day-old to 6-day-old Col-8 and ahk2-2, ahk3-3, cre1-12 single-mutant seedlings. Col,

control Columbia seedlings. Error bars represent s.e.m. (*p<0.05, n=3)


121

Cytokinin receptor mutants differ in sensitivity to cytokinin and ethylene during

hook development

The ahk2-2 mutant showed the exaggerated hook phenotype and a delayed

transition from the maintenance to the opening phases, as well as enhanced ethylene

pathways, including biosynthesis and signaling. The opposite phenotype was found in

ahk3-3 and cre1-12/ahk4, with a faster transition from the maintenance to the opening

phases, whereas the reduction in the ethylene pathway was less dramatic in cre1-12/ahk4

than in ahk3-3. Subsequently, we analyzed the sensitivity of individual cytokinin receptor

mutants to cytokinin and ethylene to evaluate their impact on apical hook establishment

in a single mutant background and to know whether the AHK receptors are necessary for

the cytokinin/ethylene-mediated or the cytokinin-specific phenotypes. Furthermore, the

comparison of the cytokinin receptor mutant sensitivity to the relevant hormones would

allow the determination of the potentially coupled function and contribution of cytokinin

and ethylene to hook development.

The sensitivity to both cytokinin and ethylene of the apical hook development in

the ahk2-2 mutant was comparable to that of its control (Fig. S2A, S2D in the

supplementary material). Conversely, the sensitivity to cytokinin of ahk3-3 was

comparable to that of the control in the formation phase, but was reduced in the

maintenance and opening phases of the hook. A reduced sensitivity to cytokinin was

demonstrated by a shorter maintenance phase and faster kinetics of the opening phase

thant those of the control upon a cytokinin treatment (Fig. S2B, S2E in the supplementary

material). The sensititivy to ethylene of ahk3-3 was similar to that of its control in the

formation and maintenance phases, but enhanced the delayed opening (Fig. S2B, S2E in

the supplementary material). The loss-of-function mutant cre1-12/ahk4 showed an

oversensitivity to both cytokinin and ethylene. Upon treatment with cytokinin, the

formation phase was faster, the maintenance phase was prolonged, and the opening phase

was delayed when compared to cytokinin-treated control. The ethylene treatment did not

affect the formation phase, whereas the maintenance and opening phases were delayed in

comparison to the ethylene-treated control (Fig. S2C, S2F in the supplementary material).

In conclusion, our data show that the ahk2-2 mutant is as sensitive to cytokinin

and ethylene as the controls. The sensitivity to cytokinin of the ahk3-3 mutant in the

formation and maintenance phases was similar to that of the control, but reduced during

the transition to the opening phase. The cre1-12/ahk4 mutant displayed an oversensitivity

Chapter 4

122

to cytokinin in all phases of hook development and to ethylene in the maintenance and

opening phases. Taken together, the AHK3 expression is required for a cytokinin-

mediated effect and AHK3 and CRE1/AHK4 are important for balancing the impact of

ethylene and of cytokinin and ethylene, respectively.

Downstream components of the cytokinin signaling pathway involved in hook

development

To understand how the cytokinin signaling cascade works as a complex during

apical hook development, we further focused on the investigation of downstream

components. In an attempt to find phenotypes resembling those of particular cytokinin

receptor mutants, the phenotype of several ARR loss-of-function mutants were analyzed,

including the B-type ARR transcriptional activators and A-type ARR negative regulators

of cytokinin signaling (Hutchison et al., 2002).

Initially, the B-type ARR response regulators were studied. No changes in the

apical hook of arr2 and arr13 were observed in the formation phase, but defects in the

maintenance phase, which was shorter in both mutants. In addition, the opening phase

was faster than that of the control in the arr2 and arr13 mutants (Fig. 5A). Conversely,

the single mutants arr1, arr10, arr11, arr12, and arr14 did not display any hook

phenotype. B-type ARR genes are known to act redundantly (Ishida et al., 2008).

Therefore, some of the available double and triple mutants were used to investigate

whether their involvement in hook establishment is masked by functional redundancy.

The most severe defect was found in the triple mutant arr1-3xarr10-5xarr12-1, in which

the hypocotyl was unable to form a proper hook that never reached the fully closed stage

and had severe problems to remain bent. Similarly, the double mutants arr1-3xarr12-1,

arr1-2xarr11-1, and arr1-2xarr11-2 displayed defects in hook development, mainly

exhibiting delayed formation phases, followed by short maintenance phases and faster

opening, but without dramatic differences in the kinetics from the controls (Fig. 5B).

These observations suggest that B-type ARR genes play essential, but redundant ,roles in

hook establishment during the cytokinin transduction pathway. Moreover, the phenotype

of the arr2 and arr13 single mutants and the arr1-3xarr12-1, arr1-2xarr11-1, arr1-

2xarr11-2 and arr1-3xarr10-5xarr12-1 multiple mutants resembled obviously that of the

cytokinin receptor mutants ahk3 and cre1-12/ahk4, suggesting a role for ARR1, ARR2,


123

ARR10, ARR11, ARR12, and ARR13 as downstream components of the cytokinin

signaling pathway (Fig. 5B, 5C).

Mutants of the A-type ARR negative regulators displayed two different

phenotypes: (i) no dramatic changes in the formation phase and a shorter maintenance

phase, followed by a faster opening as in the case of the arr3 and arr6 mutants; and (ii)

the opposite phenotype, characterized by no dramatic defects in the formation phase, but

a longer maintenance phase, followed by a slower opening as detected in the arr9 and

arr16 mutants (Fig. 5D). The multiple mutants arr3xarr4xarr5xarr6 and

arr3xarr4xarr5xarr6xarr8xarr9 exhibited no defective formation phase and a prolonged

maintenance phase, followed by slower opening, hinting at a redundant and predominant

role of the negative factors responsible for maintaining hook at the closed stage (Fig 5E,

5F). Altogether, these findings suggest that ARR1, ARR2, ARR10, ARR11, ARR12, and

ARR13 function as positive downstream regulatory elements and that ARR9 and ARR16

might provide a negative feedback of the cytokinin receptors AHK3 and CRE1-12/AHK4.

Furthermore, ARR3 and ARR6 were proposed as positive downstream regulatory elements

of the AHK2-mediated branch of the cytokinin pathway.

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A

B E

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arr - type B phenotype

arr1 no

arr2 shorter maintenance

arr10 no

arr11 no

arr12 no

arr13 shorter maintenance

arr14 no

arr1-3xarr12-1do not close, shorter

maintenace

arr1-2xarr11-1 shorter maintenance

arr1-3xarr11-2 shorter maintenance

arr1-3xarr10-5xarr12-1do not close, strong

deficient

arr - type A phenotype arr3 shorter maintenancearr4 no

arr5 no

arr6 weak, not closed

arr7 no

arr8 no

arr9 longer maintenance

arr15 no


arr3xarr4 no

arr8xarr9 no

arr3xarr4xarr5xarr6 longer maintenancearr3xarr4xarr5xarr6xarr8x


C F

Figure 5. Involvement of apical hook development by downstream components of the cytokinin

signaling pathway

(A) Kinetics of apical hook development in untreated Col-0, arr2, and arr13 seedlings. (B) Kinetics of

apical hook development in untreated Col-0 and the multiple mutants arr1-3xarr12-1, arr1-2xarr11-1,

arr1-2xarr11-2, and arr1-3xarr10-5xarr12-1. (C) Overview of all performed kinetics of apical hook

development in B-type ARR genes. (D) Kinetics of apical hook development in untreated Col-0, arr3, arr6,

arr9, and arr16 seedlings. (E) Kinetics of apical hook development in untreated Col-0, and multiple

mutants arr3xarr4xarr5,arr6 and arr3xarr4xarr5,arr6,arr8,arr9. (F) Overview of all performed kinetics of

apical hook development in A-type ARR genes. Col, control Columbia seedlings. Error bars represent s.e.m.

(*p<0.05, n=3)


125

Interaction of cytokinin and auxin in apical hook regulation

The asymmetric distribution and response of auxin are crucial for proper hook

development. Inhibition of the auxin transport by treatment with N-1-naphthylphthalamic

acid (NPA) as well as mutations affecting the auxin transport or response result in

dramatic defects in apical hook development (Schwark and Schierle, 1992; Boerjan et al.,

1995; Lehman et al., 1996; Vandenbussche et al., 2010; Ţádníková et al., 2010). To

investigate and specify the participation of cytokinin on the asymmetric auxin distribution

during apical hook formation, the expression of the auxin response reporters DR5::GUS

and DR5-GFP was monitored. During hook formation, the DR5 expression pattern did

not significantly differ in seedlings treated with cytokinin or ethylene from untreated

seedlings (Fig. 7I, 7J, 7K). The differences were most apparent in the maintenance phase

when the hook reaches its maximal curvature. In apical hooks treated with cytokinin, the

auxin response increased upon cytokinin treatment (Fig. 7J). A similar effect was

observed in ethylene-treated seedlings (Fig. 7I). Quantification of the DR5-GFP signal in

the middle cell at the concave side of seedlings 36 hours after germination did not reveal

any significant differences in the strength of the GFP signal between nontreated seedlings

and seedlings treated with cytokinin or ethylene (Fig. S5A, S5E, S5I in the supplementary

material). Interestingly, 86 hours after germination, the signal gradually disappeared in

ethylene-treated and control seedlings due to the opening of their hooks, whereas in

cytokinin-treated seedlings the signal remained present, apparently due to the

considerably longer maintenance phase (Fig. 7I, 7J, 7K).

Furthermore, the impact of the cytokinin treatment on the membrane-localized

PIN-GFP was examined and compared with the ethylene treatment to understand how

cytokinin and ethylene affect particular PIN proteins that are involved in apical hook

development (Ţádníková et al., 2010). The GFP signal was measured on cortical

transversal membranes because they contribute to the auxin redistribution (for details, see

Ţádníková et al., 2010). The GFP signal was quantified at the concave and convex sides

in the PIN3-GFP, PIN4-GFP, and PIN7-GFP seedlings 36 hours after germination; in the

PIN1-GFP seedlings, the GFP signal was measured 72 hours after germination on

transversal membranes of the epidermal cells, because PIN1-GFP is expressed only in

epidermis of opening hooks.

The PIN3-GFP signal did not change significantly upon cytokinin or ethylene

treatments (Fig. S3D, S3E, S3F in the supplementary material). The PIN4-GFP signal on

Chapter 4

126

transversal membranes of cortical cells was only slightly (7%) reduced at the concave

side of the hook upon the cytokinin treatment, whereas the ethylene treatment strongly

decreased (40%) the signal intensity at convex side and slightly (7%) at the concave side

when compared to untreated controls (Fig. S3G, S3H, S3I in the supplementary material).

No differences in the PIN7-GFP expression were observed after cytokinin treatment, in

contrast to the strong upregulation by ethylene of approximately 55% on both sides of the

hook (Fig. S3J, S3K, S3L in the supplementary material). After cytokinin application, the

PIN1-GFP signal was significantly reduced at the membranes, but simultaneously,

enhanced in the intracellular compartments, presumably the vacuoles (Fig. S3B in the

supplementary material). In contrast, upon ethylene treatment, the PIN1-GFP membrane

signal was reduced, but without increase in the intracellular signal (data not shown).

Quantification of the signal in epidermal membranes revealed a dramatic reduction by

44% in the PIN1-GFP signal upon cytokinin treatment in comparison to the untreated

wild type (Fig. S3A, S3B, S3C in the supplementary material). A similar effect was

observed upon ethylene treatment whereafter the PIN1-GFP signal was also reduced by

44% (Fig. S3C in the supplementary material).

Next, the effect of cytokinin on the auxin distribution in pin loss-of-function

mutants was inspected with DR5::DR5-GFP in mutant backgrounds. The GFP signal in

the middle cell at the concave side of the hook was quantified in DR5-GFP seedlings 36

hours after germination. Cytokinin or ethylene treatments did not cause any significant

difference in the strength of the signal in DR5-GFP seedlings in comparison to the

untreated controls. The signal in the untreated pin3-4 mutant was 43% lower than that of

the wild type (Fig. S5A, S5B, S5I in the supplementary material). This decrease was

abolished by exogenous cytokinin. In contrast, the ethylene treatment additionally

reduced the DR5 signal when compared to the untreated pin3 mutant (Fig. S5E, S5F, S5I

in the supplementary material). The DR5 signal intensity did not differ between the pin4-

3 mutants and the wild type (Fig. S5A, S5C, S5I in the supplementary material) and a

30% reduction was observed upon both cytokinin and ethylene treatments (Fig. S5C,

S5G, S5I in the supplementary material). In the pin7-1 mutant, the DR5 signal intensity

was the same as that of its control in untreated and cytokinin-treated apical hooks. In

contrast, ethylene application led to a 47% reduced DR5 signal in comparison to the

untreated mutant or control (Fig. S5A, S5D, S5H, S5I in the supplementary material).

Taken together, our data suggest that cytokinin and ethylene play a major role in

modulating the asymmetric auxin distribution by affecting the PIN expression. Cytokinin-


127

dependent changes in gene expression of individual PIN genes prompted us to test the

sensitivity to cytokinin of mutants lacking the expression of particular PIN genes. Upon

treatment with cytokinin, the pin3-4 mutant displayed a mildly reduced sensitivity to

cytokinin; the formation phase occurred later, the maintenance phase disappeared, and the

hook opened gradually when compared to its cytokinin-treated control (Fig. S4B in the

supplementary material). Surprisingly, the pin4-3 mutant exhibited a dramatically

reduced sensitivity to cytokinin with severe defects in all phases of apical hook

development in cytokinin-treated mutant seedlings (Fig. S4C in the supplementary

material). In the pin7-1 mutant, the sensitivity to cytokinin was moderately modified, the

hook formation was later, the maintenance phase was shorter, and the opening phase was

faster than those of the cytokinin-treated control (Fig. S4D in the supplementary

material). A slightly decreased sensitivity to cytokinin was observed in the pin1-3 mutant

and occurred in all phases of apical hook development (Fig. S4A in the supplementary

material).

Altogether, our data demonstrate that PIN3, PIN4, and PIN7 are particularly

important for the cytokinin control of the maintenance phase. When one of them is

absent, the cytokinin effect, represented by a prolonged maintenance phase, disappears.

Furthermore, because the pin1-3 mutant is less sensitive to cytokinin, especially in the

late maintenance phase, our data confirm the contribution of PIN1 to the auxin flux away

from the concave side of the hook at the end of the maintenance phase.

Chapter 4

128

AHK3

AHPs

Type-B ARRs

1,2,10,11,12,13

AHK2 CRE1

Ub

Ub

Ub

AHPs AHPs

Type-B ARRs

Type-A ARRs

3,6

Type-A ARRs 9,16

membrane

cytokinin

proteasome 26S

ACS expressionACS

Type-B ARRs

1,2,10,11,12,13

ACS

Figure 6. Model for a possible regulation by cytokinin of the ethylene pathway in apical hook

development

Cytokinin was reported to stabilize the ACS proteins by preventing their degradation (black pathway); this

effect might be mediated by the cytokinin receptors AHK3 and CRE1/AHK4 (grey pathways, Hansen et al.,

2009). Cytokinin binds to AHK3 and presumably also to the CRE1/AHK4 receptors that autophosphorylate

and, subsequently, transmit a phosphoryl group to the AHP genes. The AHP genes phosphorylate the B-type

or A-type ARR genes. The particular type B prevents the degradation of ACS proteins (grey pathways,

Hansen et al., 2009) and their other possible function, they might increase the ethylene biosynthesis via the

promoting expression of ACS (orange pathways). When cytokinin binds to AHK2, the signal is transduced

in a similar via other mentioned cytokinin receptors, but presumably involving different ARR regulators.

Downstream components of the AHK2 B-type ARR possibly prevent the ACS expression (red pathways).

The A-type ARR proteins negatively regulate the cytokinin signaling and ethylene. The blocking arrow

denotes negative regulation.


129

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A B

E

C D

ahk3-3xDR5-GFP cre1-12xDR5-GFP

ahk2-2xDR5-GFP

DR5-GFP

DR5-GFPF G

H *

I DR5::GUS MS

J DR5::GUS BAP


K DR5::GUS ACC

Figure 7. Involvement of cytokinin on the asymmetric auxin distribution during hook development

(A) Expression of DR5-GFP in untreated seedlings 36 hours after germination. (B) Expression of ahk2-

2xDR5-GFP in untreated seedlings 36 hours after germination. (C) Expression of ahk3-3xDR5-GFP in

seedlings 36 hours after germination. (D) Expression of cre1-12xDR5-GFP in seedlings 36 hours after

germination. (E) Quantification of the relative fluorescence of DR5-GFP in one cell in the middle of the

concave side of the hook (n = 10) of untreated seedlings 36 hours after germination. (F) Transversal section

in the middle of the hook in DR5-GFP untreated seedlings 36 hours after germination. (G) Transversal

section in the middle of the hook in ahk2-2xDR5-GFP untreated seedlings 36 hours after germination. (H)

Quantification from transversal sections (n = 10) of the cells with the GFP signal. (I) Expression of

DR5::GUS in untreated seedlings (MS) during the formation, maintenance, and opening phases. (J)

Expression of DR5::GUS upon cytokinin treatment (BAP) during the formation, maintenance, and opening

phases. (K) Expression of DR5::GUS upon ethylene treatment (ACC) during the formation, maintenance,

and opening phases. Error bars represent s.e.m. (*p<0.05, n=3)

Chapter 4

130

Figure 8. Expression analysis of PIN proteins in cytokinin receptor mutants

(A) Expression of PIN3-GFP in untreated seedlings 36 hours after germination. (B) Expression of ahk2-

2xPIN3-GFP in untreated seedlings 36 hours after germination. (C) Expression of ahk3-3xPIN3-GFP in

untreated seedlings 36 hours after germination. (D) Expression of cre1-12xPIN3-GFP in untreated

seedlings 36 hours after germination. (E) Quantification of the relative fluorescence of PIN3-GFP on

transversal membranes in cortical cells at the convex and concave sides of the hook in untreated seedlings

36 hours after germination (n = 10). (F) Expression of PIN1-GFP in untreated seedlings36 hours after

germination. (G) Expression of ahk2-2xPIN1-GFP in in untreated seedlings36 hours after germination. (H)

Expression of ahk3-3xPIN1-GFP in in untreated seedlings 36 hours after germination. (I) Expression of

cre1-12xPIN1-GFP in untreated seedlings 36hours after germination. (J) Quantification of the relative

fluorescence of PIN1-GFP on transversal membranes in epidermal cells at the concave side of the hook (n

= 10) in untreated seedlings 36 hours after germination . Error bars represent s.e.m. (*p<0.05, n=3)


131

Cytokinin receptor activity affects auxin response and transport

The asymmetric accumulation of the plant hormone auxin is indispensable for

appropriate hook establishment (Lehman et al., 1996; Ţádníková et al., 2010).). To

investigate the mutual influences of auxin and cytokinin and of cytokinin and ethylene,

we studied the impact of the cytokinin receptor activity on the auxin response and

transport.

Initially, the DR5::DR5-GFP auxin reporter (Benková et al., 2003) was crossed

into the ahk2-2, ahk3-3 and cre1-12/ahk4 mutant backgrounds. Analysis of DR5-GFP in

ahk2-2 mutant seedlings 36 hours after germination revealed widened auxin maxima and

extension of the DR5 signal to the convex side (Fig. 7A, 7B). Enlargement of the auxin

maxima in the epidermis toward concave side was clearly visible on the transverse

sections of closed hooks in the ahk2-2 mutant (Fig. 7F, 7G). By scoring the epidermal

cells with a signal in seedlings 36 hours after germination, we found that the number of

DR5-positive cells was 20% higher in the ahk2-2 mutants than that of the control (Fig.

7H). This DR5 expression pattern was reminiscent of that observed in ethylene-treated

seedlings, implying that the defective apical hook development in the ahk2-2 mutant

might be the consequence of the enhanced ethylene production (P. Ţádníková,

unpublished data). Quantification of the DR5-GFP signal in the middle cell at the

concave side of the hook did not reveal any significant difference in the signal intensity

between the ahk2-2 mutant and control seedlings (Fig. 7E). Likewise, the DR5 signal

distribution and intensity did not differ significantly in ahk3-3 mutant seedlings 36 hours

after germination in comparison to the control (Fig. 7A, 7C, 7E). In contrast, in cre1-

12/ahk4 mutant seedlings 36 hours after germination, the DR5-GFP signal was

significantly lower and the pattern less homogenous and patchy than in the control (Fig

7D). Quantification of the DR5 signal confirmed the reduction in the signal intensity by

30% compared to the control (Fig 7E). Our observations on the DR5 expression pattern in

cytokinin receptor mutants are overall in agreement with their respective apical hook

phenotypes. The ahk2-2 mutant showed an ethylene-related hook phenotype and its DR5

expression pattern corresponded to that of ethylene-treated seedlings. The low DR5

expression in the hook of cre1-12/ahk4 correlated with defects in apical hook

development. No significant changes in the DR5 expression were found in the ahk3-3

mutant.

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To examine the contribution of individual cytokinin receptors in the regulation of

the polar auxin transport, we analyzed the expression of the key auxin efflux carriers,

previously shown to be involved in control of apical hook development. PIN3::PIN3-

GFP and PIN1::PIN1-GFP (Benková et al., 2003; Ţádníková et al., 2010) were crossed

into the ahk2-2, ahk3-3 and cre1-12/ahk4 mutant backgrounds and the signal intensity

was measured on transversal membranes of cortical cells at both the convex and concave

sides of the apical hook in seedlings 36 hours after germination. Expression analysis of

PIN3::PIN3-GFP and quantification of the signal did not reveal any significant difference

between the ahk2-2 mutant and the control (Fig. 8A, 8B, 8E). In the ahk3-3 mutant, the

PIN3-GFP membrane signal had increased slightly, by 16% on the convex and by 15%

on the concave side of the apical hook (Fig. 8C, 8E). An opposite effect was detected in

the cre1-12/ahk4 mutant, in which the PIN3-GFP signal was downregulated by 15% and

8% on the convex and concave sides of the hook, respectively (Fig. 8D, 8E). Quantitative

analysis of PIN1::PIN1-GFP on transversal membranes of epidermal cells did not

disclose any dramatic difference in the PIN1-GFP signal in the ahk2-2 and ahk3-3

mutants and a mild (14%) increase of the PIN1-GFP signal at the concave side of the

hook in the cre1-1/ahk4 mutant, hinting at a possible role of PIN1 because the

upregulated PIN1-GFP accelerates auxin depletion and faster opening of the hook. These

results suggest distinct regulatory mechanisms among different cytokinin receptors and

their impact on the expression of individual auxin efflux carriers.


133

DISCUSSION

Involvement of cytokinin signaling in apical hook development

The establishment of the apical hook would not be possible without differential

growth that is driven by differences in cell elongation and division on opposite sides of

the hypocotyl (Raz and Koornneef, 2001). Differential growth is controlled mainly by the

asymmetric auxin distribution (Friml et al., 2002, Abas et al., 2006). The permanent

activity of auxin is required during all phases of the process to ensure proper hook

development (Lehman et al., 1996; Wu et al., 2010; Ţádníková et al., 2010). Another

plant hormone, ethylene, also plays a fundamental role in hook development, particularly

in the maintenance and opening phases (Raz and Ecker, 1999), whereas its function in the

formation phase is insignificant (Knee et al., 2000; Vandenbussche et al., 2010).

However, the role of cytokinin on apical hook development is still poorly understood.

Recent studies focused mainly on the root of Arabidopsis revealed that cytokinin has an

impact on the auxin distribution, mediated by the auxin efflux carriers belonging to the

PIN family that are under the transcriptional control of the auxin signaling pathway

(Dello Ioio et al., 2008; Růţička et al., 2009).

The combination of pharmacological and genetic approaches implied that

cytokinin functions specifically at the different stages of hook development and also

interplays with other plant hormones. Severe defects in apical hook development due to

depletion of endogenous cytokinin levels by overexpression of its degrading enzyme

CKX3 hints at an important role for cytokinin in the formation and maintenance phases of

hook development. Analyses by chemical treatments of the impact of cytokinin indicate

that the role of cytokinin in these developmental phases is coupled with ethylene. In

contrast, the ethylene biosynthesis inhibitor AVG, applied simultaneously with cytokinin,

could not rescue the cytokinin effect on the opening phase. Accordingly, cytokinin also

delayed the opening phase in the ethylene-insensitive mutant etr1-3, suggesting that the

cytokinin control over the hook opening might be independent of ethylene.

The cytokinin effect on the apical hook development might be linked partly with

the modulation of the auxin activity. Our results indicate that cytokinin interferes with the

auxin distribution by affecting the polar auxin transport via the regulation of the PIN

auxin efflux carriers. For instance, a strong downregulation of PIN1 and PIN4 might

underlie the phenotype observed after cytokinin treatment and migh cause defects in the

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formation and maintenance of auxin maxima at the concave side of the hook that are

required for proper apical hook development. Interestingly, we found that a fully working

polar auxin transport machinery is crucial for the impact of cytokinin. In the absence of

one of the PIN genes, the cytokinin effect, represented by a halted opening phase

prolonging opening of the hook opening, disappears.

Further, investigation of auxin response in cytokinin receptor mutants showed a

reduction in the DR5 expression in the hook of cre1-12/ahk4 that was correlated with a

defective apical hook development. This finding might indicate that a low auxin response

is not sufficient to maintain the hook closed, with a premature opening as a consequence.

The ahk3-3 mutant did not display any significant changes in the DR5 expression,

presumably because of the weaker phenotype in the opening phase than that of the cre1-

12/ahk4 mutant. Therefore, it would be interesting to investigate auxin responses in later

developmental phases, when the phenotype is more pronounced. Expression analysis of

PIN proteins revealed an upregulation of PIN3 in the ahk3-3 mutant, suggesting that PIN3

is presumably important to keep auxin maxima at the concave side of the hook during the

maintenance phase. Accordingly, downregulation of PIN3 in the cre1-12/ahk4 mutant

might reduce the auxin levels at the concave side of the hook. Analysis of PIN1-GFP in

cytokinin receptor mutants displayed only difference in the cre1-1/ahk4 mutant: the

PIN1-GFP signal had slightly increased, hinting at a possible role of PIN1 in cre1-

1/ahk4, whereas the upregulated PIN1-GFP might lead to a faster auxin depletion

followed by an enhanced opening of the hook.

Our data suggest that cytokinin might play a prominent role in (i) the formation

phase, in which, together with other hormones, it is responsible for the establishment of a

proper hook; (ii) the maintenance phase, in which, also with other hormones, it is

important to keep the hook adequately closed; (iii) the control of the transition from the

maintenance to the opening phases and the prevention of a premature hook opening; and

(iv) preservation of the auxin maxima at the concave side of the hook and inhibition of

premature diffusion of auxin from the hook zone by interference with the polar auxin

transport machinery.

Cytokinin receptors are involved in the regulation of apical hook development

Although the role of cytokinin in apical hook development has been reported to be

mediated by ethylene (i.e. by increased ethylene biosynthesis) (Cary et al., 1995; Vogel et


135

al., 1998), our data suggest that the cytokinin effect on this developmental process is

partly independent of ethylene. Additionally, particular cytokinin receptors play an

important, but distinct, role that is tightly coupled with the ethylene regulation.

Cytokinin receptors are differentially expressed during apical hook development.

AHK2 is expressed exclusively at the beginning of the opening phase; thus, AHK2 might

modulate hook development during the transition from the maintenance to the opening

phase. AHK3 is expressed during hook formation and maintenance, suggesting its

involvement in the formation and maintenance phases. Expression of CRE1-12/AHK4

was stable during all phases of hook development, implying its contribution to the entire

process of hook development.

Exclusion of cytokinin receptor genes led to particular defects in apical hook

development. The phenotype of cytokinin receptor loss-of-function mutants supports their

essential role in the maintenance phase and during the maintenance-to-opening phase

transition. However, AHK2 seems to promote the transition from the maintenance to the

opening phase and opening of the hook, whereas AHK3 and CRE1-12/AHK4 prevent

hook opening. AHK3 alone presumably plays an important role in the cytokinin

sensitivity during the transition from the maintenance to the opening phase, in which it

prevents premature hook opening. In addition, cytokinin receptors appeared to have

distinct effects on the ethylene biosynthesis and signaling: AHK2 seemed to be a

suppressor and AHK3 and CRE1-12/AHK4 presumably acted as positive regulators during

hook development. Interestingly cytokinin mutants had no impact on the ethylene

biosynthesis genes from ACO family, because in the cytokinin receptor mutants, no

dramatic changes in their expression were observed. We hypothesize that probably all the

cytokinin activity during hook development is tightly balanced and important for the

interaction with ethylene and auxin.

Interestingly, recently published data show that cytokinin increases ethylene

biosynthesis via a posttranscriptional mechanism (Vogel et al., 1998; Woeste et al., 1999)

that blocks the interaction between the ACS and ETO1 proteins, preventing the

degradation of ACS with an increased ACS protein stability as a consequence (Chae and

Kieber, 2005; Hansen et al., 2009). Although our findings support previous observations

that the cytokinin activity might be mediated through interaction with the ethylene

pathway, our data suggest a different mode of regulation, in which cytokinin via AHK3

and CRE1/AHK4 promotes transcription of the ACS genes and via AHK2 might block

their transcription.

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Downstream elements in cytokinin signaling during hook development

Once cytokinin is perceived, the receptor signals are transduced by a multistep

phosphorelay system through a complex consisting of the two-component system

pathway. All the components are present as gene families and include histidine kinases,

histidine-containing phosphotransfer proteins (AHPs), and two classes of response

regulators (ARRs) (Schmülling, 2004). First, we focused on the Arabidopsis histidine

kinases (cytokinin receptors) and then on the downstream elements of the phosphorelay

signaling, the ARRs. Our data reveal that the functional cytokinin receptors, several

transcriptional B-type ARR regulators, and negative A-type regulators are required for the

cytokinin-mediated apical hook development. Cytokinin receptor mutants displayed two

types of phenotypes: the ahk2-2 phenotype, in which the apical hook was exaggerated and

the maintenance phase was prolonged followed by a slow hook opening; and the ahk3-3

and cre1-12/ahk4 phenotype, in which the maintenance phase was shorter followed by a

faster hook opening. Real-time phenotype analyses of the B-type ARR transcription

factor mutants arr1,arr2, arr10, arr11, arr12, and arr13 reveal that phenotypes

reminiscent of ahk3-3 and cre1-12/ahk4, suggesting a function for the genes ARR1,

ARR2, ARR10, ARR11, ARR12, and ARR13 as positive regulators of the cytokinin

pathway that might act downstream of AHK3 and CRE1-12/AHK4. However, none of the

B-type ARR loss-of-function mutants that were examined displayed either an exaggerated

hook or a prolonged maintenance phase, which would be expected of B-type ARR

regulators acting downstream of AHK2. Further analyses have to be done to examine

other combinations of multiple or single mutants. Thus far, seven out of the 11 types of

ARR genes have been analyzed. Apical hook development in negative A-type ARR

mutants corresponded to that observed either in ahk2, ahk3, and ahk4/cre1 mutants.

According to their respective phenotypes and their presumed function as negative

regulators of the cytokinin pathway, we speculate that ARR9 and ARR16 mediate a

negative feedback on AHK3 and CRE1-12/AHK4 and ARR3 and ARR6 on the AHK2-

controlled branch of the cytokinin signaling pathway.

Model of cytokinin signaling in apical hook development

Based on our data on the role of the components of the cytokinin signaling

pathway in apical hook development, we propose a model to explain the possible


137

molecular scenario of the cytokinin-controlled hook development (Fig. 6). Typically, the

cytokinin signal is perceived by cytokinin receptors that autophosphorylate and transmit

phosphoryl groups to the AHPs. The AHPs activate B-type ARRs in the nucleus and,

consequently, downstream transcriptional responses. A negative feedback loop is ensured

by the A-type ARRs that inhibit, by a still unknown mechanism, the activity of the B-type

ARR transcription factors. However, the mechanism is still unknown by which cytokinin

control the common developmental process, such as apical hook development, through

multiple cytokinin receptors. The question remains whether there are specific functions

for individual receptors in this regulation. Detailed analyses of cytokinin receptors and

downstream signaling components indicate that they regulate the apical hook

development in a specific, even antagonistic, manner. Hence, we propose a model is in

which the cytokinin receptors through specific interactions with the ethylene pathway

control the ethylene activity and, thus, differentially contribute to the regulation of apical

hook development (Fig. 6).

Based on the downregulation of genes involved in ethylene biosynthesis and

signaling in the ahk3-3 mutants, the cytokinin receptor AHK3 might act as the positive

regulator of the ethylene pathway. The cytokinin signal perceived by AHK3 is

presumably further transmitted through downstream components involved in this

regulation, namely ARR1, ARR2, ARR10, ARR11, ARR12, and ARR13, the positive

regulators from the B-type ARR family. A CRE/AHK4-mediate pathway might positively

regulate the ethylene pathway as indicated by the modest downregulation of the genes

involved in the ethylene biosynthesis in the cre1-12 mutant. On the contrary, a different

mode of regulation occurs via AHK2. The AHK2-mediated pathway suppresses the

ethylene activity, because most of the ethylene biosynthesis (interestingly, not those from

the ACO family) and signaling genes are upregulated in the akh2-2 mutant. Based on the

phenotypes of the A-type loss-of-function mutants and their presumed function as

negative regulators of the cytokinin signaling pathway, we propose that ARR9 and ARR16

mediate a negative feedback on AHK3 and CRE1-12/AHK4, whereas ARR3 and ARR6 on

the AHK2-controlled branch of the cytokinin signalling pathway. We speculate that this

pathway might be parallel to the previously identified cytokinin regulation occurring at a

posttranslational level. Indeed, cytokinin has been found to stabilize the ACS enzyme

and, thus, to enhance ethylene biosynthesis (Hansen et al. 2009).

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MATERIALS AND METHODS

Plant material

The transgenic Arabidopsis thaliana (L.) Heynh. lines were acquired at the

European Arabidopsis Stock Centre (Nottingham, UK): Col-0, Col-8, Wassilewskija and

arr2-1 (SALK_043107C), arr13-1 (N655053), arr16-1 (N667506) (Alonso et al., 2003),

arr1-2 (N6368), arr10-1 (N6369), arr12-1 (N6978) arr11-1 (N6370), arr11-2

(SALK_013832C), arr1-2xarr11-1(N6988), arr1-3xarr12-1 (N6981), arr1-3xarr11-2

(N6980) (Mason et al., 2005), arr3-1 (SALK_042068C), arr4-1 (CS25266), arr5-1

(CS25267), arr6-1 (SALK_080024C), arr9-1 (CS25270), arr1-3xarr10-5xarr12-1

(N39992), arr3xarr4 (CS25271), arr3xarrx4arr5xarr6 (CS25276),

arr3xarr4xarr5xarr6xarr8xarr9 (CS25279), arr8xarr9 (CS25275) (To et al., 2004), arr7

(N858131) (part of Wisconsin DsLox T-DNA lines), arr14-1 (N875481) (Sessions et al.,

2002), arr15-1 (N411750) (part of GABI-Kat), arr1-3, arr10-5, arr12-1 (N39992)

(Argyros et al.,2008).

Other transgenic Arabidopsis thaliana (L.) Heynh. lines have been described

elsewhere: ahk4/cre1-12; ahk2-2; ahk3-3; ahk4/cre1-12xahk2-2; ahk4/cre1-12xahk3-3;

ahk2-2xahk3-3 (Higuchi et al., 2004); CRE1/AHK4::GUS; AHK2::GUS; AHK3::GUS

(Higuchi et al., 2004); (mutants of ahk4/cre1-12; ahk2-2; ahk3-3 were crossed with

DR5::GFP, PIN1::PIN1-GFP and PIN3::PIN3-GFP), ein2, etr1-3 (Roman et al., 1995),

DR5::GUS (Sabatini et al., 1999); DR5::GFP, PIN1::PIN1-GFP (Benková et al., 2003);

PIN3::PIN3-GFP (Ţádníková et al., 2010), PIN4::PIN4-GFP; PIN7::PIN7-GFP (Blilou

et al., 2005); CycB1;1::GUS (Ferreira et al., 1994); pin1-3 (Okada et al., 1991); the pin1

phenotype was analyzed on the heterozygous population (apical hook development was

recorded, seedlings were numbered, and the pin1 homozygotes were identified based on

pinformed-like shoot phenotypes in the adult stage); pin4-3 (Friml et al., 2002a); pin3-4,

pin7-1 (Friml et al.,2003) (mutants pin3-4¸ pin4-3, and pin7-1 were crossed with

DR5::GFP), EBS::GUS1-11 (Stepanova et al., 2007), 35S:IPT3 (Galihet et al., 2008),

35S:CKX3 (Pernisová et al., 2008), ERS1::GUS, ETR1::GUS (Grefen et al., 2008), acs5,

acs7, and acs8 (Tsuchisaka et al.,2009).


139

Growth conditions

Seeds were surface sterized by ethanol, sown on half-strength Murashige and

Skoog (MS) medium (Duchefa) with 1% sucrose, 0.8% agar (pH 5.7), or supplemented

either with 5 µM 1-aminocyclopropane-1-carboxylate (ACC; Sigma-Aldrich), 0.1 µM 6-

benzylaminopurine (BAP; Sigma-Aldrich), or 0.25 µM 2-aminoethoxyvinylglycine

(AVG; Sigma-Aldrich), vernalized for 2 days at 4°C, exposed to light for 12 hours at

18°C to synchronize the germination start, and cultivated in the dark at 18°C. Seeds were

used for real-time phenotype analysis and 24, 48, 72, and 84 hours after germination

seedlings were imaged by confocal microscopy, stained with GUS, or used for transverse

sectioning.

Transverse sections

DR5::DR5-GFP seedlings or mutant seedlings crossed with DR5::DR5-GFP 24 hours

after germination were processed as described previously (Ţádníková et al., 2010). The

DR5-GFP signal was quantified; the number of cells with the DR5 signal was divided by

the total number of cells in the apical hook; for example, 14 signal-positive cells out of 29

is equal to 48%. VSNI GenStat 14th was used for statistical analysis (p-value for t-test

0.05). Error bars represent standard errors.

Real-time analysis of apical hook development

Seedling development was recorded at 1-hour intervals for 7 days at 18°C with an


(Canon Rebel T2i, 550DH, spectrum enhanced camera and EF-S 18-55 mm, IS lens kit

with built-in clear filter wideband-multicoated and standard Canon accessories) and

operated by the EOS utility software. Angles between the hypocotyl axis and cotyledons

were measured by ImageJ (NIH; http://rsb.info.nih.gov/ij). Fifteen seedlings with

synchronized germination start were processed. Error bars represent standard errors.

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Histochemical analysis of GUS activity

Histochemical GUS staining was done as published (Friml et al., 2003) for

24 hours at 37°C. Seedlings mounted in chloral hydrate (Fluka) were analyzed with a

automatic virtual slide scanner dotSlide frame (BX51 microscope; Olympus) and 10x

UPLSAPO objective equipped with a digital CCD camera (2/3” CCD camera, 6.45 µm x

6.45 µm pixel size, high sensitivity, high resolution, Peltier cooled, dynamic range of 3 x

12 bit). Images were processed in Adobe Photoshop.

Confocal microscopy

For confocal microscopy images, Zeiss LSM 710 with Zeiss C-Apochromat

63x/1.20 water immersion objective and Leica TCS SP2 AOBS with HC PL APO

20x/0.70 water immersion objective were used. The green fluorescent protein (GFP)

signal after 488-nm argon laser line excitation was detected in the spectral range from

500 nm to 590 nm and 505 nm and 580 nm for the Zeiss and Leica systems, respectively.

The yellow fluorescent protein (YFP) signal after 514-nm argon laser line excitation was

detected in the spectral range from 525 nm to 600 nm for the Zeiss system.

Quantitative analysis of the GFP signal

The GFP signal on the membranes of lines PIN3::PIN3-GFP, PIN4::PIN-GFP

and PIN7::PIN7-GFP were analyzed quantitatively as described (Ţádníková et al., 2010)

and the DR5-GFP signal from full z-stack confocal images with 3D projection with the

Leica confocal software. The fluorescence intensity of DR5-GFP on average projection

pictures in the middle of the epidermal cell at the concave side was quantified by ImageJ.

The same size area was used in all measurements. At least 10 seedlings were analyzed per

treatment. VSNI GenStat 14th was used for the statistical analysis (p-value for t-test

0.05). Error bars represent standard errors.

Quantitative RT-PCR

RNA for quantitative RT-PCR was extracted with the RNeasy kit (Qiagen) from

48-hour-old Arabidopsis seedlings. A DNase treatment with the RNase-free DNase Set


141

(Qiagen) was carried out for 15 min at 25°C. Poly(dT) cDNA was prepared from 1 μg

total RNA with the iScript™cDNA Synthesis Kit (Bio-Rad) and quantified with a

LightCycler 480 (Roche) and SYBR GREEN I Master (Roche), according to the

manufacturer‟s instructions. PCR was carried out in 384-well optical reaction plates

heated for 10 min to 95°C to activate hot start Taq DNA polymerase, followed by 40

cycles of denaturation for 60 sec at 95°C and annealing/extension for 60 sec at 58°C.

Targets were quantified with specific primer pairs designed with the Beacon Designer 4.0

(Premier Biosoft International). Expression levels were normalized to the levels of

UBIQUITIN10 (UBQ10). All RT-PCR experiments were done at least in triplicate. For

the primer list, their sequence, and efficiency factor of the tested genes corresponding to

UBQ10 together with the standard error, see Table S1 in supplementary material.

Ethylene measurements

Ethylene was measured as published (Bijnen, et al., 1996).

ACKNOWLEDGMENTS

We thank Martine De Cock for help in preparing the manuscript. This work was

supported by grants from the European Research Council (Starting Independent Research

Grant ERC-2007-Stg-207362-HCPO to E.B).

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SUPPLEMENTARY MATERIAL

A ERS1::GUS

D EBS::GUS BAP


E EBS::GUS ACC

C EBS::GUS MS

B ETR1::GUS

Figure S1. Expression analysis

(A) Expression of ERS1::GUS in 26 HAG old untreated seedlings. (B) Expression of ETR1::GUS in 26

HAG old untreated seedlings. (C) Expression of EBS::GUS in untreated seedlings (MS) in formation,

maintenance an d opening phase. (D) Expression of EBS::GUS upon cytokinin treatment (BAP) in

formation, maintenance and opening phase. (E) Expression of EBS::GUS upon ethylene treatment (ACC) in

formation, maintenance and opening phase.


143

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Figure S2. Cytokinin and ethylene sensitivity of cytokinin recptor mutants

(A). Kinetics of apical hook development in untreated (MS) or cytokinin (BAP) treated Col-8 and ahk2-2

seedlings. (B) Kinetics of apical hook development in untreated (MS) or cytokinin (BAP) treated Col-8 and

ahk3-3 seedlings. (C) Kinetics of apical hook development in untreated (MS) or cytokinin (BAP) treated

Col-8 and cre1-12 seedlings. (D). Kinetics of apical hook development in untreated (MS) or ethylene

(ACC) treated Col-8 and ahk2-2 seedlings. (E) Kinetics of apical hook development in untreated (MS) or

ethylene (ACC) treated Col-8 and ahk3-3 seedlings. (F) Kinetics of apical hook development in untreated

(MS) or ethylene (ACC) treated Col-8 and cre1-12 seedlings.

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Figure S3. Changes in expression in PIN-GFP upon cytokinin treatment

(A) Expression of PIN1-GFP in 46 HAG old untreated seedlings (MS). (B) Expression of PIN1-GFP in 46

HAG old cytokinin treated seedlings (BAP). (C) Quantification of relative fluorescence on the membrane of

PIN1-GFP in 72 HAG old untreated seedlings, on transversal membranes in epidermal cells on the concave

side of the hook (n = 10). (D) Expression of PIN3-GFP in 36 HAG old untreated seedlings (MS). (E).

Expression of PIN3-GFP in 36 HAG old cytokinin treated seedlings (BAP). (F) Quantification of relative

fluorescence on the membrane of PIN3-GFP in 36 HAG old untreated seedlings, on transversal membranes

in cortical cells on the convex and concave cells of the hook (n = 10). (G). Expression of PIN4-GFP in 36

HAG old untreated seedlings (MS). (H) Expression of PIN4-GFP in 36 HAG old cytokinin treated

seedlings (BAP). (I) Quantification of relative fluorescence on the membrane of PIN4-GFP in 36 HAG old

untreated seedlings, on transversal membranes in cortical cells on the convex and concave cells of the hook

(n = 10). (J) Expression of PIN7-GFP in 36 HAG old untreated seedlings (MS). (K) Expression of PIN7-

GFP in 36 HAG old cytokinin treated seedlings (BAP). (L) Quantification of relative fluorescence on the

membrane of PIN7-GFP in 36 HAG old untreated seedlings, on transversal membranes in cortical cells on

the convex and concave cells of the hook (n = 10). Error bars represent s.e.m. (*p<0.05, n=3)


145

Figure S4. Sensitivity of pin mutants to cytokinin

(A) Kinetics of apical hook development in untreated (MS) or cytokinin (BAP) treated PIN1-3 control

seedlings and pin1-3 mutant seedlings. (B) Kinetics of apical hook development in untreated (MS) or

cytokinin (BAP) treated Col-0 seedlings and pin3-4 mutant seedlings. (C). Kinetics of apical hook

development in untreated (MS) or cytokinin (BAP) treated Col-0 seedlings and pin4-3 mutant seedlings.

(D) Kinetics of apical hook development in untreated (MS) or cytokinin (BAP) treated Col-0 seedlings and

pin7-1 mutant seedlings.

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Figure S5. Expression analysis, changes in PINs expression upon cytokinin

(A) Expression of DR5-GFP in 36 HAG old untreated seedlings (MS). (B) Expression of pin3-4x DR5-GFP

in 36 HAG old untreated seedlings (MS). (C) Expression of pin4-3xDR5-GFP in 36 HAG old untreated

seedlings (MS). (D) Expression of pin7-1xDR5-GFP in 36 HAG old untreated seedlings (MS). (E)

Expression of DR5-GFP in 36 HAG old cytokinin treated seedlings (BAP). (F) Expression of pin3-4x DR5-

GFP in 36 HAG old cytokinin treated seedlings (BAP). (G) Expression of pin4-3xDR5-GFP in 36 HAG old

cytokinin treated seedlings (BAP). (H) Expression of pin7-1xDR5-GFP in 36 HAG old cytokinin treated

seedlings (BAP). (I) Quantification of relative fluorescence of DR5-GFP in 36 HAG old untreated (MS)

and cytokinin treated (BAP) DR5-GFP seedlings and pin mutants, in one cell in the middle of the concave

side of the hook (n = 10).


147

Figure S6. Table of qPCR primers used in this study

List of the primers, which were used to inspect ethylene biosynthesis and signaling pathway in cytokinin

receptor mutants.

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Chapter 5 Conclusions and future perspectives

Conclusions and future perspectives

154


The aim of this thesis was to deepen our understanding of the hormonal regulation

of apical hook development in Arabidopsis thaliana with specific focus on auxin,

cytokinin, and ethylene (Vandenbussche and Van Der Straeten, 2004). The plant hormone

auxin has been described as an important regulator of differential growth during the

apical hook development. Previous observations have suggested that an auxin gradient is

established in the apical region, resulting in differential growth followed by hook

formation. Hook establishment depends on an asymmetric auxin distribution, highlighting

the importance of a functional polar auxin transport (Friml et al., 2002). Indeed,

imbalances in auxin levels provoke defective hook formation or hookless phenotypes, as

demonstrated by the auxin-overproducing yuc1 mutant (Boerjan et al., 1995; Zhao et al.,

2001), auxin-resistant mutants (Tian and Reed, 1999), and treatments with the auxin

efflux inhibitor 1-naphthylphthalamic acid (Schwark and Schierle, 1992; Lehman et al.,

1996). Additionally, ethylene as well has been reported to be involved in apical hook

development because it controls hook maintenance and intensifies the apical hook

curvature (Bleecker et al., 1988; Guzman and Ecker, 1990). However, the molecular

mechanisms remain unclear by which the auxin gradient is established and ethylene

prompts differential growth. Cytokinins applied on dark-grown seedlings of Arabidopsis

have been shown to be able to enhance apical hooks, similarly to ethylene, implying that

cytokinins play a role on apical hook development through an increase in ethylene

biosynthesis (Cary et al., 1995; Vogel et al., 1998b). Although the interactions of most

plant hormones have been studied extensively, the molecular basis for the hormonal

crosstalk between auxin, ethylene, and cytokinin is still largely unknown, especially in

apical hook development (see chapter 1 for details).

From a technical point of view, it was essential to set up experimental methods

that could be further used to study apical hook development. Previously, numerous

reports have documented phenotypic changes in seedlings at a few time points, i.e., at 48

hours after germination (HAG) and 72 HAG, but these analyses did not provide detailed

information on the temporal changes in the phenotypes and were technically unable to

follow precisely the whole process of apical hook development. Therefore, a real-time

phenotypic analysis was developed based on the continuous monitoring of seedlings

grown in the dark by means of a special camera (Canon Rebel T2i-550DH). As the


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complete developmental process could be monitored from seed germination to apical

hook opening, only the germination-synchronized seedlings were analyzed, thus avoiding

comparison between seedlings at different developmental phases due to germination

variability. Our real-time approach also allowed the detailed monitoring of each

developmental phase of the hooks and even minor differences between mutants or treated

seedlings could be identified properly. This new technique was implemented throughout

the elaboration of the thesis. All the results of the phenotypic analysis were obtained with

real-time studies of apical hook development.

Similarly, the determination of the confocal microscopy parameters as well as the

quantitative analysis of the GFP reporter signal were required. The GFP localization in

the apical hook zone was examined by confocal z-sectioning and by further 3D

reconstruction of the obtained sections. Each confocal micrograph presented in this thesis

represents either a single focal plane or a 3D projection of individual images taken as a z-

series from a collection of at least 10 images through the cortex and epidermal layers.

Even though the observation of the signals in the upper parts of the hypocotyls had been

improved by the confocal microscopy, some difficulties remained, such as the detection

with the GFP signals of the exact cell layers. Therefore, it was essential to establish a

method in which it would be possible to explore the GFP signals on transversal sections.

The vibratome was used to prepare transversal sections of paraformaldehyde-fixed hooks

expressing the GFP material. So, it was possible to investigate the exact place of the GFP

expression in all layers of the hypocotyl.

Concluding remarks

To assess the role of auxin in apical hook development, the auxin-responsive

synthetic promoter DR5 (Ulmasov et al., 1997) was fused with the GUS or GFP reporters.

During hook formation, the DR5 expression was localized on the concave (inner) side of

the hook, remained throughout the maintenance phase, and gradually disappeared during

the opening phase. Subsequently, the contribution of the PIN proteins on the asymmetric

distribution in the apical hook was investigated. The PIN genes were found to display

specific, partly overlapping, spatial and temporal expression patterns in the hooks.

Moreover, because the elimination of some PIN proteins by mutation interfered with

particular phases of the hook development, we concluded that PIN1, PIN3, PIN4 and

PIN7 participate in the dynamic auxin redistribution during hook development. Next,


156

ethylene was shown to postpone the transition between the formation and maintenance

phases, presumably by changes in the auxin distribution and by the interaction with the

polar auxin transport machinery through modulation of the expression of several PIN

genes. Based on these results and combined with published data on the auxin influx

(Vandenbussche et al., 2010), we proposed a model in which the coordinated activities of

LAX3 and the ethylene-regulated AUX1 and PIN proteins control the direction of the

auxin transport. Whereas the auxin transport in the cotyledons, meristem, and hypocotyl

tip is basipetal toward the root, a lateral transport occurs toward the outer hypocotyl tissue

layers, establishing an auxin gradient that regulates the apical hook development (chapter

2).

In a further research, we attempted to answer the following questions: (i) which

are the requirements for the spatial auxin redistribution during the apical hook formation;

(ii) how does ethylene impact on the auxin redistribution; and (iii) which is the role of cell

division during the apical hook development? First, data from the quantification analysis

on the PIN3-GFP signal were combined with computational approaches. The asymmetric

expression of PIN genes on the outer (convex) side of the hook was shown to be a driving

force for the auxin redistribution and consequent establishment of the auxin maximum on

the inner (concave) side of the hook. The computer model predictions were validated by

quantitative experimental analyses of the auxin response (i.e. the DR5 signal) and the

dynamic localization of the PIN proteins in both the wild-type and mutant plants. Our

dynamic model explains where and how the auxin gradient is translated into differential

growth in the hook (chapter 3).

Subsequently, to explore the contribution of cytokinins to the differential growth

regulation in apical hook development, we focused on the molecular mechanisms

underlying the hormonal crosstalk between auxin and cytokinin (chapter 4). Our results

suggest that cytokinin is an important regulator of differential growth for proper hook

establishment, with particular functions at the different stages of hook development.

Analysis of the cytokinin impact revealed the role of cytokinins is coupled with ethylene

in the both formation and maintenance phases, whereas the control of the opening phase

by the cytokinins is independent of ethylene. In addition, by combining gene expression

and mutant phenotype analysis, we observed that individual cytokinin receptors play

distinct roles in apical hook development. AHK2 operates as a negative regulator of the

formation phase and as a positive regulator of the hook opening. In contrast, AHK3 and

CRE1/AHK4 are important positive regulators of the maintenance phase, but negatively


157

regulate the opening phase. The functions of cytokinin receptors are closely linked with

the ethylene biosynthesis and signaling, but are opposite in their effect on the ethylene

pathway. AHK2 probably acts as a suppressor of the ethylene biosynthesis and signaling,

whereas AHK3, together with CRE1/AHK4, seems to induce the above-mentioned

pathways. Moreover, we identified downstream regulatory components that allowed us to

propose a model in which the potential role is elucidated of the cytokinin signaling in the

regulation of the ethylene pathway during apical hook development. Finally, our results

demonstrated that cytokinin interferes with the polar auxin transport by modulating the

transcription of several PIN auxin efflux carriers and that a fully working polar auxin

transport machinery is crucial for the cytokinin effect.

Future perspectives

The apical hook is a well described developmental process that depends on

differential growth and that it is tightly regulated by several plant hormones. The full

development of the apical hook can be followed easily and is simple to work with,

making the apical hook a very attractive model system to study the differential growth-

controlling hormonal crosstalk.

Although the data obtained in this thesis have improved our understanding on the

hormonal regulation of the apical hook development, many questions remain unanswered.

The various components affecting the different stages of the apical hook development

should be further elucidated, hence, deepening our understanding of the hormonal

regulation and the implications of differential growth in apical hook formation.

In dicotyledonous plants, the apical hook is formed soon after germination, after

emergence of young seedlings from the seed coat (Raz and Ecker, 1999). Thus, it would

be interesting to study which components govern apical hook formation and also to

explore whether this process is already predetermined during embryogenesis. Moreover,

it would be of remarkable importance to elucidate whether apical hook formation is

determined by internal factors, as suggested by Darwin (Darwin and Darwin, 1881) and

Rubinstein (Rubinstein, 1972), or by external forces, such as gravity (Karve, 1964) or

light (Liscum and Hangarter, 1993a).

All the research done so far on apical hook development has been focused on

exploring the asymmetric auxin distribution with the DR5 reporter only, a synthetic auxin

response promoter (Ulmasov et al., 1997; Vandenbussche et al., 2010; Ţádníková et al.,


158

2010). In the future, it would be appealing to use the novel auxin signaling sensor DII-

VENUS (Vernoux et al., 2011; Brunoud et al., 2012). This reporter forms a yellow

fluorescent protein (Shaner et al., 2005) fused to the Aux/IAA interaction domain

(designated domain II or DII) (Tan et al., 2007) and is expressed under a constitutive

promoter. Thus, any increase in auxin levels would cause the degradation of the reporter.

The use of DII-VENUS could allow us to visualize dynamically the auxin input into the

signaling pathway leading to apical hook development. This experiment might provide

spatio-temporal information about the auxin distribution and response during the whole

process of apical hook development.

Although the PIN auxin efflux carriers seem to be crucial auxin transporters that

mediate the auxin redistribution in the hook (Ţádníková et al., 2010), it might be

important to investigate the implication of other auxin transporters possibly involved in

the asymmetric auxin accumulation during the hook development. For example, it would

be interesting to explore the role of the recently discovered novel putative auxin transport

facilitator family, designated PIN-LIKES (PILS) (Barbez et al., 2012) during apical hook

development. As the PILS proteins regulate the intracellular auxin accumulation at the

endoplasmic reticulum and, thus, the auxin availability for the nuclear auxin signaling,

they might have an important function in hook development. Here too, further

investigation of other endoplasmic reticulum-localized auxin efflux carriers in apical

hook development should be assessed; for example, the recently published auxin

transporters from the PIN family, PIN5 (Mravec et al., 2009) and PIN8 (Ding et al., 2012)

have been reported to regulate differential growth, hinting at their involvement in apical

hook development as well. In addition, P-glycoproteins (PGPs) have been found to

interact with PIN proteins and, together, to affect the efflux system dependent on PIN and

PGP that is required for asymmetric distribution during several developmental processes

(Mravec et al., 2008).

Most of the recent analyses on apical hook development have been carried out

exclusively under dark conditions, in which the apical hook could open passively without

any light pulse (De Grauwe et al., 2005b; Vandenbussche et al., 2010; Ţádníková et al.,

2010; Gallego-Bartolome et al., 2011b; Willige et al., 2011). However, under normal

conditions, light is the major factor that influences the hook opening. It would be

extremely interesting to investigate the molecular mechanisms related to light signaling

during apical hook development as well as to answer questions, such as (i) which are the

requirements for light-mediated hook opening; (ii) do the auxin redistribution and the


159

polar auxin transport machinery interact; and (iii) are any other hormones involved in this

pathway, such as cytokinin or ethylene? Undoubtedly, future experiments and answers to

the open questions would help elucidate the molecular mechanisms that govern

differential growth resulting in apical hook development.


160

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309.

Summary

163

Summary

Developmental programs of plants and animals have evolved separately for

millions of years. Despite the fact that both kingdoms share several common cell

signalling and gene regulation mechanisms, differences between them are already

obvious during embryogenesis. While animals develop their organs during zygotic

embryogenesis and their mature embryo constitutes a miniature version of the adult

organism, plant embryogenesis generates a relatively simple structure and the adult plant

morphology is formed by post-germination activity of apical meristems. The meristem is

a special tissue consisting of undifferentiated, actively growing and dividing cells,

responsible for the maintenance of a stable stem cell pool and plant growth. Just after

seed germination during early plant growth towards the soil surface, the exposed shoot

apical meristem could be damaged. Therefore, plants evolved specialized mechanisms to

protect the delicate meristematic tissue. In dicotyledonous plants, the hypocotyl bends

and forms the apical hook, in which the meristematic tissue is placed below the hook

region and thus is protected from mechanical injuries. Similar to many of the

morphogenetic changes in developing plants, the apical hook development also must be

precisely regulated. Plant hormones have been described as important endogenous

regulators of several physiological processes in plant development. Nevertheless, the

specific roles of plant hormones during apical hook development remain largely unknown

(chapter 1).

The main goal of this thesis was to understand how plant hormones regulate

differential growth during apical hook formation, with a special attention to hormonal

interactions among auxin, ethylene and cytokinin.

We show that the progress of the apical hook development through distinct phases

depends on the dynamic, asymmetric distribution of auxin, which is controlled by auxin

efflux carriers of the PIN family. Several PIN proteins have specific, partly overlapping

spatio-temporal expression patterns in the hook. Elimination of PIN proteins by mutation

or knock-out interferes with particular phases of hook development. Furthermore,

ethylene-induced enhancement of apical hook formation and subsequent hook curvature

exaggeration is accompanied by changes in the expression of several PIN genes and,

consequently, in auxin distribution (chapter 2).

Summary

164

Experimental data from signal quantification of PIN3-GFP on cortical membranes

revealed enhanced expression of PIN3 on convex side of the hook. These data were

combined with computational approaches. The mathematical model helped us to gain a

better and more detailed insight into the requirements for spatial auxin redistribution

during hook formation. Computer model predictions were further validated with the

quantitative analysis of PIN-GFP dynamics and inspection of auxin response in wild type

and PIN mutant seedlings, and it was demonstrated that asymmetric expression of PINs

auxin efflux carriers on the outer (convex) side of the hook is a driving force for the

redistribution of auxin toward concave side and consequent establishment of auxin

maximum on the inner (concave) side of the hook. A dynamic model explains where and

how the auxin gradient is translated into differential growth in the hook zone. Moreover,

our findings reveal the crosstalk between auxin and ethylene that affects PIN protein

dynamics, and thus auxin distribution, during apical hook development (chapter 3).

Subsequently, the contribution of cytokinins to differential growth during apical

hook development was evaluated. By combining genetic approaches and expression

analysis, the role of cytokinin signalling during hook development was explored. We

found that the cytokinin activity is coupled with ethylene in early stages of apical hook

development while cytokinin controls the opening phase independently of ethylene. Next,

opposite functions of particular cytokinin receptors and downstream components of the

signalling pathway in the regulation of apical hook development was revealed. Part of

these specific effects might be related to differences in the interaction with the ethylene

pathway. Finally, we show that cytokinin modulates auxin distribution during hook

development by affecting polar auxin transport via interfering with the expression of

particular PINs (chapter 4).

Overall, this PhD research provides novel and important insights into the

regulation of differential growth in plants through the coordinated crosstalk between

individual hormones.

Acknowledgements

165

Acknowledgements

I would like to take this opportunity to express my gratitude to the numerous

people who supported me in the achievement of this degree. Foremost, my supervisor Eva

for providing me an opportunity to work in her excellent lab and to give me an

interesting, fascinating project. Her guidance, advice and enthusiasm have been

invaluable and her patience commendable.

I especially would like to give thanks to:

- Jiří, for his support and mainly for convincing me to change a clearly

predetermined future as a pharmacist to an unsure life as a scientist:);

- Martine de Cock for great help in preparing my thesis;

- all of my Crosstalks‟ colleagues - Agnieszka, Anas, Candela, Christa, Jerome,

Jose, Maria, Marleen and Peter for a pleasant time during our lunch time,

friendship and a enjoyable atmosphere in the lab;

- Eva and Hanka for their friendship, very nice conversations, the parties and a

lot of other things that I will keep in my heart;

- Peter, without his excellent company, and all the lunches, dinners and

enjoyable discussions we had together, my life here would not have been as

pleasant;

- all the people from different groups in the department for their kindness,

professionalism and for sharing their knowledge. Special thanks go to the

Auxin, Brassinosteroids, Root development and Proteomic groups for creating

an enjoyable working atmosphere;

- everyone from the Bioenergy lab, especially Brecht, Lisa, Elis, Dorien and

Ward. Thank you for accepting me so graciously into your group and for

including me in all the dinners, coffees, trips, and much, much more;

- the Flamingo house mates (Mira, Vanesa, Peter) for the special, friendly

atmosphere and nice discussion;

- Krzysiek for all the tips and scientific discussions;

- all of my swimming and skate-mates, especially to Camilla, Yuliya, Boris,

Nathalie, Kristof. Great mates for great time spent together;

- everybody from the PSB department for making this period one towards which

I will look back with joy and nice memories, and I will never forget you all;

- my family (my sister and my mum) for support in critical phases of my life.

I also wish, at this time, to acknowledge Igor, for which I could name plenty of

reasons, but here I will mention excellent support and great inspiration.

And last, this thesis is dedicated to my dad, Štěpán Ţádník, who passed away

during my Ph.D. studies. He remains my inspiration. Due to his advice, I took this

challenge to do my Ph.D. studies. Without him and his support, I would never have been

where I am now and who I am.

Thank you very much!

Curriculum vitae

167

Curriculum Vitae

Personal data

Name: Petra Žádníková Date of Birth: October 22, 1982

Place of Birth: Uherské Hradiště (Czech Republic)

Nationality: Czech

Address: Nedakonice 471, 687 38 Uherské Hradiště (Czech Republic)

Phone: +420 605 865 496

e-mail: [email protected]

Languages: Czech (native)

English (fluent)

French (basic)

Education

2007 - 2012: PhD in Plant Biotechnology

- 2010-2012 Department of Plant Biotechnology and Bioinformatics, Ghent

University and Department of Plant Systems Biology, VIB, Gent (Belgium)

- 2008-2009 Masaryk University, Laboratory of Molecular Plant Physiology,

Department of Functional Genomics and Proteomics, Institute of Experimental

Biology, Brno (Czech Republic)

- 2007-2008 Department of Plant Biotechnology and Bioinformatics, Ghent

University and Department of Plant Systems Biology, VIB, Gent (Belgium)

- Subject: The role of hormonal interactions in differential plant growth

- Promoter: Prof. Dr. Jiří Friml; co-promoter: Dr. Eva Benková

2002 - 2007: Master in Pharmacy

- University of Veterinary and Pharmaceutical Sciences Brno, Faculty of

Pharmacy, Brno (Czech Republic)

- Undergradute thesis: Influence of programmed cell death (apoptosis) in relation

to the development of disease

- Promoter: Prof. Luděk Beneš

Publications

- Vandenbussche F, Callebert P, Žádníková P, Benková E, Van Der Straeten D:

(2012). Brassinosteroid control of shoot gravitropism interacts with ethylene

and depends on auxin signaling components. Am. J. Bot., in press (accepted).

- Guenot B, Bayer E, Kierzkowski D, Smith RS, Mandel T, Žádníková P,

Benková E, Kuhlemeier C (2012). PIN1-independent leaf initiation in

Arabidopsis. Plant Physiol. 159 (4):1501-1510.

- Gallego-Bartolomé J, Arana MV, Vandenbussche F, Žádníková P, Minguet

EG, Guardiola V, Van Der Straeten D, Benková E, Alabadí D, Blázquez MA

(2011). Hierarchy of hormone action controlling apical hook development in

Arabidopsis. Plant J. 67(4): 622-634.

Curriculum vitae

168

- Žádníková P, Petrášek J, Marhavý P, Raz V, Vandenbussche F, Ding Z,

Schwarzerová K, Morita MT, Tasaka M, Hejátko J, Van Der Straeten D, Friml

J, Benková E (2010). Role of PIN-mediated auxin efflux in apical hook

development of Arabidopsis thaliana. Development 137(4): 607-617.

- Vandenbussche F, Petrášek J, Žádníková P, Hoyerová K, Pešek B, Raz V,

Swarup R, Bennett M, Zaţímalová E, Benková E, Van Der Straeten D. (2010).

The auxin influx carriers AUX1 and LAX3 are involved in auxin-ethylene

interactions during apical hook development in Arabidopsis thaliana

seedlings. Development 137(4): 597-606.

- Bartl T, Žádníková P, Jiříkovská R, Maroušková B, Beneš L. (2008).

Substances modifying the activity of caspases. Česká Slov Farm. 57(1): 29-34.

Manuscripts in preparation

- Žádníková P, Wabnik K, Friml J, Prusinkiewicz P, Benková E. Coupling

auxin distribution to asymmetric growth explains the formation of-apical hook

in Arabidopsis.

- Žádníková P, Vandenbussche F, Van Der Straeten D, Benková E. Role of

cytokins in apical hook development.

- Žádníková P, Benková E. Dark side of the plant growth: role of plant

hormones in apical hook development in Arabidopsis thaliana.

Meetings

Attended meetings

- International Symposium - Growth and Development of Roots, Louvain-la-

Neuve (Belgium), January 27, 2011

Oral presentation

- International Ph.D. Students Conference, Mendel University, Brno (Czech

Republic), June 30-July 1, 2009

- XII Student's Scientific Conference, University of Veterinary and

Pharmaceutical Sciences, Brno (Czech Republic), April 22, 2007

Poster presentation

- International Arabidopsis Conference, Vienna (Austria), July 3-7, 2012

- Plant Growth Biology and Modeling, Elche (Spain), September 19-21, 2011

- Auxins and Cytokinins in Plant Development - International Symposium,

Prague (Czech Republic), July 10-14, 2009

- VIIth Interdisciplinary Meeting of Young Biologists, Biochemists and

Chemists, Devět Skal - Ţďárské vrchy (Czech Republic), June 12-15, 2006

- 35th Conference of Synthesis and Analysis of Drugs, Velké Karlovice (Czech

Republic), June 12-15, 2006

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