Organogenesis in vitro under Altered Auxin Signaling ......in vitro. under Altered Auxin Signaling Conditions . Tatiana Smirnova . Master of Science . Graduate Department of Cell and

Organogenesis in vitro under Altered Auxin Signaling Conditions

by

Tatiana Smirnova

A thesis submitted in conformity with the requirements for the degree of Master of Science

Graduate Department of Cell and System Biology University of Toronto

© Copyright by Tatiana Smirnova 2011

Organogenesis in vitro under Altered Auxin Signaling Conditions

Tatiana Smirnova

Master of Science

Graduate Department of Cell and System Biology University of Toronto

2011

Abstract

The ratio of auxin to cytokinin determines de novo organogenesis in plants. Relatively little is

known about the effect of genetically altered auxin signaling on in vitro organogenesis. Here,

callusogenesis, shoot, and root formation were studied in loss- (LOF) and gain-of-function

(GOF) alleles in two phylogenetically related Auxin Response Factors (ARFs), MONOPTEROS

(MP/ARF5) and NON-PHOTOTROPHIC HYPOCOTYL 4 (NPH4/ARF7). Reduced MP activity

greatly diminished shoot regeneration, and partially diminished callusogenesis and root

formation. LOF in NPH4 strongly decreased callusogenesis, and mildly decreased shoot and root

regeneration in particular categories of explants. By contrast, organogenesis responses were

strongly increased in aerial explants carrying the GOF transgene dMP. Thus, both MP and NPH4

seem to act as positive regulators of certain organogenesis processes and the GOF dMP

transgene may be of interest for stimulating organogenesis in plant species with poor

regeneration properties. Also, organogenesis in vitro may reveal unknown developmental ARF

functions.

ii

Table of Contents Abstract……………………………………………………………………………………..……..ii

Table of Content…………………………………………………………………………..……...iii

List of Figures…………………………………………………………………………………....vii

List of Tables……………………………………………………………………………………..ix

List of Abbreviations……………………………………………………………………………...x

Chapter 1

Introduction………………………………………………………………………………..1

1.1 Plant hormone signals in plant regeneration………………………………………………1

1.1.1 The balance of auxin to cytokinin in a culture medium determines the type of

organogenesis……………………………………………………………………………...2

1.2 The auxin signal transduction pathway…………………………………………………...2

1.2.1 Biosynthesis, transport, and distribution of auxin in planta…...…………….........3

1.2.2 Auxin-mediated regulation of gene expression in Arabidopsis…………………...4

1.2.3 Aux/IAA genes and proteins……………………………………………………....4

1.2.4 ARF genes and proteins…………………………………………………………...7

1.2.5 Auxin response factors MONOPTEROS (MP/ARF5) and NON-PHOTOTROPIC

HYPOCOTYL 4 (NPH4/ARF7)…………………………………………………………..10

1.3 Roles of cytokinins.……………………………………………………………………...12

1.4 Cell proliferation and callusogenesis in plant tissue culture…………………………….13

1.5 De novo organogenesis in plant tissue culture …………………………………………..15

iii

1.5.1 Roles of shoot apical meristem genes during de novo shoot regeneration………18

1.5.2 Mutations affecting shoot regeneration in Arabidopsis…………….....................21

1.5.3 Mutations in auxin signaling genes affecting shoot regeneration……………......21

1.6 Regeneration of adventitious roots in vitro………………………………………………23

1.7 General Research Objectives………………………………………………………….....25

Chapter 2 Materials and Methods...........................................................................................27

2.1 Plant material………………………………………………………………………….....27

2.2 General growth conditions…………………………………………………………….....27

2.3 Hormone stocks………………………………………………………………. ………...30

2.4 Plant DNA extraction………………………………………………………………….....30

2.5 General procedure for polymerase chain reaction (PCR)……………… …………….....31

2.6 Genotyping…………………………………………………………………………….....31

2.7 Microtechniques and microscopy………………………………………………………..32

2.8 Plant tissue culture experiments……………………………………………………….....33

2.8.1 Rooting of dissected mpG12, dMP, nph4-1 and wild type seedlings……………33

2.8.2 Preparation of explants for tissue culture………………………………………...35

2.8.3 Preparation of tissue culture media………………………………………………36

Chapter 3 Callusogenesis under conditions of altered auxin signal transduction…………..37

3.1 Background and rationale…………………………………………………………….....37

3.2 Specific research objectives……………………………………………………………...38

iv

3.3 Results……………………………………………………………………………………39

3.3.1 Types of calluses on CIM………………………………………………………..39

3.3.2 Callusogenesis of cotyledon explants after 12 days on CIM…………………….42

3.3.3 Callusogenesis of leaf explants after 12 days on CIM……………………….......42

3.3.4 Callusogenesis of petiole explants after 12 days on CIM……………………......43

3.3.5 Callusogenesis of hypocotyl explants after 12 days on CIM…………………….45

3.3.6 Callusogenesis of adventitious root explants after 12 days on CIM…………......45

3.3.7 Callus proliferation in aerial explants after 6 days on CIM……………………...50

3.4 Discussion and Conclusions……………………………………………………………..51

Chapter 4 Shoot regeneration under conditions of altered auxin signal transduction………54

4.1 Background and rationale………………………………………………………………..54


4.3 Results……………………………………………………………………………………56

4.3.1 Shoot regeneration in cotyledon derived calluses………………………………..56

4.3.2 Shoot regeneration in leaf derived calluses………………………………………56

4.3.3 Shoot regeneration in calluses derived from petiole explants……………………59

4.3.4 Shoot regeneration in calluses derived from hypocotyl explants……………......61

4.3.5 Shoot regeneration in calluses derived from adventitious root explants………...62

4.3.6. Effect of duration of CIM preculture on shoot regeneration on SIM in calluses

derived from adventitious root explants………………………………………………….64

4.3.7 Greening of calluses derived from adventitious root explants on SIM…… …….68

v

4.4. Discussion and Conclusions………………………………………..…………………...71

Chapter 5 Root regeneration under conditions of altered auxin signal transduction………..74

5.1 Background and rationale………………………………………………………………..74


5.3 Results……………………………………………………………………………………76

5.3.1 Root regeneration in cotyledon derived calluses on RIM……………………......76

5.3.2 Root regeneration in leaf derived calluses on RIM………………………………80

5.3.3 Root regeneration in calluses derived from adventitious roots on RIM…………80

5.3.4 Root formation in adventitious root explants on SIM…………………………...82

5.4 Discussion and Conclusions……………………………………………………………..86

Chapter 6 General Discussion and Conclusions………………………………………….....89

6.1 Effects of changed MP/ARF5 and NPH4/ARF7 levels on callusogenesis, de novo shoot

and root regeneration……………………………………………………………..……...89

6.1.1 NPH4/ARF7 in auxin and cytokinin mediated developmental processes in tissue

culture…………………………………………………………………………………....89

6.1.2 MP/ARF5 in auxin and cytokinin mediated developmental processes…………..92

6.1.3 Enhancement of de novo organogenesis in the dMP genotype………………......94

6.2 General Conclusion………………………………………………………………………94

References………………………………………………………………………………………..96

vi

List of Figures 1.1 Model for the regulation of auxin signaling by auxin levels and auxin-induced

Aux/IAA protein degradation……………………………………………………………..5

1.2 MP/ARF5 and dMP/dARF5 protein structures and functions…………………………….8

1.3 Phases of organogenesis in Arabidopsis tissue culture…………………………………..17

2.1 Seedlings and inflorescence phenotypes in LOF mutants nph4-1, mpG12, and GOF

transgene line dMP……………………….………………………………………………29

2.2 Scheme of the tissue culture experiments and scored traits…………………………...…34

3.1 Fresh weight of calluses and frequency of the “whole” type of calluses after 12 days on

CIM……………………………………………………………………………………....40

3.2 Callus initiation and callus proliferation in cotyledon and leaf explants after 6 and 12

days on CIM……………………………………………………………………………...41

3.3 Callus initiation and callus proliferation in petiole and hypocotyl explants after 6 and 12

days on CIM……………………………………………………………………………...44

3.4 Callus initiation and callus proliferation in adventitious root explants

after 6 and 12 days on CIM………………………………………………………………46

3.5 Cotyledon and leaf explants after 12 days on CIM……………………………………....47

3.6 Petiole and hypocotyl explants after 12 days on CIM…………………………………...48

3.7 Adventitious root explants after 12 days on CIM………………………………………..49

4.1 Frequency of shoot regenerating calluses, and mean number of shoots per shoot

regenerating callus after 21 days on SIM………………………………………………...57

4.2 Calluses derived from cotyledon and leaf explants after 21 days on SIM……………….58

4.3 Calluses derived from petiole and hypocotyl explants after 21 days on SIM…………....60

vii

4.4 Calluses derived from adventitious root explants after 21 days on SIM………………...63

4.5 Frequency of shoot regenerating calluses and mean number of shoots per shoot

regenerating callus in calluses derived from adventitious root explants after 21 days

on SIM depending on the time on CIM (6, 9, 12 and 16 days)…………………………..65

4.6 Greening of calluses and root outgrowth in calluses derived from adventitious

root explants on SIM after 8, and 13 days on SIM………………………………………69

4.7 Frequency of green versus whitish calluses after 8 days on SIM in adventitious

root derived calluses ……………………………………………………………………..70

5.1 Frequency of root forming calluses, and the mean numbers of roots per root

forming callus in calluses derived from cotyledon, leaf and adventitious root

explants after 21 days on RIM…………………………………………………………...77

5.2 Cotyledon and leaf derived calluses after 21 days on RIM. …………………………….78

5.3 Root initiation in cotyledon explants after 6 days on RIM……………………………....79

5.4 Calluses derived from adventitious root explants after 21 days on RIM………………...81

5.5 Calluses derived from adventitious root explants after 12 and 21 days on RIM...............83

5.6 Frequency of root regenerating calluses, and the mean numbers of roots per

root regenerating callus in calluses derived from adventitious root explants after

21 days on SIM…………………………………………………………………………..84

5.7 Root outgrowth in calluses derived from adventitious root explants after 8 and 13

days on SIM……………………………………………………………………………...85

viii

List of Tables 2.1 Main characteristics of the mutant genotypes, mpG12, dMP,

and nph4-1…………………………………………………………………...…………………..28

2.2 Types of explants, genotypes, approximate sizes, and DAG at the moment

of dissection and introduction in tissue culture…………………………………………..35

2.3 The hormone composition of tissue culture media: CIM, SIM, and RIM……………….36

ix

List of Abbreviations 2,4-D 2,4-dichlorophenoxyacetic acid

2-iP gamma, gamma-Dimethylallylamino purine

A adenine

ARF AUXIN RESPONSE FACTOR

AuxRE auxin response element

B5 Gamborg B5 media

Bar Basta resistance gene

Basta glufosinate-ammonium

bp base pair

C cytosine

°C degrees Centigrade

CIM callus initiation medium

cm centimeter

Col-0 Columbia-0

DAG days after germination

DMSO methylsulfoxide

DNA deoxyribonucleic acid

dNTP deoxyribonucleotide triphosphate

E Einstein

EDTA ethylenediaminetetraacetic acid

f forward

g gram

G guanine

GFP GREEN FLUORESCENT PROTEIN

GOF gain-of-function

GUS β-glucuronidase

HCL hydrochloric acid

IAA 3-indoleacetic acid

IBA indole-3-butyric acid

KOH potassium hydroxide

x

xi

L litre

LB left border

LOF loss-of-function

m meter

M molar

min minute

MES 2-(4-Morpholino)ethanesulfonic acid

MP MONOPTEROS

MS Murashige and Skoog media

NaCL sodium chloride

NPH4 NON-PHOTOTROPIC HYPOCOTYL 4

dNTP deoxyribonucleotide triphosphate

ORF open reading frame

PCR polymerase chain reaction

r reverse

RB right border

RFLP restriction fragment length polymorphism

RIM root initiation medium

RNA ribonucleic acid

rpm revolutions per minute

RT room temperature

s second

SAM shoot apical meristem

SE standard error of the mean

SDS sodium dodecyl sulfate

SIM shoot initiation medium

T thymine

Tris Tris (hydroxymethyl) aminomethane

UTR untranslated region

v/v volume per volume

w/v weight per volume

wt wild type

Chapter 1

Introduction

1.1 Plant hormone signals in plant regeneration

Plant tissue culture and molecular genetics are core techniques for the genetic engineering of

plants (Li and Gray, 2005). The methodology is based on the unique ability of plants to

regenerate into complete plants from their parts (organs, tissues, or cells) (Schwarz et al., 2005).

Plant tissue culture is also an increasingly important tool for studies in plant development,

especially when integrated with techniques of molecular biology and genetics (Zuo et al., 2002;

Zhang and Lemaux, 2004; Caponetti et al., 2005; Gray, 2005; Mordhorst et al., 2005; Zhang and

Lemaux, 2005).

In 1957, Skoog and Miller demonstrated that the ratio of auxin to cytokinin in a culture medium

determines the outcome of developmental processes in tissue culture, and results in callus, shoot,

or root formation (Skoog and Miller, 1957). During the last two decades, the main foci of tissue

culture research were on cytokine signaling and shoot meristem genes, in light of their effect on

de novo shoot meristem formation (Howell, 2003; Zhang and Lemaux, 2004; Zhang and

Lemaux, 2005). However, auxin response genes may also play a crucial role in organogenesis

and shoot regeneration processes in vitro, given their key role in most developmental processes

in planta (Berleth et al., 2004; Teale et al., 2006), and given the requirement for auxin and

cytokinin balance in the culture medium in vitro (Skoog and Miller, 1957). Thus, the study of

auxin response genes may provide insight into the molecular basis of de novo organogenesis in

tissue culture.

Auxin signaling operates through a family of transcription factors called auxin response factors

(ARFs), which regulate expression of the primary auxin responsive genes (Berleth et al., 2004;

Teale et al., 2006). Two ARFs with partially overlapping functions in embryo shoot meristem

patterning, ARF5 (MONOPTEROS) (MP/ARF5) and ARF7 (NONPHOTOTROPHIC

HYPOCOTYL 4) (NPH4/ARF7) (Hardtke et al., 2004), have been tested in this research to

determine their roles during callus formation and organogenesis in vitro.

1

1.1.1 The balance of auxin to cytokinin in a culture medium determines the

type of organogenesis

Various types of developmental responses and organogenesis in vitro can be triggered by

particular ratios of auxin and cytokinin in a culture medium (Skoog and Miller, 1957). Generally,

high cytokinin to auxin levels promote de novo shoot regeneration; while auxin alone, or auxin in

combination with low cytokinin levels, triggers de novo root formation. An intermediate ratio of

auxin to cytokinin is required for callus proliferation (Skoog and Miller, 1957; Gaba, 2005).

However, it is still unclear, how the ratios of these hormones direct the variations of

organogenesis responses (Howell, 2003; Zhang and Lemaux, 2004; Zhang and Lemaux, 2005).

Yet, it is clear that a molecular understanding of the signal transduction pathways of both

hormones will aid in the research on plant regeneration properties.

1.2 The auxin signal transduction pathway

Physiological responses which are directed by auxin are central to plant structure and

functioning. The fate of developing tissues may be determined by their sensitivity to auxin, to the

concentration of active auxin, and to the relative concentrations of other phytohormones

(reviewed in Teale et al., 2006). Natural auxin (mainly indole-3-acetic acid, IAA) is a key

regulator of remarkably diverse developmental processes in plants, including root initiation,

embryo and fruit development, organogenesis, vascular tissue differentiation, root and embryo

patterning, photo- and gravitropism, apical dominance, and phyllotaxy. On the cellular level,

auxin mediates cell division, elongation, and differentiation, although how exactly it is involved

in each process is not completely understood (reviewed in Hagen and Guilfoyle, 2002; Berleth et

al., 2004; Davies, 2004; Paciorek and Friml, 2006; Teale et al., 2006).

Auxin plays a central role within the plant transduction network, frequently acting in conjunction

with other signals and signaling transduction pathways to regulate complex developmental

processes. There is evidences of auxin ‘cross-talk’ with other pathways: cytokinin, ethylene,

ABA (abscisic acid), GA (gibberellic acid), brassinosteroids, jasmonic acid, and with the light

signaling system (reviewed in Swarup et al., 2002). That is why many “auxin-response” mutants

exhibit pleiotropic signaling defects: they are a reflection of the existence of wide ‘cross-talk’

among signaling pathways (Hobbie and Estelle, 1994; Swarup et al., 2002).

2

The complex and diverse processes mediated by auxin operate through the signaling cascades

that recruit specific transcriptional factors to regulate executor genes, the ones which carry out

the required responses. The specificity of the processes is provided by the diversity of the

transcription factors and by the auxin distribution. Both the auxin distribution and the expression

profiles of certain classes of transcription factors contribute to the specificity of various auxin

responses (Vogler and Kuhlemeier, 2003).

1.2.1 Biosynthesis, transport, and distribution of auxin in planta

Auxin is actively transported basally from the sites of its biosynthesis in the aerial parts to the

roots (Davis, 2004). According to Ljung et al. (2001), all plant organs can synthesize auxin, with

the highest rate of auxin synthesis in young leaves. Patterns of auxin distribution are not only

dependent on auxin synthesis and catabolism, but also on auxin transport, processes which in

turn are regulated through the cells’ capacity for auxin influx and efflux. Auxin active transport

and redistribution involve many proteins. Among them, the family of polarly localized plasma-

membrane proteins, PIN-FORMED (PIN) proteins, which mediate auxin efflux (Petrasek et al.,

2006; Teale et al., 2006; Vieten et al., 2007; Vanneste and Friml, 2009), is intensely studied.

Several other protein families involved in auxin transport are known, among them, the presumed

influx carrier (AUX1), and two more proteins mediating auxin transport (reviewed in Leyser,

2006; Vieten et al., 2007; Vanneste and Friml, 2009).

Several reporters were created to visualize the spatial patterns of auxin responses as an indication

for the distribution of auxin. Among them, the DR5 reporter is the one most commonly used

(Ulmasov et al., 1997; Benkova et al., 2003). DR5 is a composite, synthetic auxin responsive

element (AuxRE) which consists of 5-7 tandem repeats of the auxin-responsive TGTCTC element

(Ulmasov et al., 1997). To visualize auxin responses, the DR5 element can be fused to either the

β-glucurunidase (GUS) reporter gene (Ulmasov et al., 1997), or to the green fluorescent protein

(GFP) gene sequence (Friml et al., 2003). Routs of auxin transport are typically monitored

through a translational fusion of the PIN1 auxin efflux carrier protein to GFP (PIN1::PIN1-GFP

transgene construct) (Benkova et al., 2003; Friml et al., 2003; Heisler et al., 2005; Scarpella et

al., 2006; Gordon et al., 2007).

3

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Several reporters were created to visualize auxin distribution and routes of auxin flow. One of

the most commonly used reporters to monitor sites of auxin accumulation is DR5::GUS/GFP, a

composite of 5-7 repeats of the auxin response element (AuxRe) TGTCTC, found in promoters

of auxin response genes (Ulmasov et al., 1997).

1.2.2 Auxin-mediated regulation of gene expression in Arabidopsis

Active auxin transport by PIN proteins influences the cellular auxin concentration that then

triggers the transcriptional control of the expression of auxin dependent genes. The gene

regulatory auxin responses in Arabidopsis are mediated by two major protein families: Aux/IAA

proteins, and auxin response factors (ARFs) (reviewed in Berleth et al., 2004; Teale, 2006;

Guilfoyle and Hagen, 2007; Vanneste and Friml, 2009). A key feature of the auxin response

system is the ubiquitin-mediated proteolysis of Aux/IAA proteins (reviewed in Berleth et al.,

2004; Gray 2004; Weijers et al., 2005; Vanneste and Friml, 2009). A summary of auxin

signaling is represented in Fig. 1.1.

1.2.3 Aux/IAA genes and proteins

The 29 members of the Aux/IAA protein family are small, short-lived nuclear proteins, which

are transcriptional regulators of early auxin-response genes. Aux/IAA proteins are encoded by

the primary auxin response genes. Aux/IAA proteins consist of four domains (I-IV). Domain I

repress the transcription factors to which it becomes attached. Domain II contains a “degron”

sequence, through which AUX/IAA proteins are targeted towards the SCF/TIR ubiquitin ligase.

Interaction between AUX/IAA proteins and SCF/TIR is enhanced by auxin. Through this

mechanism, auxin promotes the ubiquination and degradation of Aux/IAA proteins (Gray et al.,

2001; Berleth et al., 2004; Vanneste and Friml, 2009). Domains III and IV play an important role

in homo- and hetero-dimerization with other Aux/IAA proteins and ARFs (Ulmasov et al.,

1999b; Berleth et al., 2004; Vanneste and Friml, 2009). The functions of Aux/IAA dimers are

not clear (Paciorek and Friml, 2006). Heterodimerization of Aux/IAA proteins with ARFs results

in the ARFs’ repression (Ulmasov et al., 1999b; Berleth et al., 2004; Vanneste and Friml, 2009).

When the concentration of auxin in a cell is low, Aux/IAA proteins are available to bind to the

corresponding ARFs, resulting in the inhibition of ARF. When the auxin concentration in a cell

increases, the Aux/IAA proteins are targeted for degradation, which results in the ARFs’ de-

4

Figure 1.1 Model for the regulation of auxin signaling by auxin levels and auxin-induced Aux/IAA protein degradation

Low auxin levels result in the binding of the Aux/IAA proteins to ARFs via shared dimerization domains III and IV (shown in yellow and green in both Aux/IAA and ARF) and a repression of ARF function. The repression of ARFs is mediated via domain I (red) of Aux/IAA protein. The ARF proteins are bound to the promoter part of the target genes via their DNA binding domain (DBD, in purple).

Increased auxin levels promote Aux/IAA protein degradation and de-repression of ARF protein function. Auxin-induced Aux/IAA protein degradation occurs via the ubiquitin-26S proteasome protein degradation pathway (on the right). Ubiquitin molecules are conjugated to the Aux/IAA protein through the concerted action of three enzymes, ubiquitin-activating enzyme (E1), ubiquitin-conjugated enzyme (E2), and ubiquitin protein ligase (E3). The SCF-type ligases represent the largest family of E3 enzymes in Arabidopsis. They consist of four subunits, three of which, SCP1-like protein (ASK1/2 here), Cullin (CUL1/AXR6 here), and an F-box protein (the TIR1 protein here) gave the name of this E3 complex – SCF or SCFTIR1. The TIR1 protein acts as an auxin receptor. The binding of auxin to TIR1 promotes substrate-specific binding and enzymatic activity of TIR1 to the substrate Aux/IAA protein, and its ubiquitination and degradation. The conserved domain II (in blue) of Aux/IAA proteins is necessary for the interaction with the TIR1 subunit, and the mutation in this domain results in increased stability of the Aux/IAA protein and auxin resistance.

Disassociation of Aux/IAA from ARF allows for the dimerization of ARFs via their shared domains III and IV, resulting in further potentiation of ARF activity.

(Figure modified from Berleth et al., 2004; the legend based on Berleth et al., 2004; Hagen, 2004, Kepinski et al., 2004, Dharmasiri et al., 2005, Krogan, 2006).

5

IAA

REPRESSON

ACTIVATION

POTENTIATION

LOW AUXIN

HIGH AUXIN

E1

E2

E3 SCF complex

TIR1

6

repression and their activation as transcriptional factors. As long as the auxin concentration

remains high, Aux/IAA proteins turn over at a high rate due to the targeted proteolysis (Hagen et

al., 2004). The rapid, auxin-dependent turnover of Aux/IAA proteins makes them highly

responsive to changes in auxin signaling (Gray and Estelle, 2000).

Some AUX/IAA protein interactions with ARFs have been investigated. IAA28 interacts with

ARF5, 6, 7, 8 and 19 (Rybel et al., 2010); IAA14/SLR and IAA13/SHY2 interact with

NPH4/ARF7 and ARF19 (Peret et al., 2009, and references therein). IAA12/BDL interact with

MP/ARF5 (Hamann et al., 2002; Weijers et al., 2005; Rybel et al., 2010). Gain-of-function

(GOF) mutations in AUX/IAA proteins typically affect the “degron” in domain II and lead to

stable AUX/IAA, whose abundance is no longer affected by auxin (Gray et al., 2001; Weijers et

al., 2005). GOF mutants of iaa12/bdl, and, to a lesser degree, with constitutively repressed

MP/ARF5 function, cause phenotypes similar to those found in mp mutants (Hamann et al.,

2002; Hardtke et al., 2004; Weijers et al., 2005). GOF mutations in iaa28 and iaa14/slr cause a

lack of lateral roots, similar to that found in nph4 arf19 double mutants (Rogg et al., 2001;

Okushima et al., 2005; Peret et al., 2009; Rybel et al., 2010).

1.2.4 ARF genes and proteins

ARFs are transcription factors encoded by a gene family of 23 members. ARFs are characterized

by a conservative amino-terminal DNA binding domain (DBD), which interacts directly with the

auxin response element (AuxRE). AuxREs (TGTCTC) are found in the promoters of most

primary auxin-response genes. ARFs comprise another, only moderately conserved domain,

called the middle domain, which determines the functional properties of an ARF as a

transcriptional activator or repressor. Carboxy-terminal domains III and IV are similar to those in

AUX/IAA proteins, and mediate dimerization with Aux/IAAs or other ARFs (Ulmasov et al.,

1999a, Ulmasov et al., 1999b; Berleth et al., 2004). Recently Shin et al. (2007) showed that ARF

activity can also be modulated by interaction with other factors, such as MYB77, which increase

the combinatorial possibilities in auxin-dependent transcriptional regulation.

Whether through differential expression, differential affinities for target promoters, or both,

individual ARFs have a distinct function in developmental processes (Vogler and Kuhlemeier,

7

Figure 1.2 MP/ARF5 and dMP protein structures and functions:

A) When the auxin concentration is low, the Aux/IAA protein, BDL/IAA12, binds to MP/ARF5

through their shared dimerization domains III and IV. As a result, MP/ARF5 does not function as

a transcription activator and the expression of early auxin response genes downstream of

MP/ARF5, is repressed.

B) The proposed model suggests that, when the dimerization domains III and IV are truncated, as

in the dMP protein, no binding of the Aux/IAA protein occurs. As a result, dMP functions

constitutively as a transcriptional activator regardless of auxin levels.

AuxRE – auxin response element; DBD (purple) – DNA binding domain; MR (light blue) –

middle region; III (yellow) and IV (green) – the shared dimerization domains in MP/ARF5 and

BDL/IAA12; I (red) and II (blue) – domains I and II in BDL/IAA12 protein.

8

Inhibition of the MP/ARF5 function and expression of early auxin gene by the binding of Aux/IAA protein

AuxRE

I IMP/ARF5

Aux/IAA12

MP/ARF5

Constitutive dMP/dARF5 function and early auxin gene expression when Aux/IAA protein is not able to bind to dMP/dARF5

AuxRE

DBDDBD MRMR

dMP

IVI I II II

Aux/IAA12

I

DBDDBD MRMR IVIVI I II I I

dMPa) b)

Low/high auxin

III IV

Low auxin

9

2003). However, the precise functions and downstream gene targets of most ARFs are still not

known (Okushima et al., 2005; Guilfoyle and Hagen, 2007).

A systematic reverse genetics approach with ARF genes failed to reveal an abnormal phenotype

of most ARF mutations, which suggested that most of the ARFs have largely redundant functions

(Okushima et al., 2005; Teale, 2006). The analyses of ARF1 to ARF10 have shown that their

mRNAs are ubiquitously expressed in most major organs, as well as in cultured undifferentiated

Arabidopsis cells (Ulmasov et al., 1999b). Participation in multiple transduction pathways was

confirmed for several ARFs. For example, ARF19 participates in both auxin and ethylene

signaling (Li et al., 2006), and ARF6 and ARF8 promote jasmonic acid production (Nagpal et

al., 2005). Unlike Aux/IAA genes, ARF genes do not belong to the group of early auxin response

genes. ARFs have been found to be permanently expressed and bound to promoters of the target

genes containing the AuxRE sequence. ARF functions are activated when the auxin

concentration increases and Aux/IAA proteins dissociate from ARFs, allowing a homo- or

hetero-dimerization of ARFs with each other (Berleth et al., 2004; Hagen et al., 2004).

Interestingly, there is a growing body of information on posttranscriptional regulation of ARFs’

mRNA levels by micro-RNAs, and by small interfering RNAs. This posttranscriptional

regulation of ARFs seems to be as important as their transcriptional regulation (Guilfoyle and

Hagen, 2007).

1.2.5 Auxin response factors MONOPTEROS (MP/ARF5) and NON-

PHOTOTROPIC HYPOCOTYL 4 (NPH4/ARF7)

Loss-of-function mutations have been identified by single mutant phenotype in only 4 of 23

ARFs, arf2/hss, arf3/ett, arf5/mp, and arf7/nph4 (Okushima et al., 2005). They include those in

MONOPTEROS (MP/ARF5) and NON-PHOTOTROPIC HYPOCOTYL 4 (NPH4/ARF7). Both

belong to the group of transcriptional activators with corresponding middle domains (Ulmasov et

al., 1999a; Ulmasov et al., 1999b). Both ARFs, MP and NPH4, are capable of homo- and hetero-

dimerization with each other (Hardtke et al., 2004). Especially in their DNA binding domains,

MP and NPH4 have a high degree of sequence similarity, and they have partially redundant

functions (Liscum and Reed, 2002; Hardtke et al., 2004; Remington et al., 2004; De Smet et al.,

2010).

10

Loss of function (LOF) mutations in MP interfere with embryonic patterning, and with the

formation of embryonic hypocotyl and primary roots (Mayer et al., 1991; Berleth and Jurgens,

1993). LOF mutations in MP also severely reduce auxin sensitivity, auxin transport, and vascular

development (Li et al., 2006; Przemeck et al., 1996; Mattsson et al., 2003; Schuetz et al., 2008).

Finally, they result in the formation of “pin-like” inflorescences, which lack leaves and flowers

(Berleth and Jurgens, 1993; Przemeck et al., 1996). The expression of PIN1 (auxin transport) and

DR5 (auxin response) reporters is defective in mp mutant leaves, suggesting that these responses

were diminished in mp mutants (Mattsson et al., 2003; Wenzel et al., 2007).

LOF mutations in NPH4 lead to hypocotyls that no longer bend towards the light, as well to

changes in leaf shapes and in other aspects of post-embryonic morphology (Stowe-Evans et al.,

1998). LOF nph4 mutants exhibit auxin resistance, and severely impaired expression of many

primary auxin-dependent genes (Okushima et al., 2005). Expression of the auxin-responsive

reporter DR5 was also significantly reduced in nph4 mutants (Stowe-Evans et al., 1998; Wang et

al., 2005). Also, Watahiki and Yamamoto (1997) demonstrated that nph4 mutants showed a 15 to

20 fold increase in auxin resistance (2,4-D) in the aerial parts, whereas the root parts were of

normal sensitivity to auxin (Watahiki and Yamamoto, 1997; Okushima et al., 2005). In the

presence of exogenous IAA, the excised nph4 roots formed fewer lateral roots, and the nph4

excised hypocotyls formed fewer adventitious roots than did wild-type roots and hypocotyls

(Wilmoth et al., 2005). However, almost no adventitious or lateral roots were formed in nph4

arf19 double mutants (Okushima et al., 2005; Wilmoth et al., 2005).

The developmental functions of MP and NPH4 are far more related than is suggested by their

single mutant phenotypes. Both MP/ARF5 and NPH4/ARF7 can act in the regulation of

embryonic pattern formation (Hardtke et al., 2004). Although the single mutant phenotype of

NPH4 implicates the gene only in conditional responses during post-embryonic development,

NPH4 is already expressed in embryos, and LOF mutations in NPH4 dramatically enhance the

mp embryo phenotype (Hardtke et al., 2004).

MP is expressed in broad domains in embryos and in emerging organs. The early functions of the

MP gene are related to vascular and body patterning, and to organ initiation (Hardtke and

Berleth, 1998; Hardtke et al., 2004; Schuetz et al., 2008; Cole et al., 2009; De Smet et al., 2010).

However, upon organ maturation, the expression of MP is gradually narrowed to the central

11

domains, and then to the vascular tissues (Hardtke et al., 2004). By contrast, NPH4 is fairly

ubiquitously expressed throughout plant development (Hardtke et al., 2004; Okushima et al.,

2005).

Selective deletion of domains III and IV of MP results in a GOF version of MP that is associated

with the dominant enhancement of auxin-related traits. Therefore, it was named dMP (for

dominant MONOPTEROS) (Krogan et al., 2006). Those traits included increased adventitious

root formation, when dMP cotyledons were rooted in a liquid medium containing IBA (Krogan

et al., 2006). By contrast, adventitious root formation in LOF mp cotyledons was strongly

decreased (Mattsson et al., 2003). Also, MP together with PIN1 is involved in the process of

embryonic shoot meristem formation and patterning, through control of the expression of the

meristem genes CUC2 and STM (Aida et al., 2002). It is also involved in the regulation of

meristem size (Schuetz et al., 2008); and in the regulation of the activity of some A-type

ARABIDOPSIS RESPONSE REGULATORS (A-type ARRs, negative cytokinin response

regulators) in SAM (Zhao et al., 2010). The MP/ARF5 protein interacts with BODENLOS

(IAA12/BDL) resulting in inhibition of the MP/ARF5 function (Fig.1.2 A; Hardtke et al., 2004;

Weijers et al., 2005). However, dMP is partially epistatic over bdl (Krogan et al., 2006).

1.3 Roles of cytokinins

Cytokinins were discovered by their ability to stimulate plant cell division (del Pozo et al., 2005,

and references therein). Later, Skoog and Miller introduced their hypothesis, according to which

plant morphogenesis is under the control of the auxin to cytokinin ratio (Skoog and Miller,

1957). Cytokinins control key aspects of plant growth and development, such as cell division,

shoot meristem initiation, control of the stem cell pool in the shoot apical meristem, leaf and root

differentiation, vascular patterning, chloroplast biogenesis, photomorphogenesis, fertility, seed

development, stress tolerance, and senescence (reviewed in Muller and Sheen, 2007a; Muller and

Sheen, 2007b). Together with auxin, cytokinin can reprogram terminally differentiated cells to

stem cells, and initiate de novo shoot formation in culture (Gaba, 2005; Zhang and Lemaux,

2005). The most common cytokinin in plants is zeatin (Davies, 2004). Although the important

sites of cytokinin biosynthesis seem to be in the root meristems (Davies, 2004; Nordstrom et al.,

2004), it has been shown recently that cytokinin biosynthesis occurs in certain quantities in all

types of aerial organs. The major subcellular compartments of cytokinin biosynthesis are

12

plastids. The major steps in cytokinin signaling are reviewed in detail in Maxwell and Kieber

(2004), and in Muller and Sheen (2007 a, b).

Many studies were done to illustrate the interaction of auxin and cytokinin signaling pathways,

and their mutual control of growth and differentiation in plants (Coenen and Lomax, 1997;

Nordstrom et al., 2004; Moubayidin et al., 2009; Zhao et al., 2010). The existence of complex

and multilevel networks of synergistic, antagonistic, and additive interactions between these two

plant hormones and their signaling systems have been demonstrated in a number of studies (Klee

and Estelle, 1991; Hobbie and Estelle, 1994; Coenen and Lomax, 1997; Nordstrom et al., 2004;

Moubayidin et al., 2009; Zhao et al., 2010). One of the indicators of interaction, between auxin

and cytokinin signaling pathways, is the number of mutants cross resistant to both hormones

(Hobbie and Estelle, 1994; Hobbie et al., 1994; Coenen and Lomax, 1997; Swarup et al., 2002;

Moubayidin et al., 2009). Auxin signaling mutants, such as aux1, axr1, axr2, axr3, iaa28, and

shy2, have demonstrated pleiotropic signaling defects affecting other signaling pathways,

including cytokinin signaling (Hobbie and Estelle, 1994; Swarup et al., 2002). Also, several

mutants, initially characterized as affected in cytokinin perception, have been shown to have

altered auxin responses (Smalle et al., 2002; Tajima et al., 2004; Laxmi et al., 2006). The

molecular basis of auxin-cytokinin cross talk is largely unclear (Gray, 2004; Hartig and Beck,

2006), although important advances in its understanding have been recently made (reviewed in

Moubayidin et al., 2009).

1.4 Cell proliferation and callusogenesis in plant tissue culture

Plant callusogenesis is a process also observed under natural growth conditions, where two types

of triggers can lead to the development of callus tissues – tumor formation in response to

infection by Agrobacterium tumefaciens, and wounding with subsequent regeneration of

damaged tissues (Frank et al., 2000; Gordon et al., 2007). Callus proliferation can be induced in

vitro, when the culture medium contains a certain ratio of auxin to cytokinin (Skoog and Miller,

1957). Cell proliferation during callusogenesis leads to disruption of the ordered morphology of

differentiated tissues (Cary et al., 2002).

The mechanisms leading to callus proliferation are not well understood (Cary et al., 2002; Zhang

and Lemaux, 2005). Generally, cell-cycle genes, such as cyclins and cyclin dependent kinases

(CDKs), play a major role in the mediation of hormonal effects, or in response to wounding

13

signals, leading to cell proliferation (Hemerly et al., 1993; Soni et al., 1995; Shaul et al., 1996;

Riou-Khamlichi et al., 1999; Sugiyama, 1999; Richard et al., 2002; Che et al., 2007). It has been

shown that the level of CDK expression correlates with competence for cell proliferation

(Hemerly et al., 1993; Zhang et al., 2005); and, CDK expression and cell proliferation are

increased upon wounding in tissue culture (Hemerly et al., 1993).

Callus initiation, callus growth on callus inducing medium (CIM), and subsequent shoot

formation on SIM were completely prevented in the Arabidopsis root explants cultured during

the first 1 to 2 days on the CIM supplied with reversible cell cycle inhibitors aphidicolin (APH)

or hydroxyurea (HU), which block G1-S transition (Che et al., 2007). Overexpression of cyclin

D3 (CycD3) resulted in cytokinin-independent callus growth, but these calluses failed to

regenerate shoots (Riou-Khamlichi et al., 1999). Richard et al. (2002) found that, during the

initiation of a cell suspension culture, the different members of the CDK or CYC families were

differentially up-regulated by mitogenic factors, including auxin and cytokinin. Moreover, cis-

acting regulatory elements specific for particular plant hormones, such as AuxREs for auxin,

were found in the promoters of the cell cycle genes (Richard et al., 2002). Auxin responsive

elements (AuxREs) were found in the promoter regions of many cyclin genes, indicating that

they might be primary auxin responsive genes (Perrot-Rechenmann, 2010).

Gordon et al. (2007) studied the dynamics of the auxin and cytokinin responses, and the pattern

of the distribution of hormonal responses during developmental processes in tissue culture.

According to their data, auxin responses, visualized by DR5 and PIN1 driven reporters, were

strongly increased only during the initiation of callus proliferation on CIM, but they were

downregulated after 5 days on CIM. They were not observed within the large callus outgrowths

after 1 week on CIM, and were not detected after 10 days on CIM. Conversely, the expression of

the cytokinin signaling marker pARR5::GFP, and of CUP-SHAPED COTYLEDON 1 and 2

(CUC1 and CUC2), were detected in an already established culture after 8 days on CIM. The

signal expanded throughout the callus after 2 weeks of induction (Gordon et al., 2007). No

expression of WUSCHEL (WUS), SHOOT MERISTEMLESS (STM), CLAVATA3 (CLV3),

FILAMENTOUS FLOWER (FIL), or REVOLUTA (REV) reporters was detected during 2 weeks

on CIM, indicating that expression of these key SAM genes require incubation on SIM (Cary et

al., 2002; Gordon et al., 2007). Interestingly, both the CUC2 and ARR5 reporters were expressed

at the site of callus formation, even on the medium lacking cytokinin, and containing auxin as the

14

sole hormone. By contrast, the culture of explants with the hormone kinetin as the sole hormone

did not lead to callus proliferation and CUC2 reporter expression, suggesting that auxin

containing medium is required for callus proliferation and the CUC2 gene expression (Gordon

et al., 2007).

Global analysis of gene expression during the first 4 days of callus proliferation on CIM

demonstrated that more than 250 genes were significantly upregulated, and about 500 genes were

downregulated, during the early stages of callus formation, usually associated with the stage of

acquisition of competence for organogenesis (Che et al., 2006). A dramatic increase in the

expression of genes involved in hormone responses, particularly Aux/IAA genes, was detected on

the second day of CIM incubation. These genes were downregulated after the transfer of explants

on SIM (Che et al., 2002). When cells proliferated and formed callus tissue (after 10 days of

CIM incubation), the main categories of upregulated genes were stress-related transcription

factors and stress-related proteins (Che et al., 2006). However, the authors concluded that the

genes associated with the acquisition of competence, and those specific for callus formation,

were more difficult to categorize compared to those that are part of the characterization of gene

expression during shoot formation on shoot inducing medium (SIM), or root formation on root

inducing medium (RIM) (Che et al., 2006).

Based on the requirement of a high auxin concentration in CIM (Skoog and Miller, 1957),

mutations in auxin signaling most likely can interfere with callusogenesis processes. Thus,

explants of the auxin-resistant mutant axr1-3 required more auxin for callus induction, and

tolerated more auxin to grow calluses, while sensitivity to kinetin was normal in tissue culture

(Lincoln et al., 1990; Kubo and Kakimoto 2000; Sieberer et al., 2003). Another well known

auxin-resistant mutant, aux1, also required higher auxin concentrations for callus initiation, and

demonstrated a tendency to form roots rather than calluses on CIM (Kakani et al., 2009). Altered

auxin transport in pin1 mutants resulted in the proliferation of callus-like tissues, instead of

lateral roots, when exogenous auxin was applied to the roots (Benkova et al., 2003).

1.5 de novo organogenesis in plant tissue culture

Plant tissues can potentially develop into somatic embryos, shoots, and roots. The specificity of

the regeneration is determined by the hormone composition of the culture medium (Schwarz et

al., 2005, and references therein). Primordia originate de novo from callus cells. The cell, or

15

cells, that can be direct progenitors are stimulated to undergo a number of rapid cell divisions,

resulting in the formation of a meristemoid. A meristemoid represents an aggregate of small,

isodiametric, thin-walled, micro-vacuolated cells. Early in their development, meristemoids are

capable of developing into different types of organ primordia (Schwarz et al., 2005, and

references therein; Atta et al., 2009). The capabilities of plants for de novo shoot and root

organogenesis are among the most important qualities required for successful micropropagation

in vitro (Schwarz et al., 2005).

Organogenesis in vitro can be direct or indirect. In direct organogenesis, explants are fully

competent to respond to inductive hormones, and do not require a callus proliferation phase

(Schwarz et al., 2005). Recently, the capability of Arabidopsis thaliana roots and hypocotyl

explants for direct shoot organogenesis has been described (Atta et al., 2009). In general,

however, the explants of most plants, including Arabidopsis, can undergo organogenesis only

following a callus growth phase on CIM. During this phase, it is hypothesized that genetic or

developmental factors blocking the competence to respond to the inductive signals (such as

inducing shooting or rooting hormone ratio in tissue culture medium) are overcome. As a result,

many efforts to induce organogenesis focus on overcoming these blocks to the competence to

respond to the inductive signals (Cary et al., 2002, and references therein).

According to the most commonly accepted model (Christianson and Warnick, 1985; Schwarz et

al., 2005), regeneration occurs in three sequential phases (Fig. 1.3): first, when explants are

cultured on CIM, they acquire competence to respond to organogenesis stimuli during the

following stage (on SIM or RIM); second, the determination phase, shortly after calluses are

transferred either to SIM or RIM, when the type of organogenesis (shoot or root regeneration) is

determined, and, third, the organogenesis phase, during which meristems are forming, and

shoots/roots emerging. The third phase, the formation of organ primordia, and organ emergence,

is considered to be hormone independent (Christianson and Warnick, 1985). Ozawa et al. (1998)

argued that “competence for shoot regeneration is assumed to be always acquired additively,

over and above the competence for root redifferentiation”. Shown first in leaf explants of

Convolvulus arvensis (Christianson and Warnick, 1983, 1984, 1985, 1988), the organogenic

phases were widely confirmed for other plant species and cultivars, including pine, tobacco,

apple, Arabidopsis, mangosteen, etc. (Flinn et al., 1988; Attfield and Evans, 1991; Hicks, 1994;

16

Phase I“acquisition

of competence” on CIM

Phase II“determination”on SIM or RIM

Phase III“differentiation”,

hormone-independent

CIM:high auxin

low cytokinin

explants

SIM:high cytokinin

low auxin,shoots

RIM:auxin,roots

Figure 1.3 Phases of organogenesis in Arabidopsis tissue culture

According to the model of Christianson and Warnick (1985), de novo regeneration occurs in three sequential phases: the first phase, the acquisition of competence, i.e. of the competence to respond to the stimuli for organogenesis during dedifferentiation on callus inducing medium (CIM); the second phase, called the determination phase, where the type of organogenesis is determined depending on the type of medium; and, the organogenesis phase, when meristems are fully developing shoots or roots are emerging. The first phase, the acquisition of competence, occurs on auxin-rich CIM. The second, the determination of either shoots or roots, requires either a cytokinin-rich shoot initiation medium (SIM), or an auxin-containing root initiation medium (RIM). The third phase, the formation of organ primordia and organ emergence, is considered to be hormone independent.

(Figure modified from Howell et al., 2003)

17

de Klerk et al., 1995; Lakshmanan et al., 1997; Sugiyama, 1999; Ozawa et al., 1998; Banno et

al., 2001; Cary et al., 2002; Che et al., 2007).

During in vitro organogenesis, a high cytokinin level in SIM initiates and promotes the processes

of meristemoid specification, and the “canalization” of a developmental program for the

formation of shoot primordia (Schwarz et al., 2005). In Arabidopsis, the origin of the shoot

meristem in vitro has recently been shown to be initiated from dividing xylem pericycle cells, or

from lateral root meristem - like protuberances, which are formed during culture on CIM (Che et

al., 2007; Atta et al., 2009; Sugimoto et al., 2010). The pericycle cells remain more meristematic

than the surrounding cells, and retain the expression of cell cycle genes. In particular, the xylem

pericycle cells have a higher capacity to re-enter the cell cycle, and to form new meristems when

compared to the other cells (Atta et al., 2009).

Many authors have indicated that the ability for shoot regeneration differs dramatically among

Arabidopsis ecotypes. In particular, the Columbia (Col) ecotype, widely used in research, has a

low shoot regeneration potential (Akama et al., 1992; Cary et al., 2002; Zhao et al., 2002;

Chatfield and Raizada, 2008). As a rule, Arabidopsis ecotypes more capable of shoot

regeneration, such as C24, and Wasilewskija, need less time on SIM to be committed to

regeneration, and to form green foci and shoots (Cary et al., 2002; Zhao et al., 2002). The

regeneration potential also varies depending on the types of explants, and on explant age - partly

because of the different endogenous hormone concentrations (Akama et al., 1992; Zhao et al.,

2002; Gordon et al., 2007).

1.5.1 Roles of shoot apical meristem genes during de novo shoot

regeneration

Several key genes, many encoding transcription factors, are known to have critical roles in the

initiation and maintenance of a shoot apical meristem (SAM). Some are expressed as early as the

embryonic heart stage, and usually play crucial roles in post-embryonic meristem maintenance

(Aida et al., 1999; Long et al., 1996; Cary et al., 2002). The SHOOT MERISTEMLESS (STM)

gene of Arabidopsis encodes a specific, knotted-like 1 (KN1-like), type homeodomain protein

(Barton and Poethig, 1993; Zhang and Lemaux, 2005). The STM gene is required for SAM

formation and maintenance (Long et al., 1996). It promotes cell division, and suppresses cell

18

differentiation in the SAM (Lenhard et al., 2002). The WUSCHEL (WUS) gene is a member of

another subtype of homeodomain transcription factors, required for specifying and maintaining a

stem cell population in the SAM (Mayer et al., 1998; Zhang and Lemaux, 2005). The WUS and

STM genes are activated independently, but mutations in WUS results in the loss of expression of

STM and vice versa (Lenhard et al., 2002). The CLAVATA3 (CLV3) gene encodes a small

protein, which appears to be a ligand for CLV1 receptor kinase (Fletcher et al., 1999). CLV1 and

CLV3 restrict cell proliferation activity in SAM (Fletcher et al., 1999). The CLV3 gene is

activated by WUS, and functions in a negative feedback loop that antagonizes WUS activity, and

thereby controls the size of the central stem cell population (Gallois et al., 2004, and references

therein). The redundant genes CUP-SHAPED COTYLEDON 1 and 2 (CUC1 and CUC2) encode

transcription factors with similarities to the NO APICAL MERISTEM (NAC) protein. The latter

acts in the development of embryos and flowers in petunias (Aida et al., 1997, and references

therein; Takada et al., 2001). These types of proteins are known to play important roles in

establishing SAM and in separating cotyledons (Aida et al., 1997, and references therein; Takada

et al., 2001). Loss of function mutations in STM, WUS, or CUC1 and CUC 2 (in double mutants)

block SAM formation in planta (Cary et al., 2002; Hibara et al., 2003). All four genes are also

used as reporters of shoot meristem initiation in vitro (Cary et al., 2002; Zhang and Lemaux,

2005; Gordon et al., 2007). De novo shoot organogenesis can be a useful model for the study of

developmental processes in the SAM (Cary et al., 2002; Che et al., 2002; Che et al., 2006;

Gordon et al., 2007).

The dynamics of gene expression of STM, CUC1, CUC2, WUS, and CLV1 during shoot

regeneration have been studied (Cary et al., 2002; Daimon et al., 2003). They showed that CUC1

and CUC2 were strongly expressed at 3 to 6 days on SIM, before the expression of other

markers, and prior to shoot commitment, or any visible green foci or shoot meristem

organization. The increase of the WUS gene expression occurred after 6 days on SIM, and

gradually increased until 15 days on SIM. STM and CLV1 transcript levels rose later than those

of WUS, CUC1, and CUC2, and were significantly higher only after 11 -15 days on SIM, at

about the time of shoot commitment (Cary et al., 2002).

A live imaging approach enabled visualization of the expression of CUC1 and CUC2 via a GFP

fluorescent marker (Cary et al., 2002). This showed that the initially broad expression during

preculture on auxin rich CIM, and the first days on SIM, was progressively localized to the sites

19

of presumptive shoot formation at later stages, finally coinciding with green foci formation and

the development of SAMs (Cary et al., 2002; Gordon et al., 2007). The authors hypothesized that

“restriction of gene expression to the new pattern might be a critical event in shoot commitment”

(Cary et al., 2002). Similar results were obtained by another group, when they analyzed the

expression patterns of the CUC1, CUC2, and STM genes on CIM, and on SIM (Daimon et al.,

2003).

Gordon et al. (2007) studied the dynamics and patterns of expression of several major SAM

genes, such as CUC2, WUS, STM, PIN1, REV, CLV1, and FIL, as well as cytokinin (ARR5) and

auxin (DR5) reporters, during de novo SAM formation in Arabidopsis tissue culture. The authors

demonstrated that CUC2 and PIN1 were expressed within the same domains in the callus tissues

on CIM and in the promeristems on SIM. The distribution of the signals from the hormone

reporters ARR5 and DR5 showed that shoot meristems can be initiated in areas of low auxin and

high cytokinin responses (Gordon et al., 2007). At later stages of meristem development, the

DR5 signal was detected following the PIN1 reporter upregulation at the sites of future leaf

primordia formation. In contrast to DR5, the cytokinin responsive ARR5 reporter was expressed

in the areas of shoot meristem initiation, and within the developing meristems; but, was

downregulated in leaf primordia, and in the areas of callus where no shoots or root meristems

were formed (Gordon et al., 2007). Based on the results, Gordon et al. (2007) suggested that de

novo shoot organogenesis can be broken down into distinct events: callus induction, cytokinin-

induced specification of cell identity within the callus, radial patterning within shoot progenitors,

and meristem morphogenesis. They hypothesized that the non-homogenous distribution of

cytokinin and auxin may be a key factor for the partition of the cell identity within a callus on

SIM. They also hypothesized that the hormone gradient must be gradually reorganized from the

disrupted initial conditions, during shoot induction in culture (Gordon et al., 2007).

In summary, the expression of key meristem genes during de novo SAM formation occurs during

late CIM (4 days) or early SIM (3-6 days) incubation times in Arabidopsis, which appears to be

related to the acquisition of competence on CIM, and to the shoot commitment stage during the

first days on SIM (Che et al., 2002; Che et al., 2006; Christianson and Warnick, 1985; Gordon et

al., 2007).

20

1.5.2 Mutations affecting shoot regeneration in Arabidopsis

Cytokinins mediate de novo shoot regeneration in tissue culture (Skoog and Miller 1957; Gaba,

2005). Accumulating evidence supports the idea of molecular interactions between cytokinin

signaling, the cell cycle, and shoot meristem developmental pathways (Riou-Khamlichi et al.,

1999; Sieberer et al., 2003; Zhang and Lemaux, 2004; Zhang and Lemaux, 2005). Key SAM

genes, and the genes of cytokinin signaling are among the primary candidates for study in tissue

culture to determine their possible effects on the efficiency of shoot regeneration in vitro (Zhang

and Lemaux, 2004; Zhang and Lemaux, 2005). Among the genes involved in meristem initiation

and maintenance, CUC1, CUC2, STM, WUS, ENHANCER OF SHOOT REGENERATION 1 and

2 (ESR1, and ESR2) showed a significant effect on shoot regeneration. Typically, mutations in

these genes affect shoot meristem initiation, but not the callus initiation stages (Barton and

Poethig, 1993; Aida et al., 1997; Banno et al., 2001; Daimon et al., 2003; Gordon et al., 2007).

An elevation of endogenous cytokinin levels generally results in increased shoot regeneration in

cytokinin overproducing mutants, and often in shoot formation on cytokinin-free or hormone-

free media (Chaudhury et al., 1993; Kunkel et al., 1999; Frank et al., 2000; Kakimoto, 2001;

Catterou et al., 2002; Zubko et al., 2002; Sun et al., 2003; Zhang and Lemaux, 2004; Zhang and

Lemaux, 2005). However, genes belonging to other signaling pathways can be implicated in

shoot regeneration as well, and some of them will be discussed in the following sections.

1.5.3 Mutations in auxin signaling genes affecting shoot regeneration

Although SIM contains auxin (Skoog and Muller, 1957), relatively limited information is

available about the direct effects of auxin, and auxin signaling genes on shoot regeneration in

tissue culture. Typically, auxin signaling genes are not among those reviewed in the context of

shoot regeneration in vitro (Zuo et al., 2002b; Howell, 2003; Zhang and Lemaux, 2004; Zhang

and Lemaux, 2005). Nevertheless, auxin response marker levels are highly elevated in shoot

meristems in planta (Kakani et al., 2009). Evidence of the effects of mutations in auxin signaling

genes on shoot regeneration has been demonstrated (Coenen and Lomax, 1998; Chatfield and

Raizada, 2008; Kakani et al., 2009). Also, many auxin signaling genes are implicated in cross

talk with other signaling pathways (Hobbie and Estelle, 1994). In addition, impairment of auxin

genes can affect coordinated interaction among multiple developmental processes (Gordon et al.,

2007; Kakani et al., 2009).

21

For example, the AUXIN RESISTANT1 (AUX1) gene encodes a permease-like membrane protein,

facilitating auxin influx into plant cells. Loss of function aux1 mutants result in resistance to

auxin, ethylene, and cytokinin (Hobbie and Estelle, 1994). The AUX1 gene is highly expressed

within the SAM, in the root tips of plants (Swarup et al., 2004), and in callus culture (Kakani et

al., 2009). Remarkably, aux1 LOF mutants regenerated roots instead of shoots on SIM (Kakani

et al., 2009).

Loss of the PIN1 gene function, encoding the auxin efflux protein and facilitating polar auxin

transport, resulted in a moderate decrease of shoot regeneration (Gordon et al., 2007). However,

the PIN gene family consists of eight members with partly redundant functions. Therefore an

effect on shoot regeneration could be more prominent if multiple PIN mutations were tested in

tissue culture.

The ALTERED AUXIN RESPONSE (AXR1) gene encodes a subunit of the protein RUB1 which

is required for auxin-dependent ubiquitin-mediated degradation of Aux/IAA proteins (Gray et

al., 2001; del Pozo et al., 2002). The AXR1 gene is highly expressed in growing cells throughout

plant organs. Loss of AXR1 function results in decreased auxin responses, auxin resistance in all

plant organs, reduced apical dominance, fewer lateral roots and root hairs, low fertility, impaired

callusogenesis, and abnormal expression of the Aux/IAA and SAUR genes (Lincoln et al., 1990;

del Pozo et al., 2002). Cross-resistance to cytokinin was also reported for axr1 mutants (Hobbie

and Estelle, 1994). Remarkably, the axr1 mutants were dramatically impaired in shoot

regeneration, which was abolished in strong axr1 alleles, and reduced to only 2-3% in weaker

alleles (Chatfield and Raizada, 2008). Similar to aux1 mutants, axr1 mutants also developed

roots rather than shoots on SIM (Chatfield and Raizada, 2008).

Constitutively auxin-overproducing superroot1 (sur1) explants demonstrated auxin-autonomous

growth, proliferation, and root formation on a hormone free medium. Unlike the wild type, they

were capable of shoot regeneration on a medium with cytokinin as the sole hormone (Boerjan et

al., 1995). In contrast, superroot2 (sur2) explants, which also demonstrated elevated auxin

levels, failed to produce shoots on a cytokinin containing medium (Delarue et al., 1998). The

authors discussed the difference between the sur1 and sur2 phenotypes. They proposed that,

while sur1 constantly overproduces auxin, sur2 accumulates auxin as a result of an impaired

mechanism of auxin conjugation, and possibly of impaired auxin transport (Delarue et al., 1998).

22

1.6 Regeneration of adventitious roots in vitro

In tissue culture, regeneration of adventitious roots occurs in response to high auxin and low

cytokinin levels, or when auxin is the sole hormone in the root inducing medium (RIM) (Miller

and Skoog, 1957; Gaba, 2005). Rooting is often a critical step in plant propagation, because

many species are extremely recalcitrant to adventitious root formation (de Klerk, 2002; Schwarz

et al., 2005). Although roots have been routinely induced in response to auxin in plant

propagation (Callis, 2005; Schwarz et al., 2005), the molecular mechanisms of this event are not

completely clear (Schwarz et al., 2005; Rose et al, 2006).

Three temperature-sensitive mutants of Arabidopsis, shoot redifferentiation 1, 2, and 3 (srd1,

srd2 and srd3), defective in shoot regeneration, were isolated as tools for the study of

organogenesis (Yasutani et al., 1994). Genetic analysis indicated that mutations resulted from

single, nuclear, recessive mutations in three different genes located on chromosome 1,

designated SRD1, SRD2, and SRD3. Products of these SRD genes function at different stages of

shoot regeneration (Ozawa et al., 1998). Using srd1, srd2, srd3 mutants, Ozawa et al. (1998)

were able to break down more precisely the time frame for acquiring organogenesis competence

in Arabidopsis calluses. They demonstrated that the competence to form organs is acquired

sequentially: first roots, and then shoots. They also showed that competence to form roots

depends on the type of explant. While hypocotyl explants required CIM preincubation before

transfer to RIM, no CIM stage was required for root explants (Ozawa et al., 1998).

Histological examination of adventitious root formation was performed on leaf explants in

Medicago truncatula (Rose et al., 2006), and in tobacco (Attfield and Evans, 1991). In both

studies, root primordia were formed from procambial cells in veins, which function as

pluripotent stem cells, with the ability to form either root primordia or vascular tissues in

response to added auxin (Attfield and Evans, 1991; Rose et al., 2006). Rose et al. (2006)

concluded that pools of stem cells exist in vascular tissues, which - in combination with auxin

and other factors, such as ethylene and other hormones--drive the diversity of plant

developmental responses.

Cytokinin inhibits root formation in planta (Aloni et al., 2006; Riefler et al., 2006; Kyozuka,

2007), and in vitro (Sugiyama, 1999; Kubo and Kakimoto, 2000). Excessive development of

roots on explants cultured on cytokinin containing media can be attributed either to a low

23

cytokinin to auxin ratio, to an impairment of cytokinin signaling, or to a low cytokinin sensitivity

(Kubo and Kakimoto, 2000; Kakani et al., 2009). When cytokinin responses were elevated,

opposite traits were observed. Such explants typically failed to produce roots, as upon

overexpression of the CYTOKININ-INDEPENDENT1 (CKI1) gene or the

ISOPENTENYLTRANSFERASE (IPT) gene, which mediate cytokinin responses (CKI1) and

cytokinin biosynthesis (IPT) (Kakimoto, 1996; Kunkel et al., 1999; Kakimoto, 2001; Sun et al.,

2003).

Auxin promotes the initiation of root regeneration, but the growth of already formed root

primordia is largely a hormone independent process (Christianson and Warnick, 1985; reviewed

in Schwarz et al., 2005). Auxin overproducing explants of sur1 mutants were capable of root

initiation on a hormone free medium (Boerjan et al., 1995). Also, Frank et al. (2000) isolated

several lines of autotrophic calluses producing roots on a hormone free medium. A molecular

analysis of these lines demonstrated that these rooting calluses had either auxin overproduction,

or had increased levels of Aux/IAA genes, such as IAA1, IAA2, or IAA9 (Frank et al., 2000). The

authors suggested that loss-of-function mutations of auxin response repressors of the AUX/IAA

family can result in a constitutive auxin response, and root overproduction (Frank et al., 2000).

Auxin insensitivity often causes impairment in root regeneration (Sugiyama, 1999, and

references therein). Thus, auxin-tolerant rac mutants in tobacco are defective in primary root

formation, and regeneration of adventitious roots. RAC can be involved in the auxin transduction

pathway, modifying the levels of auxin sensitivity required for adventitious root formation

(Sugiyama, 1999, and references therein).

Che et al. (2006) demonstrated - using the gene profiling method - that molecular signatures for

callus formation and root regeneration are quite similar, because both represent developmental

processes induced by an auxin-rich medium. Competence for root regeneration correlated with

an increased expression of IAA genes on CIM. Many IAA genes were downregulated shortly after

transfer on SIM, at the same time as loss of root commitment occurred (Cary et al., 2002; Che et

al., 2006). The main difference between callusogenesis and root formation was in the expression

of root specific genes associated with the formation of the cell wall, and with vascular

development (Che et al., 2006).

24

The similarity of callus induction and root development pathways was also reflected in auxin

deficient root initiation defective (rid1 and rid2) mutants, which were impaired in both callus

and root formation (Konishi and Sugiyama, 2003). However, one of the mutants, rid3, was

specifically affected only in adventitious root growth, but not callus formation (Konishi and

Sugiyama, 2003). Interestingly, the rid3 mutants were also significantly affected in shoot

initiation, and in the expression of some SAM genes. This demonstrated that the regeneration of

adventitious roots and shoots in culture can also share partly overlapping developmental

pathways, presumably at the stages of acquiring competence, and of promeristemoid formation

(Christian and Warnick, 1985; Schwarz et al., 2005; Tamaki et al., 2009). Recently, it has been

shown that lateral root meristem (LRM)-like structures are formed during CIM incubation; when

calluses are transferred on RIM or SIM, these LRM-like primordia convert to either root or shoot

meristems (Che et al., 2007; Atta et al., 2009; Sugimoto et al., 2010). The enhanced adventitious

root (on RIM) and shoot (on SIM) regeneration in increased organ regeneration 1(ire1) mutants

also supports the model of shared developmental pathways during root and shoot regeneration

(Cary et al., 2001).

1.7 General Research Objectives

In this study, the two related ARFs, MP/ARF5 and NPH4/ARF7, were investigated with regard to

their roles in callusogenesis and de novo organogeneses (shoot and root formation). Mutations in

the MP/ARF5 gene result in the formation of rootless seedlings (Mayer et al., 1991; Berleth and

Jurgens, 1993), and severely reduced auxin sensitivity and auxin transport (Przemeck et al.,

1996; Mattsson et al., 2003; Li et al., 2006; Schuetz et al., 2008). Moreover, MP is upstream of

several meristematic genes, such as CUC and STM, during embryonic meristem formation (Aida

et al., 2002); and, of the ESR1/DRN gene during cotyledon development (Cole et al., 2009).

These genes are also known to be required for shoot regeneration in vitro (Barton and Poethig,

1993; Banno et al., 2001; Daimon et al., 2003; Tamaki et al., 2009). Thus, MP/ARF5 may

possibly be involved in de novo organogenesis. NPH4/ARF7 plays a central role in the

modulation of auxin-dependent differential growth (Stowe-Evans et al., 1998; Watahiki et al.,

1999; Hardtke et al., 2004). nph4 mutants exhibit severely reduced auxin sensitivity, severely

impaired expression of many primary auxin dependent genes, but they have normal responses to

other hormones, including cytokinin (Stowe-Evans et al., 1998; Okushima et al., 2005; Wang et

al., 2005).

25

The general research objectives addressed in this thesis are therefore:

1) To explore the effect of altered auxin signaling on callusogenesis, and shoot and root

regeneration efficiency in vitro;

2) To investigate the impact of explant organ origin on callusogenesis and organogenesis

responses in the genotypes of Arabidopsis thaliana with altered auxin signaling;

Each objective is investigated for both loss- and gain-of-function alleles of specific ARFs:

a) LOF mutations in MP/ARF5 (mpG12) and NPH4/ARF7 (nph4-1);

b) MP GOF transgene dMP, which should no longer be regulated by AUX/IAA proteins (Fig.

1.2; Krogan, 2006)

26

27

Chapter 2

Materials and Methods

2.1 Plant material

All genotypes used in this research were in the Columbia (Col-0) background of Arabidopsis

thaliana. The severe allele mpG12, induced by gamma-ray mutagenesis (Hardtke and Berleth,

1998) was used as a MP LOF mutant. The dMP transgene line, generated by Krogan (2006), was

used in the experiments as a GOF mutation in the MP gene. The dMP line was obtained as a

result of the insertion into the Col-0 background of a truncated copy of the MP gene, which lacks

regulatory and dimerization domains III and IV (Krogan, 2006). For the nph4 LOF allele, I used

the presumed null allele nph4-1, generated by fast neutron mutagenesis (Harper et al., 2000). For

all mutant alleles and the dMP transgene line only homozygous genotypes were used for the

experiments. Some characteristics of the genotypes are represented in Table 2.1; Fig. 2.1.

2.2 General growth conditions

For the experiments and seed propagation, all seeds were dry-sterilized. For this procedure, a

small amount of seeds was placed inside open Eppendorf tubes into an exicator. Sterilization of

the seeds occurred by chlorine gas produced as a result of the reaction when 3 ml of concentrated

hydrochloric acid were added to 100 ml of a 3% bleach solution inside the closed exicator. After

4 hours of sterilization in chlorine gas, the seeds were aseptically transferred into a sterile

laminar box and sown on tissue culture dishes (100x20 mm, Sarstedt, USA) containing 0.5X

Murashige and Skoog (MS) salts (Sigma, USA), 0.5mg/L morpholino ethane sulfonic acid

(MES), 1X Gamborg’s vitamin solution, 1.5% (w/v) sucrose, pH 5.7 (KOH), solidified by 4-5

g/L Bacto-agar (BioShop, Canada) for the purpose of seed propagation, or solidified by 2.4 g/L

Phytogel (Sigma) for the tissue culture experiments. The density of the seeds was about 1-2

seeds per 1 cm2. For seed stratification, plates with seeds were kept in the dark in a refrigerator at

40C for 4 days. After stratification, the seeds were germinated and grown at 24-260C under

constant light (100μEm-2sec-1). “Days after germination” (DAG) are defined as the number of

days the seeds were cultivated in light conditions after stratification.

Table 2.1: Main characteristics of the mutant genotypes, mpG12, dMP, and nph4-1

Genotypes Type of

mutation, and

the strength of

the allele

Type of

mutagenesis

Character of molecular changes in the mutated gene

sequence

Phenotype in

homozygous

seedlings in normal

condition

References

1.

mpG12 recessive, LOF,

presumed null

allele

gamma-ray

mutagenesis

.

the molecular lesion in the allele G12 has not been fully

characterized. It is associated with multiple RFLPs

rootless seedlings,

reduced vasculature,

pin like

inflorescences

Hardtke and

Berleth, 1998;

Hardtke et al.,

2004

2.

dMP dominant, GOF insertion of

transgene

dMP transgene express the truncated MP protein lacking the

C-terminal part from the middle of domain III (absence of

dimerization domains III and IV presumably prevent the

Aux/IAA and ARF proteins binding the following up or

down regulation of dMP protein function)

elongated

cotyledons and

narrow shape of

leaves; increased

vasculature, infertile

flowers

Krogan, 2006

3.

nph4-1 recessive, LOF,

presumed null

allele

mutagenesis by

fast neutrons

A alteration of the coding sequence as a result of the

combination of inversion with internal deletion between

exons 11 and 12

epinastic or

hyponastic rosette

leaves in seedlings

Stowe-Evans et

al., 1998; Harper

et al., 2000

28

mpG12

A B

C DD

E F G

wt nph4

dMP mpG12

wt mpG12 dMP

Figure 2.1 Seedling (A-D), and inflorescence (E-G) phenotypes in the LOF mutants nph4-1 (B),

mpG12 (D, F), and the GOF transgene line dMP (homozygous) (C, G) compared to wild type (A,

E). Genotypes as indicated on top right.

(Modified from Krogan, 2006; Hardtke et al., 2004)

29

For the propagation of the dMP transgene line, dMP seeds were sown on ½ MS medium without

sucrose, supplemented with 10 μM Basta. This allowed selective growth of homozygous and

heterozygous dMP seedlings, while wild type seedlings were eliminated by Basta. The dMP

homozygous seedlings can be distinguished from heterozygous dMP seedlings based on their

elongated cotyledons and narrowed first leaves (Table 2.1; Fig. 2.1). Only heterozygous dMP

plants are fertile and can be used for propagation; and only homozygous dMP seedlings were

used for the experiments. Seedlings intended for propagation were transferred into Promix BX

soil at 12 DAG, and grown under fluorescent light (100 μE/m2/s2) in an 8-hour dark cycle, at

22°C.

2.3 Hormone stocks

Several auxin and cytokinin plant hormones were used for the experiments. The auxins 2,4-

dichlorophenoxyacetic acid (2,4-D, Sigma), indole-3-butyric acid (IBA, Sigma), and 3-

indoleacetic acid (IAA, Sigma) were dissolved in liquid dimethyl sulfoxide (DMSO) at a

concentration of 50 mg/ml. Cytokinin stock solutions, such as kinetin (Sigma) and 6-

(gamma,gamma-Dimethylallylamino)purine (2-iP, Sigma), were used as commercially prepared

(Sigma) hormone solutions, dissolved in water (sterile filtered, concentration 1 mg/ml). All

stock hormone solutions were stored at -200C.

2.4 Plant DNA extraction

A crude, rapid DNA isolation was done to collect templates for PCR analyses and genotyping.

For crude DNA isolation a small rosette leaf or a small piece of tissue culture was flash-frozen in

liquid nitrogen, and ground in a 1.5 ml Eppendorf tube with a micropestle on ice. Then 500 μL of

extraction buffer [250mM Tris-HCl, 250mM NaCl, 25mM EDTA, 0.5% (w/v) SDS] were added,

and the tube was vortexed for 5 sec. To discharge debris the tube was centrifuged for 1 minute in

a table-top centrifuge at 14000 rpm/RT. The supernatant was transferred into a new Eppendorf

tube and 300 μl of Isopropanol was added to 300 μl of supernatant, vortexed, and left to sit for 2

minutes at RT. Then the mixture was centrifuged at 14000 rpm for 5 minutes at RT, and the

supernatant was discarded. The pellets were washed once with 750 μL of ice-cold 70% (v/v)

ethanol, air-dried, and resuspended with a pipette tip in 30 μL of water. The mix was briefly

centrifuged one more time at 14000 rpm/RT for 1 minute, and the supernatant was carefully

30

removed and stored at -200C. 1μL of crude DNA extract was used as a template per 25μL or

50μL of PCR mix.

2.5 General procedure for polymerase chain reaction (PCR)

PCRs were performed in Biometra UNO-Thermoblock (Germany). The PCR mix (25μL total

volume) contained 1 μl of DNA template, 2.5 μL Taq buffer (supplied with polymerase), 2.5 μL

of MgCl2, 2.5 μL of each primer (Sigma-Genosys), 2.5 μL of dNTPs (Fermentas), and Taq DNA

polymerase 1 unit. Double distilled autoclaved water was added to a total volume of 25 μL was

reached, and then all was overlaid with light mineral oil. The general thermocycling regime was

performed as follows: 94°C for 90s, 40 cycles of (94°C for 30s, 55°C for 30s, 72°C for 2min),

72°C 10min; hold at 4°C.

2.6 Genotyping

Several screening methods can be applied to distinguish between plant samples containing dMP

insertion (homogenous or heterogeneous dMP genotypes) and those of wild type. First, only the

seeds containing the dMP transgene (homogenous or heterogeneous dMP seedlings) can survive

on Basta plates, while wild type seedlings will not germinate, or will die soon after germination

(see Section 2.2, General growth conditions). The homogenous and heterogeneous dMP

seedlings can be distinguished based on a difference in their phenotypes, where homozygous

dMP seedlings had elongated cotyledons and narrowed first leaves (Table 2.1; Fig. 2.1).

To differentiate DNA samples containing dMP (homo or heterozygous insertion) from wild type

the PCR primers for the Basta resistance gene (Bar) were designed. The primers were designed

at the beginning and the end of the Bar gene and gave a PCR product of 403bp. The primer

sequences are:

BST-3-F: 5’-GGTCTGCACCATCGTCAACCACTA-3’

BST-3-R: 5’- CCAGTTCCGTGCTTGAAGCC-3’

The thermocycling regime was performed as follows: 940C for 2min, 35 cycles of (940C for 30s,

620C for 30s, 720C for 1 min), 720C for 5 min; hold at 40C.

31

Homogenous mpG12 seedlings do not form primary roots and hypocotyls (Table 2.1; Fig. 2.1).

This phenotypic trait allows clear distinction of the homogenous mpG12 seedlings from the

heterogeneous (MP/mpG12) or wild type. Additionally, PCR diagnostics were performed on the

genomic DNA to verify a genotypic identity of individual samples (plant or tissue culture

origin). To distinguish between mpG12 homozygous samples and those with at least one wild

type MP copy, primers BS1354-F (5’-GAGATGGCCTGGTTCTAAGTGGC-3’; corresponding

to a central portion of MP ORF) and BS1354-R (5’-GCCAGTTCAACATCTCGGTTATCG-3’;

corresponding to a region of MP 3’-UTR) were used in diagnostic PCR reactions. The presence

of at least one wild type MP gene copy will support amplification of a 2660bp product. In

homozygous mpG12 samples there is no amplification of any PCR products. The thermocycling

regime was performed as follows: 94°C for 90s, 45 cycles of (94°C for 30s, 60°C for 30s, 72°C

for 3min), 72°C 10min; hold at 4°C.

Genotyping of nph4 homozygous individuals was based on PCR diagnostics of plant and tissue

culture samples. To distinguish between nph4-1 homozygous individuals and those with at least

one wild type NPH4 copy, primers NPH4.5 (5’- TCCTGCTGAGTTTGTGGTTCCTT -3’) and

NPH4.6 (5’- GGGGCTTGCTGATTCTGTTTGTTA -3’) were used in diagnostic PCR reactions.

The presence of at least one wild type NPH4 gene copy supports amplification of a 823bp

product, whereas the genomic DNA of nph4-1 homozygous individuals does not support

amplification of any PCR products. The thermocycling regime was performed as follows: 94°C

for 90s, 45 cycles of (94°C for 30s, 63°C for 30s, 72°C for 2min), 72°C 10min; hold at 4°C.

2.7 Microtechniques and microscopy

To introduce explants in tissue culture, seedlings were dissected under a Hund Wetzlar

stereomicroscope (GmbH, Germany) in aseptic conditions in a flowhood. Samples were viewed

under dark field illumination with a Leica MZFLIII stereomicroscope (Leica Microsystem,

Wetzlar, Germany), equipped with a Canon EOS D60 digital camera (Canon inc., Tokyo, Japan).

All images were assembled using Adobe Photoshop 7.0 (Adobe Systems, Mountain View, CA,

USA). The fresh weight of the samples was measured using Sartorius analytical balances

(Sartorius, GmbH, Germany).

32

2.8 Plant tissue culture experiments

Cotyledons, leaves, petioles, hypocotyls, and adventitious roots were used as explants in tissue

culture experiments (Table 2.2). Because mpG12 seedlings lack hypocotyls and primary roots,

no mpG12 hypocotyl explants could be tested. To obtain root explants in mpG12 seedlings, a

special rooting procedure was applied to mpG12 and the rest of the genotypes to obtain

adventitious roots (see Section 2.8.1 for more details). The main steps of the tissue culture

experiments are summarized in Fig. 2.2.

At least three independent replicas of the experiment were taken unless otherwise stated.

Student’s t-test was used to evaluate the significance of the comparison between the mean

responses, such as fresh weight of calluses, number of shoots or roots per callus, etc. For the

purposes of this research, each of the evaluated genotypes was compared against wild type via

Student’s t-test (Microsoft Excel, 2003).

2.8.1 Rooting of dissected mpG12, dMP, nph4-1 and wild type seedlings

Seeds of mpG12, nph4-1, dMP, and wild type were germinated as described above (see for

details Section 2.2, General growth conditions). 5-6 DAG seedlings were dissected under a Hund

Wetzlar stereomicroscope (GmbH, Germany) in aseptic conditions. In rootless mpG12 seedlings,

peg parts (short hypocotyl-like outgrowths below the cotyledon node) were cut with a sharp

scalpel. Seedlings of other genotypes were similarly dissected by cutting off the roots with a part

of the hypocotyl. Dissected seedlings were placed in tissue culture dishes (100x20 mm, Sarstedt,

USA) with root inducing medium (RIM), containing 1 mg/L IBA (Table 2.3), and cultured at 24-

260C under constant light (100μEm-2sec-1). After 7-9 days on RIM, about 5-10% of the mpG12

dissected seedlings, and most of the seedlings of the other genotypes formed adventitious roots.

Rooted seedlings were transferred on hormone-free ½ MS medium, to restore normal root

morphology; and, after 14 days of culture, they were dissected and their adventitious roots were

used for tissue culture experiments.

33

Explants

CIM 6 days

SIM 21 days

Callusogenesis:% callus patterns,

fresh weight

Shoot regeneration:% shoot regeneration,

# shoots per callus

CIM 12 days RIM 21 days

Root regeneration:% root regeneration,

# roots per callus

Figure 2.2 General scheme of the tissue culture experiments and scored traits, unless otherwise

specified in the description of the particular experiment.

34

2.8.2 Preparation of explants for tissue culture

For the tissue culture procedure, seedlings of a particular age (Table 2.2) were aseptically

dissected into explants under a Hund Wetzlar stereomicroscope (GmbH, Germany) in aseptic

conditions. The types of explants, the genotypes, and the age of the seedlings at the time of

explant dissection (DAG) are summarized in Table 2.2.

Each culture dish (100x20 mm, Sarstedt, USA) was divided into sections corresponding to the

genotypes, and explants of each genotype were cultured together. About 18-20 explants per

culture dish were used during callus initiation on CIM, 12 explants per culture dish were used for

shoot initiation on SIM, and about 16 explants per dish were used for root initiation on RIM. The

culture dishes were sealed with Parafilm (Parafilm® M, USA). Tissue culture was maintained

under fluorescent light (100 μE/m2/s2), at 24-250C.

Table 2.2. Types of explants, genotypes, approximate sizes, and DAG at the moment of

dissection and introduction in tissue culture.

Explants Cotyledons Leaves Petioles Hypocotyls Adventitious

roots

Age (DAG) 8 11 11 9 26-28

Size (mm) 4-5 4-5 3-5 4-6 5-7

Genotypes mpG12,

nph4-1,

dMP,

wt (Col-0)

mpG12,

nph4-1,

dMP,

wt (Col-0)

mpG12,

nph4-1,

dMP,

wt (Col-0)

nph4-1,

dMP,

wt (Col-0)

mpG12,

nph4-1,

dMP,

wt Col-0

35

For shoot and root regeneration experiments, explants were precultured for 6 days on CIM and

then transferred on SIM or RIM. For callusogenesis experiments explants were left on CIM for 6

more days (12 days in total). Results of shoot or root regeneration were scored after 21 days on

SIM or RIM. For these purposes explants were screened directly in tissue culture dishes under a

Hund Wetzlar stereomicroscope (GmbH, Germany) or under a dark field illumination with a

Leica MZFLIII stereomicroscope (Leica Microsystem, Wetzlar, Germany), equipped with a

Canon EOS D60 digital camera (Canon inc., Tokyo, Japan). The schema of the experiments is

shown in Fig. 2.2.

2.8.3 Preparation of tissue culture media

The tissue culture procedure followed the protocol suggested by Valvekens et al., (1988).

Following the protocol, Gamborg B5 (B5) salt base (Sigma, USA) was used for the preparation

of CIM and SIM. For both media, CIM and SIM, B5 salt base was supplemented with 2%

sucrose, 0.5 g/L MES, 1x Gamborg’s vitamin solution (Sigma, USA), and pH 5.7 (KOH) before

autoclaving. The hormone compositions varied depending on the type of the tissue culture

medium (Table 2.3). The media were solidified with 2.4 g/L Phytogel (Sigma) and sterilized for

20 min at 1210C and 15 psi in the autoclave (Hirayama, Japan). The liquid hot medium was

poured into tissue culture dishes (about 20 ml/dish) and stored at -40C till the time of the

experiment.

Table 2.3. The hormone composition of tissue culture media: CIM, SIM, and RIM.

CIM SIM RIM

2,4-D - 0.5 mg/L

Kinetin - 0.05 mg/L

IAA - 0.15 mg/L

2-IP - 5 mg/L

IBA - 1 mg/L

36

Chapter 3

Callusogenesis under conditions of altered auxin signal

transduction

3.1 Background and rationale

Callus proliferation is induced in vitro at a certain relative concentration of auxin and cytokinin

in the culture medium (Skoog and Miller 1957). Cell proliferation during callusogenesis leads to

disruption of the ordered morphology of differentiated tissues (Cary et al., 2002). The

mechanisms leading to callus proliferation are not well understood (Cary et al., 2002; Zhang and

Lemaux, 2005). Generally, cell-cycle genes, such as cyclins and cyclin dependent kinases

(CDKs), play a major role in the mediation of hormonal effects, or in the response to wounding

signals, leading to cell proliferation (Hemerly et al., 1993; Soni et al., 1995; Shaul et al., 1996;

Riou-Khamlichi et al., 1999; Sugiyama 1999; Richard et al., 2002; Che et al., 2007). Cis-acting

regulatory elements, specific for the binding of hormone transcription factors, such as AuxREs

for auxin signaling, have been found in the promoters of some cell cycle genes (Richard et al.,

2002; Perrot-Rechenmann, 2010).

Auxin responses, visualized by DR5 and PIN1 driven reporters (reviewed in Chapter 1.2), were

strongly increased during the initiation of callus proliferation. Then they were downregulated

after 5 days on CIM (callus inducing medium). After 10 days on CIM, DR5 and PIN1 reporters

were not detectable in the callus culture (Gordon et al., 2007). Among the other reporters

associated with proliferating callus growth, are the meristem genes CUC1 and CUC2, and the

ARR5 gene, which is involved in cytokinin signaling (reviewed in Chapter 1.4; Cary et al., 2002;

Gordon et al., 2007). The expression of the corresponding reporter genes was observed after 8

days on CIM, and continuously increased during CIM cultivation (Gordon et al., 2007). When

callusogenesis was induced on the medium supplemented with auxin as the sole hormone, CUC2

and ARR5 reporters were still expressed. In contrast, only weak callus proliferation occurred, and

no expression of the callus proliferation reporters, CUC1, CUC2 and ARR5, was observed, when

the explants were cultured on the medium supplemented with cytokinin only (Gordon et al.,

37

2007). This observation emphasizes the importance for callus proliferation of the presence of

auxin, rather than cytokinin, in the culture medium.

As described in Chapter 1 (1.2.5), the auxin response factors MP (ARF5) and NPH4 (ARF7)

belong to the group of the transcriptional activators whose potency is enhanced in the presence

of auxin (Ulmasov et al., 1999a; Ulmasov et al., 1999b; Berleth et al., 2004). Among other

described defects (Hardtke et al., 2004), the loss of MP or NPH4 function severely reduces auxin

sensitivity. The expression of the auxin-responsive reporter DR5 and auxin-inducible genes was

significantly reduced in nph4 and mp mutants (Stowe-Evans et al., 1998; Watahiki et al., 1999;

Hardtke et al., 2004; Li et al. 2006; Przemeck et al., 1996; Mattsson et al., 2003; Okushima et

al., 2005; Wang et al., 2005; Schuetz et al., 2008).

The NPH4 gene is expressed throughout the Arabidopsis life cycle, and NPH4 transcripts are

generally present in all major organs (Hardtke et al., 2004). The MP gene is expressed in broad

domains in embryos and in emerging organs. Later, upon organ maturation, the expression of

MP is gradually narrowed to central domains and then to vascular tissues (Hardtke and Berleth,

1998; Hardtke et al., 2004). The mRNAs of both genes, MP and NPH4, were found to be

ubiquitous and uniformly expressed in Arabidopsis cells in suspension culture (Ulmasov et al.,

1999b; reviewed in Chapter 1.2.4). The genotype dMP generated by the deletion of protein

interaction domains in the MP gene is described in the General Introduction (Chapter 1.2.5; Fig.

1.2; Krogan, 2006). As these interaction domains are required to bind negative regulators of ARF

function, it seems likely that dMP represents a constitutively active MP gene.

In summary, the MP and NPH4 genes play important roles during many auxin-triggered

processes in plant development. Because callus proliferation is triggered by the hormone auxin,

changes in MP and NPH4 function in the loss-of-function mutants may have an impact on

callusogenesis.

3.2 Specific research objectives

The investigation of the possible influence of altered auxin signaling through LOF and GOF

alleles of ARF5/MP and ARF7/NPH4 on callusogenesis was the purpose of this part of the

project. To this end, the loss-of-function mutants mpG12 (Hardtke and Berleth, 1998), and nph4-

1 (Harper et al., 2000), and the gain-of-function transgene line dMP (Krogan, 2006) with 38

truncated dimerization domains III and IV (reviewed in Chapter 1.2.5; Fig. 1.2), were tested to

determine the influence of these genotypes on callus formation in vitro. For a comprehensive

comparison of callus proliferation among the genotypes, different types of explants (cotyledons,

leaves, petioles, hypocotyls, and adventitious roots) were cultured on CIM for 12 days (for

details see Material and Methods, Chapter 2.8). Afterward, the patterns of callusogenesis and the

fresh weights of the calluses were scored. The frequencies of the “whole” type of calluses, and

the fresh weights of the calluses (mean weights of calluses ± SE) were determined based on the

pooled data of all replicas of the experiments. All genotypes used in the experiments were in the

Col-0 background and were compared to the Col-0 wild type. Explants of approximately similar

size and age (DAG) were used for the callusogenesis experiments. This minimized the variations

in the fresh weights of the calluses on CIM (for details see Chapter 2.8.2; Table 2.2).

3.3 Results

3.3.1 Types of calluses on CIM

Callus proliferation on CIM depended on the genotype, and on the type of organ used as explant

(Fig. 3.1). When aerial explants were compared, two major types of calluses were detected after

12 days on CIM. The first type was characterized by even callus proliferation across the whole

explant surface (Fig. 3.2; 3.3). As a result, this pattern of callusogenesis was called the “whole”

type. In the second type, callus proliferation was limited to some patches on the explant surface,

with some areas free of callus proliferation (Fig. 3.2; 3.3). Often, in the latter type of calluses,

cell proliferation occurred mainly near the wounded areas (cut parts) of the explants, with or

without a combination of local patches along the explant surface. Because of the localized

character of the callus proliferation, this second type of callus was called the “patchy” type. No

noticeable difference in the callusogenesis patterns was observed in the adventitious root

explants after 12 days on CIM. The adventitious root explants were relatively evenly covered by

callus cells along their axes (similar to the “whole” type described for callusogenesis in aerial

organs) (Fig. 3.4).

39

0 0.01 0.02 0.03 0.04 0.05 0.06 0.07

roots

hypocotyls

petioles

leaves

cotyledons

Fresh weight, g

wtdMPnph4-1mpG12

A

n=137n=133

n=133n=135

n=161n=161

n=174n=167

n=108n=73

n=109n=62

n=137n=92

n=132

n=116n=114n=111

n=110

0 10 20 30 40 50 60 70 80 90 100

hypocotyls

petioles

leaves

cotyledons

Frequency of whole type calluses, %

wtdMPnph4-1mpG12

n=137n=133

n=133n=135

n=161n=161

n=174n=167

n=108

n=73

n=73n=109

n=62

n=137n=92

n=132

B

“whole” type calluses, %

Figure 3.1. Fresh weight of calluses (mean numbers ± SE) (A), and percentage of the “whole”

type of calluses (B) after 12 days on CIM; n, total number of explants; bars represent SE.

40

A B

C D

6 d CIM, whole type 12 d CIM, whole type

6 d CIM, patchy type 12 d CIM, patchy type

cotyledon cotyledon

cotyledon cotyledon

6 d CIM, whole type

6 d CIM, patchy type

12 d CIM, whole type

12 d CIM, patchy type

E F

G H

leaf leaf

leaf leaf

Figure 3.2 Callus initiation and callus proliferation in cotyledon (A-D) and leaf (E-H) explants

after 6 (A, C, E, G)) and 12 (B, D, F, H) days on CIM. Origin of explant as shown on top right;

days on CIM and types of callus growth (“whole” versus “patchy”) as indicated, bottom right;

arrows indicate sites of callus proliferation

41

3.3.2 Callusogenesis of cotyledon explants after 12 days on CIM

Callus proliferation, indicated by the fresh weights of the cotyledon derived calluses, differed

among the genotypes. The fresh weights of the dMP calluses were similar to those of the wild

type calluses (the wild type calluses 52±1.4 mg, n=137; dMP 52±1.5 mg, n=133) (Fig. 3.1 A). In

contrast, the fresh weights of the nph4-1 and the mpG12 calluses were significantly less than

those of the wild type calluses (nph4-1 13±6 mg, n=135, P<0.001; and mpG12 16±6 g, n=133,

P<0.001) (Fig. 3.1 A). Thus, there were two distinct phenotype classes: the dMP calluses

growing like the wild type ones versus the strongly impaired callus growth in nph4-1 and mpG12

(Fig. 3.1 A).

The distribution of the callusogenesis types (the “whole” versus the “patchy” types) in the

cotyledon explants also dramatically differed among the genotypes (Fig. 3. 1 B; 3. 5 A). All wild

type (n=137) and all dMP cotyledon explants (n=133) had the “whole” type of callusogenesis

(Fig. 3.1 B). In contrast, mpG12 and nph4-1 showed mostly the “patchy” type of callusogenesis.

The “whole” type calluses were found in only 4% of the mpG12 explants (n=135), and in 8% of

the nph4-1 explants (n=133) (Fig. 3.1 B). The impaired callus proliferation in the arf mutants,

indicated by the lower fresh weights of the calluses, was combined with a strong prevalence of

the “patchy” type calluses in these genotypes. In contrast, the dMP and the wild type calluses,

which defined normal callus growth, were exclusively the “whole” type.

3.3.3 Callusogenesis of leaf explants after 12 days on CIM

The fresh weights of the leaf calluses also differed among the genotypes. As with the cotyledon

explants, the fresh weights of the calluses derived from the leaf explants were similar between

wild type and dMP (wild type 60±2.4 mg, n=161; and dMP 65±1.6 mg, n=161). However, the

fresh weights of the nph4-1 calluses (18±1 mg, n=174; P<0.001), and of the mpG12 calluses

(40±0.6 mg, n=167; P<0.001) were significantly less than those of wild type (Fig. 3.1 A). Also,

the fresh weights of the mutant genotypes, nph4-1 and mpG12, differed significantly: the mean

fresh weight of the nph4-1 calluses was only about half that of mpG12 (nph4-1 18±1 mg, n=174;

mpG12 40±0.6 mg, n=167; P<0.001, for the comparison of the mean responses between nph4-1

and mpG12) (Fig. 3.1 A).

42

The distribution of the callusogenesis types (the “whole” versus the “patchy” types) in the

calluses derived from leaf explants also differed among the genotypes (Fig. 3.1 B; 3.5 B). In wild

type, the “whole” type calluses formed in only 54% of the explants (n=161). The frequency of

the “whole” type calluses among the dMP explants (94%, n=161) was much higher compared to

that of the wild type ones (Fig. 3.1 B). The frequencies of the “whole” type calluses were

significantly lower in the mutant genotypes. Only 33% of the mpG12 (n=167), and only 7% of

the nph4-1 (n=174) leaf explants developed the “whole” type calluses (Fig. 3.1 B). The rest of

the mpG12 and nph4-1 leaf explants were only partly covered by callus patches (Fig. 3.5 B).

Thus, analysis of the fresh weights and of the distribution of callusogenesis types in the leaf

explants once again revealed differences in callus proliferation among the genotypes.

Compromised callus proliferation was detected in the nph4-1, and, to a lesser degree, in the

mpG12 leaf explants. Increased callus proliferation was found in the dMP explants, as indicated

by the frequency of the “whole” type calluses.

3.3.4 Callusogenesis of petiole explants after 12 days on CIM

The fresh weights of the petiole derived calluses showed genotype and organ-specific

differences. The fresh weights of the dMP calluses were almost twice those of the wild type

calluses (wild type 22±1 mg, n=108; dMP 38±3 mg, n=73; P<0.001) (Fig. 3.1 A). The fresh

weights of the nph4-1 and mpG12 calluses were significantly lower than those of the wild type

ones (nph4-1 13±1 mg, n=105, P<0.001; mpG12 18±1 mg, n= 62; P<0.05) (Fig. 3.1 A). Further,

between the mutants, the fresh weights of the nph4-1 calluses were significantly less than those

of the mpG12 ones (nph4-1 13±1 mg, n=105; mpG12 18±1 mg, n= 62; P<0.001 for the

comparison of the mean responses between nph4-1 and mpG12).

There were also difference among the genotypes in the distribution of the callus types in the

petiole explants among the genotypes (Fig. 3.1 B; 3.6 A). Almost all wild type (95%) and dMP

(97%) petiole explants developed the “whole” type of calluses (Fig. 3.1 B). By contrast, only

27% of the nph4-1 petiole calluses were of the “whole” type. However, the frequency of the

“whole” type calluses in the mpG12 petiole explants was relatively high (79%), which was closer

to the frequency in wild type than to that in nph4-1.

43

A

6 d CIM, the “whole” type

petiole

C

6 d CIM, the “patchy” type

petiole

B


petiole

D


petiole

E


hypocotyl

G


hypocotyl

F


hypocotyl

H


hypocotyl

Figure 3.3 Callus initiation and callus proliferation in petiole (A-D) and hypocotyl (E-H)

explants after 6 and 12 days on CIM. Origin of explant as indicated on top right; days on CIM

44

and types of callus growth (“whole” versus “patchy”) as indicated, bottom right; arrows indicate

sites of callus proliferation.

To summarize, the distribution of the fresh weights and of the types of calluses in the petiole

explants indicated the restricted character of callus proliferation in the nph4-1 explants; and, to a

lesser degree, in the mpG12 petiole explants. By contrast, callus proliferation was elevated in the

dMP explants, as indicated by the fresh weights of the dMP calluses, the vast majority of which

were of the “whole” type.

3.3.5 Callusogenesis of hypocotyl explants after 12 days on CIM

The fresh weights of the hypocotyl calluses also showed genotype related differences. However,

these differences were weaker compared to those of the other organs. The fresh weights of the

dMP calluses were significantly higher than those of the wild type ones (wild type 19±1 mg,

n=137; dMP 23±1 mg, n=92; P<0.001). The hypocotyl explants were the only type of aerial

explants where the fresh weights of the nph4-1 calluses were not less than those in wild type

(nph4-1 19±1 mg, n=132) (Fig. 3.1 B). No mpG12 hypocotyl explants could be tested, because

of the mpG12 seedlings’ lack of a hypocotyl.

The distribution of the callus types in the hypocotyl explants differed among the genotypes (Fig.

3.1 B; 3.6 B). Most of the wild type explants (82%, n=137) and most of the dMP explants (97%,

n=92) had the “whole” type of calluses (Fig. 3.1 B). In contrast, there were only 35% of the

“whole’ type calluses in nph4-1 (n=132) (Fig. 3.1 B).

To summarize, the comparison of the callus growth and type data suggests that the “whole” type

calluses are not just the result of vigorous growth. Specifically, the fresh weights of the nph4-1

calluses were similar to those of the wild type ones in hypocotyl explants. The “patched” calluses

dominated in this group. Among the genotypes, callus proliferation was once again greatest in

dMP calluses.

3.3.6 Callusogenesis of adventitious root explants after 12 days on CIM

Because mpG12 seedlings lack primary roots (Berleth and Jurgens, 1993), the procedure to

initiate adventitious roots in mpG12 and in the other genotypes preceded the callusogenesis

45

A 6 d CIM, wt

B 6 d CIM, dMP

E 12 d CIM, wt

12 d CIM, dMPD

wt, 6 d CIM wt, 12 d CIM

dMP, 6 d CIM

A B

C

mpG12, 12 d CIMH

nph4-1, 12 d CIMFC 6 d CIM, nph4-1

D 6 d CIM, mpG12

nph4-1, 6 d CIM

mpG12, 6 d CIM

E

G

Figure 3.4 Callus initiation and callus proliferation in adventitious root explants after 6 (A, C, E,

G) and 12 (B, D, F, H) days on CIM. The genotypes and days on CIM as indicated on top right;

arrows indicate sites of callus proliferation.

46

wt

mpG12 nph4-1

dMP

A

cotyledons

dMP

mpG12

wt

nph4-1

B

leaves

Figure 3.5 Cotyledon (A) and leaf (B) explants after 12 days on CIM. Genotypes indicated

within plate sectors; arrows point at “patchy” type calluses

47

dMP

mpG12

wt

nph4-1

A

petioles

dMP

wt

nph4-1

B

hypocotyls

Figure 3.6 Petiole (A) and hypocotyl (B) explants after 12 days on CIM. Genotypes indicated

within plate sectors; arrows point at the “patchy” type calluses.

48

dMP

nph4-1mpG12

wt

roots

Figure 3.7 Adventitious root explants after 12 days on CIM. Genotypes indicated within plate

sectors.

49

experiments (see Chapter 2.8.1 for details). Seedlings of all the genotypes were dissected by

cutting off the lower part of their hypocotyl (or the low part of the peg in the mpG12 seedlings).

The dissected seedlings were re-rooted on the auxin (IBA) containing medium, and then

transferred to the hormone-free 1/2MS medium to allow the outgrowth of adventitious roots.

These adventitious roots were used as explants in the experiments described below (for details

see Chapter 2.8).

In this category, root explants, the distribution of the fresh weights of the calluses among the

genotypes was different compared to those of the aerial organs (Fig. 3.1A). The fresh weights of

the calluses were: wild type 56±0.4 mg, n=116; the similar nph4-1, 57±0.4 mg, n=111; and, the

significantly lower (by more than half) mpG12, 26±0.2 mg, n=110 (P<0.001). In contrast to the

growth of calluses on all the aerial organs, the growth of the dMP calluses on the root explants

was slightly, but significantly, lower than that of the wild type calluses (dMP 4±0.4 mg, n=114;

P<0.01) (Fig. 3.1 A).

Thus, in this category of root explants, callus growth among the genotypes was quite different

than that in the aerial explants. No genotype related difference in callus types was detected in

this category, and the calluses of all the genotypes looked relatively similar after 12 days on

CIM, resembling the “whole” type calluses of the aerial explant group.

3.3.7 Callus proliferation in aerial explants after 6 days on CIM

The difference in phenotypic classes between genotypes observed after 12 days on CIM in

calluses derived from aerial explants (the “whole’ or the “patchy’’ types of calluses) raises the

question of the origin of this difference. Two distinct possibilities can be considered. First, the

difference in callus types might simply reflect the difference in callus growth rate. In this case,

two callus types could be initiated as local callus patches along the explant surface, and around

the wounded areas. Later, these areas would merge to form the “whole” calluses, when callus

proliferation was strong, or stay “patchy”, when the callus growth rate was low. Alternatively,

the two types could reflect genuinely different growth patterns observable right from the

beginning of callus growth. To distinguish between these two possibilities, the explants were

visually examined after 6 days on CIM.

50

When aerial explants (cotyledons, leaves, petioles and hypocotyls) were examined after 6 days

on CIM, clear differences were detected between explants later developing the “whole” type or

the “patchy” type calluses. Most of the explants (more than 90%), which later developed the

“whole" type of calluses, were getting visibly friable and partly translucent. Their original dense

green color was changing to whitish or yellowish green (Fig. 3.2; 3.3). Callus proliferation in

these explants occurred evenly across the entire explant surface. Further, when cotyledon and

leaf explants were examined, the callus proliferation was denser along the veins and the petiole

parts (Fig. 3.2). All these changes made the explants developing into the “whole” type calluses

look quite distinct from those developing into the “patchy’ type calluses (Fig. 3.2; 3.3).

By contrast, in the majority of the explants (more than 90%) which later developed the “patchy”

type of calluses, relatively large areas of the explants retained their original appearance after 6

days on CIM. In these areas, the tissues did not look friable and translucent. The color of the

explants did not change from the original dense green to whitish or yellowish shades. Initiation

of callus proliferation in these explants was limited to the wounded (cut) areas (in all organ types

of the aerial explants), and to some locally restricted patches of proliferating cells on the explant

surface, when the cotyledon and leaf explants were examined (Fig. 3.2; 3.3). The rest of the

explant tissues stayed “untouched” and mainly preserved their original color and structure (Fig.

3.2; 3.3).

In summary the results indicated that the differences in the callus types (the “whole” type versus

the “patchy” type) were due to the distinctive initiation of callus proliferation in the explants,

rather than to the differences in the callus growth rate, even though there were differences in the

callus growth rates.

3.4 Discussion and Conclusions

The results in this chapter indicate genotype and organ related differences in callus proliferation

responses among the genotypes. These differences affect both the callus growth rate, as well as

the pattern of proliferation.

In the calluses derived from the aerial explants (cotyledons, leaves, petioles, and hypocotyls),

two major phenotypic classes of callus type were observed: the “whole” type, with even callus

proliferation along the entire explant surface; and the “patchy” type, where callus proliferation 51

occurred as isolated patches, often localized to the wounded areas. The callus types and the

callus weights differed among the tested genotypes, falling into two major groups. The first

group was comprised of the dMP and wild type genotypes. Callusogenesis in dMP was either

equal to, or sometimes greater than that in wild type, as indicated by the fresh weights and the

frequencies of the “whole” type calluses. The second group was represented by the arf mutants,

nph4-1 and mpG12. In these, callusogenesis was partly impaired and significantly weaker

compared to that in wild type. This was indicated by the lower fresh weights of the nph4-1 and

the mpG12 calluses, and usually by much lower frequencies of the “whole” type calluses. Also,

of these arf genotypes, callusogenesis compromise was greater in nph4-1 than in mpG12, when

their aerial explants were cultured on CIM. The differences in the callus types (the “whole” type

versus the “patchy” type) were as a result of the distinct initiation of callus proliferation

(throughout the explant surface in the “whole’ type, or in local patches in the “patchy” type of

calluses), rather than as a result of differences in the callus growth rate.

The callusogenesis responses were different when calluses derived from the adventitious root

explants were compared. Although no distinct callus types were detected, genotype related

differences in callusogenesis were indicated by the fresh weights of the calluses. Dissimilar to

calluses derived from the aerial explants, no impairment in callusogenesis was detected in nph4-

1root explants. These data are compatible with previous observations that only the aerial organs,

but not the roots, of nph4 plants were resistant to toxic concentrations of the synthetic auxin 2,4-

D (Watahiki and Yamamoto, 1997). In contrast to nph4-1, callusogenesis in mpG12, as indicated

by the fresh weights of the calluses, was significantly weaker compared to that of wild type.

Also, the callus weights were slightly, but significantly, lower in the dMP calluses. Generally,

however, in this category of root explants, differences in callusogenesis among the genotypes

were more subtle than those in the calluses derived from the aerial explants. Given the artificial

character of the procedure for the induction of adventitious roots (the long exposure to an auxin

containing medium followed by culture on a hormone free medium), the observed differences in

callus weights might reflect the genotype specific response to preculture conditions during

adventitious root induction, rather than a true genotype related difference in callus proliferation.

In nature, callusogenesis is a part of the mechanism of protection facilitating the repair of

wounded or damaged tissues (Frank et al., 2000). This mechanism involves auxin signaling,

which is crucial for callus initiation (Frank et al., 2000; Gordon et al., 2007).Thus, it is plausible 52

that auxin insensitive arf mutants are impaired in callusogenesis. The relative degree of

impairment may reflect the degree to which a given ARF gene is involved in auxin signaling in a

specific organ or tissue. It may also reflect the degree to which a particular ARF gene is related

to the regulation of the cell-cycle genes mediating cell proliferation responses (Hemerly et al.,

1993; Richard et al., 2002; Che et al., 2007; Perrot-Rechenmann, 2010).

The dMP transgene represents a gain-of-function allele of MP, possibly a constitutively active

allele (Krogan, 2006). Callusogenesis was strong in the dMP transgene line in aerial, but not in

root explants. In this case, one may speculate that this reflects the enhanced auxin signal

transduction features of dMP tissue (Krogan, 2006).

To summarize, loss of MP and particularly NPH4 functions in mpG12 and nph4-1 mutants

resulted in the impairment of callus proliferation in particular categories of explants. The

presumably elevated MP function in the dMP genotype may increase callus proliferation in aerial

explants, and decrease the same in adventitious root explants which needs further evaluation.

53

Chapter 4

Shoot regeneration under conditions of altered auxin signal

transduction


Shoot regeneration in vitro is triggered by a high cytokinin to auxin ratio in the tissue culture

medium (Skoog and Muller, 1957; Gaba, 2005). Although the phytohormone auxin is always

used in the composition of a shoot inducing medium (SIM), relatively little is known about the

effect of auxin on shoot regeneration in tissue culture. Typically, the role of auxin in this process

is only discussed in the context of the ratio of auxin to cytokinin (Howell 2003; Zhang and

Lemaux, 2004; Gaba, 2005; Zhang and Lemaux, 2005). However, several factors can contribute

to the effect of auxin signaling on shoot regeneration. First, auxin plays a key role in shoot

meristem patterning (Reinhardt et al., 2000); auxin levels are high in shoot meristems in planta

(Kakani et al., 2009). Therefore, proper auxin signaling might be required for de novo shoot

meristem assembly (Coenen and Lomax, 1998; Gordon et al., 2007; Chatfield and Raizada,

2008; Kakani et al., 2009). Second, many genes linked to auxin signaling may also be implicated

in other hormone pathways, due to existing cross talk between signaling pathways (Hobbie and

Estelle, 1994; Swarup et al., 2002). As a result, auxin gene impairment may affect coordinated

interactions among multiple developmental processes (Gordon et al., 2007; Kakani et al., 2009).

Third, shoot regeneration is severely impaired in a number of auxin signaling mutants, such as

aux1 (Kakani et al., 2009) and axr1 (Chatfield and Raizada, 2008), and mildly impaired in pin1

(Gordon et al., 2007). Thus, genes involved in auxin signaling, auxin homeostasis, and auxin

transport may affect processes of de novo shoot regeneration in vitro (reviewed in Chapter 1.7.5).

The auxin response factors MP (ARF5) and NPH4 (ARF7) interact with each other and have

partly overlapping functions. They are both transcriptional activators (Ulmasov et al., 1999a;

Ulmasov et al., 1999b, Berleth et al., 2004; Hardtke et al., 2004). Mutations in the MP gene

affect the formation of the embryonic root meristem and result in rootless seedlings (Berleth and

Jurgens, 1993), and in severely reduced auxin sensitivity, auxin signaling and auxin transport (Li

et al. 2006, Przemeck et al., 1996, Mattsson et al., 2003, Schuetz et al., 2008). Moreover, in

54

planta, MP is involved in the regulation of several meristematic genes, such as the CUC genes

and STM, during embryonic meristem formation (Aida et al., 2002). MP is also linked to the

ESR1/DRN gene during cotyledon development (Cole et al., 2009). Because all these genes are

known to be crucial for de novo shoot meristem formation (Zhang and Lemaux, 2005), the MP

gene may have a significant effect on shoot regeneration.

Regarding nph4 mutants, they exhibit auxin resistance in the aerial parts of plants, indicated by

their resistance to toxic concentrations of auxin, and by the low signal of the DR5 auxin response

reporter. Further, the levels of many primary auxin response genes are severely affected in nph4

mutants (Stowe-Evans et al., 1998, Wang et al., 2005). Notably, nph4 mutants represent a case of

auxin-resistant mutants that are not cross-resistant to other plant hormones, including cytokinin

(Watahiki and Yamamoto, 1997, Wang et al., 2005).

The genotype dMP, generated by the deletion of protein interaction domains in the MP gene has

been described (Krogan, 2006; see also Chapter 1.2.5; Fig. 1.2). As these interaction domains are

required for the binding of regulatory proteins, it seems likely that dMP represents a

constitutively active MP gene.

In this chapter, the roles of the MP and NPH4 genes in de novo shoot regeneration are explored.

The extent to which shoot regeneration in each of the investigated genotypes is further

influenced by the origin of the tissue sample is also explored.


Loss-of-function mutants mpG12 (Hardtke and Berleth, 1998), and nph4-1 (Harper et al., 2000),

and the gain-of -function transgene line dMP with truncated dimerization domains III and IV

(reviewed in Chapter 1.2.5; Fig. 1.2; Krogan, 2006) were tested to determine the influence of

three genotypes on shoot regeneration in vitro. All genotypes used in the experiments were in the

Col-0 background and were compared to the Col-0 wild type. Explants of different organs

(cotyledons, leaves, petioles, hypocotyls, and adventitious roots) were first cultured on CIM for 6

days and then transferred to SIM (Chapter 2.8). The frequency of shoot regeneration was based

on the pooled data of all replicas of the experiments, and the mean number of shoots per shoot

regenerating callus (mean ± SE) was compared among the genotypes after 21 days on SIM.

55

4.3 Results

4.3.1 Shoot regeneration in cotyledon derived calluses

Calluses derived from cotyledon explants were generally not capable of shoot regeneration, with

the notable exception of dMP calluses (Fig. 4.1 and 4.2 A). In all other genotypes, including wild

type, shoot regeneration was almost absent (Fig. 4.1 A). There could be more than one shoot per

shooting callus in dMP (5±0.49), but the small number of shooting calluses in the other

genotypes precluded any statistical evaluation of this feature (Fig. 4.1 B). Also, because of small

numbers, it was not feasible to statistically compare the mean number of shoots per shoot

regenerating callus among all the genotypes. Nevertheless, 2 shoots were scored in the wild type

single shoot regenerating callus, and 5 shoots in the nph4-1 shoot regenerating callus. The mean

number of shoots in the mpG12 calluses was 1.6±0.6, range 1 to 4 in 5 shoot regenerating

calluses (Fig. 4.1 B). The mean number of shoots in the dMP calluses was relatively high

(5±0.49, range 1 to 20 in 171 calluses) (Fig. 4.1 B).

4.3.2 Shoot regeneration in leaf derived calluses

Calluses derived from leaf explants showed a clear difference among the genotypes in their

ability for shoot regeneration (Fig. 4.1 and 4.2 B). Generally, shoot regeneration in calluses

derived from leaf explants was stronger in all tested genotypes compared to those in cotyledon

derived calluses (Fig. 4.1 A). In wild type, 24% of the calluses generated shoots (59 out of 245,

where n represents the total number of explants in five independent experiments). Shoot

regeneration in nph4-1 was 21% (50 out of 241 calluses), which was only slightly less than that

in wild type (Fig. 4.1 A). Shoot regeneration in mpG12 was only 9.5% (17 out of 179 calluses),

which was much lower than that in wild type, and also in nph4-1 (Fig. 4.1 A). Whereas shoot

56

0 20 40 60 80 100

roots

hypocotyls

petioles

leaves

cotyledons

Frequency of shoot regenerating calluses, %

wtdMPnph4-1mpG12

A

n=173n=171

n=132n=132

n=245n=264

n=241n=179

n=108n=148

n=140n=118

n=153n=145

n=147

n=247n=254

n=238n=228

shoot regenerating calluses, %

0 2 4 6 8 10

roots

hypocotyls

petioles

leaves

cotyledons

# shoots per shoot regenerating callus

wtdMPnph4-1mpG12

B

n=63

n=5

n=59n=229

n=50n=17

n=22n=105

n=22n=9

n=97n=127

n=109

n=120n=67

n=70n=15

Figure 4.1 Percentage of shoot regenerating calluses (A), and mean number of shoots per shoot

regenerating callus (B) after 21 days on SIM (following 6 days on CIM to produce the calli).

In (A) “n” is the total number of explants used in the experiments; in (B) “n” is the number of

shoot regenerating calluses. Bars indicate SE.

57

A

dMP

mpG12

wt

nph4-1

cotyledons

B

dMPwt

nph4-1mpG12

leaves

Figure 4.2 Calluses derived from cotyledon (A) and leaf (B) explants after 21 days on SIM

(following 6 days on CIM to produce the calli). Genotypes indicated within plate sectors; arrows

indicate shoot regenerating calluses.

58

regeneration in mpG12 was reduced, it was very high in dMP (87%, or 229 out of 264 calluses)

(Fig. 4.1 A).

The number of shoots per shoot regenerating callus also showed differences in shoot

regeneration among the genotypes (Fig 4.1 B). In wild type, the mean number of shoots per

shoot regenerating callus was 2.2±0.22, range 1 to 8 in 59 calluses. The mean number of shoots

was different in nph4-1 and in mpG12. In nph4-1 calluses, the mean number of shoots per shoot

regenerating callus was 2.1±0.21, range 1 to 7 in 50 calluses. This was comparable to that in wild

type (Fig. 4.1 B). However, mpG12 calluses had only 1.1±0.08 shoots per callus, range 1 to 2 in

17 mpG12 calluses (P<0.001 for the comparison of the mean responses between mpG12 and wild

type, and between mpG12 and nph4-1 calluses) (Fig. 4.1 B). In contrast, in dMP calluses, the

mean number of shoots was much higher than that in wild type (6.2±0.27, range 1 to 24 in 229

calluses, P<0.001) (Fig. 4.1 B).

Collectively, these data indicate a strongly elevated shoot regeneration capability in dMP

calluses derived from leaf explants compared to that in wild type. This was indicated by a

difference in the proportion of shoot generating calluses, and in the mean number of shoots

formed by shoot regenerating calluses. The arf mutant genotypes, nph4-1 and mpG12, were very

different in their shoot regeneration ability. Shoot regeneration in nph4-1 was relatively similar

to that in wild type, whereas shoot regeneration in mpG12 was much lower than that in wild

type. These differences between the arf mutants were indicated by both the frequency of the

shoot regenerating calluses, and the mean number of shoots per callus.

4.3.3 Shoot regeneration in calluses derived from petiole explants

Calluses derived from petiole explants showed clear genotype related difference in their ability

for shoot regeneration on SIM (Fig. 4.1 and 4.3 A). In addition, their shoot regeneration profile

generally resembled that of leaf derived calluses (Fig. 4.1). In wild type, 20% of calluses (22 out

of 108) regenerated shoots (where n represents the total number of explants in four independent

replicas of the experiment). Shoot regeneration in the arf mutants, nph4-1 and mpG12, was

generally lower than that in wild type, but distinctly different in these two genotypes. Thus, shoot

regeneration in nph4-1 was slightly less than that in wild type, and 16%, or 22 out of 140

59

petioles

A

dMP

mpG12

nph4-1

wt

B

dMP

nph4-1

wt

hypocotyls

Figure 4.3 Calluses derived from petiole (A) and hypocotyl (B) explants after 21 days on SIM

(following 6 days on CIM to produce the calli). Genotypes indicated within plate sectors; arrows

indicate shoot regenerating calluses.

60

calluses, regenerated shoots (Fig. 4.1 A). Shoot regeneration in mpG12 was much lower than that

in wild type, and only 8%, or 9 out of 118 calluses, developed shoots (Fig. 4.1 A). Once again,

shoot regeneration in dMP was very strong compared to that in wild type, and a majority of dMP

calluses (71%, or 105 out of 148) formed shoots (Fig. 4.1 A).

The mean number of shoots per shoot regenerating callus also showed differences in shoot

regeneration ability among the genotypes (Fig. 4.1 B). In wild type, the mean number of shoots

per shoot regenerating callus was 2.7±0.42, range 1 to 7 in 22 calluses. The mean number of

shoots was significantly lower in nph4-1 and mpG12 calluses compared to that in wild type. In

nph4-1, the mean number of shoots was 1.8±0.28, range 1 to 6 in 21 calluses (P<0.1) (Fig. 4.1

B). The mpG12 calluses had 1.4±0.43 shoots per callus, range 1 to 5 in 9 calluses (P<0.05) (Fig.

4.1 B). In this category of calluses, the mean number of shoots in nph4-1 and mpG12 did not

differ significantly. In dMP calluses the mean number of shoots per shoot regenerating callus

was 5.9±0.39, range 1 to 24 in 230 calluses (P<0.001) (Fig. 4.1 B).

Collectively, these data showed strongly elevated shoot regeneration ability in dMP calluses

derived from petiole explants. This was indicated by the higher frequency of shoot regenerating

calluses, and by the higher number of shoots formed in the shoot regenerating calluses. Shoot

regeneration in the arf mutant genotypes, nph4-1 and mpG12, was generally lower than that in

wild type. Shoot regeneration was slightly decreased in the nph4-1 calluses, and greatly

decreased in the mpG12 calluses. Thus, the presumably elevated MP function in the dMP

calluses resulted in greatly increased shoot regeneration. Further, the loss of the MP function in

the mpG12 mutants resulted in greatly decreased shoot regeneration. Meanwhile, the loss of the

NPH4 function in the nph4-1 mutants had a more moderate effect on shoot regeneration, when

the calluses derived from petiole explants were compared on SIM.

4.3.4 Shoot regeneration in calluses derived from hypocotyl explants

The genotype related difference in shoot regeneration ability was relatively moderate when

calluses derived from hypocotyl explants were compared on SIM (Fig. 4.1 and 4.3 B). Generally,

shoot regeneration in calluses derived from these explants was strongest among all the types of

explants (cotyledon, leaf, petiole, and adventitious roots) (Fig. 4.1). All the tested genotypes

(wild type, dMP, and nph4-1) had a relatively high rate of shoot regeneration on SIM (Fig. 4.1).

61

No mpG12 calluses could be tested in this category, because of the lack of hypocotyls in the

mpG12 seedlings. In wild type, 63%, or 97 out of 153 calluses, regenerated shoots (where n

represents the total number of explants in four independent replicas of the experiment). Shoot

regeneration in nph4-1 was slightly higher than that in wild type, and 74%, or 109 out of 147

nph4-1 calluses, formed shoots (Fig. 4.1 A). Once again, shoot regeneration in dMP calluses was

stronger compared to that in wild type, and a majority of dMP calluses (88%, or 127 out of 145)

produced shoots (Fig. 4.1 A).

The mean number of shoots per shoot regenerating callus indicated the difference in shoot

regeneration ability among the genotypes more clearly (Fig. 4.1 B). In wild type, the mean

number of shoots per shoot regenerating callus was 4.14±0.23, range 1 to 10 in 97 calluses. The

mean number of shoots was significantly less in the nph4-1 calluses compared to that in the wild

type ones (3.4±0.19 shoots per callus, range 1 to 10 in 109 calluses, P<0.05). In the dMP

calluses, the mean number of shoots was much higher than that in the wild type calluses

(7.68±0.39, range 1 to 24 in 127 calluses, P<0.001).

Collectively, these data showed the increase in shoot regeneration in dMP calluses derived from

hypocotyl explants. This was indicated by a greater frequency of shoot regenerating calluses, and

by a higher mean number of shoots formed by the shoot regenerating calluses. Shoot

regeneration in the arf mutant genotype nph4-1 gave mixed results. On the one hand, the

frequency of shoot regenerating calluses in nph4-1 was higher than that in wild type. On the

other hand, the average number of shoots per shoot regenerating callus was significantly lower

than that in wild type. Thus, when compared to wild type, dMP calluses again showed increased

shoot regeneration on SIM. nph4-1 calluses demonstrated a moderate increase in the frequency

of shoot regenerating calluses, which produced significantly fewer shoots per callus compared to

those in wild type.

4.3.5 Shoot regeneration in calluses derived from adventitious root explants

Shoot regeneration in calluses derived from adventitious root explants partly differed from shoot

regeneration profiles in other types of explants (cotyledon, leaves, petioles, and hypocotyls) (Fig.

4.1 and 4.4). In this category of calluses, the strongest shoot regeneration was observed in wild

type, where 49%, or 120 out of 247 calluses, regenerated shoots (where n represents the total

62

dMP

mpG12

wt

nph4-1

adventitious roots

Figure 4.4 Calluses derived from adventitious root explants after 21 days on SIM (following 6

days on CIM to produce the calli). Genotypes indicated within plate sectors; arrows indicate

shoot regenerating calluses.

63

number of explants in four independent replica of the experiment). Shoot regeneration in the arf

mutants, nph4-1 and mpG12, was less than that in wild type, and also differed between these arf

genotypes. Thus, in nph4-1, 29%, or 70 out of 238 calluses, regenerated shoots, while in mpG12

only 7%, or 15 out of 228 calluses, did so (Fig. 4.1 A). This category of calluses was the only

one where the frequency of shoot regeneration in the dMP calluses was lower than that in the

wild type calluses. Only 26%, or 67 out of 254 dMP calluses, formed shoots (Fig. 4.1 A).

The mean number of shoots per shoot regenerating callus also indicated differences among the

genotypes (Fig. 4.1 B). In wild type, the mean number of shoots per shoot regenerating callus

was 3.1±0.19, range 1 to 10 in 120 calluses. The mean number of shoots was significantly less in

the nph4-1 and mpG12 calluses compared to that in the wild type calluses. In nph4-1 calluses,

the mean number of shoots was 2.25±0.22, range 1 to 12 in 70 calluses (P<0.01) (Fig. 4.1 B).

The mean number of shoots per shoot regenerating callus was even less in the mpG12 calluses

(1.7±0.24, range 1 to 4 in 15 calluses, P<0.001) (Fig. 4.1 B). In the dMP calluses, the number of

shoots per shoot regenerating callus was slightly and insignificantly higher than that in the wild

type calluses (3.5±0.51 shoots per callus). However, the maximum number of shoots scored in

the dMP calluses was more than twice that in the wild type calluses, ranging from 1 to 23 in 67

dMP shoot regenerating calluses (Fig. 4.1 B).

Thus, the dMP transgene did not affect shoot regeneration in calluses derived from adventitious

root explants. However, once again, the loss of the MP function in the mpG12 mutants resulted

in strongly decreased shoot regeneration. The loss of the NPH4 function in nph4-1 mutants also

resulted in decreased shoot regeneration, yet had a less prominent effect on shoot regeneration

than the loss of MP function. This is based on the comparison on SIM of calluses derived from

adventitious root explants.

4.3.6. Effect of duration of CIM preculture on shoot regeneration on SIM in

calluses derived from adventitious root explants

To determine the effect of the duration of CIM preculture on subsequent shoot regeneration on

SIM, adventitious root explants of the tested genotypes were precultured on CIM for different

periods of time (6, 9, 12, or 16 days on CIM), and then transferred on SIM for shoot

regeneration. The results of the experiment showed genotype related changes in the efficiency of

64

0

10

20

30

40

50

60

70

80

90

100

wt 6d n=

72

wt 9 d

n=35

wt 12d n

=22

wt 16 d

n=12

dMP 6d n=73

dMP 9d n

=34

dMP 12d

n=24

dMP 16d n=

14

nph4-1

6d n=67

nph4-1 9d

n=36

nph4

-1 12d n=25

nph4

-1 16d n=12

mpG12 6d n

=68

mpG12 9d n=32

mpG12 12d n

=24

mpG12 16 n=

12

Freq

uenc

y of

sho

ot re

gene

ratin

g ca

lluse

s, %

A

0

2

4

6

8

10

12

14

16

wt 6d n

=47

wt 9d n

=31

wt 12d

n=6

wt 16d

n=1

dMP 6d

n=18

dMP 9d

n=29

dMP 12

d n=13

dMP 16

d n=5

nph4

-1 6d n=

22

nph4

-1 9d n=

29

nph4

-1 12d n=

17

nph4

-1 16d n=

4

mpG12

6d n=11

mpG12

9d n=5

mpG12

12d n=2

mpG12

16d n=1

# sh

oots

per

sho

ot re

geer

atin

g ca

llus

B

Figure 4.5 Percentage of shoot regenerating calluses (A) and mean number of shoots per shoot regenerating callus (B) in calluses derived from adventitious root explants after 21 days on SIM depending on the time on CIM (6, 9, 12 and 16 days).

In (A) “n” represent the total number of explants used in the experiments; in (B) “n” is the number of shoot regenerating calluses. Bars indicate SE.

65

shoot regeneration depending on the time on CIM (Fig. 4.5). The results of a single experiment

are presented below.

In the experiment, when the adventitious root explants were exposed to CIM for 6 days (a

standard protocol, see Materials and Methods, Chapter 2.8), shoot regeneration was strongest in

the wild type calluses, and 64%, or 46 out of 72 calluses, formed shoots. Shoot regeneration

among all other genotypes was much lower compared to that in wild type. Specifically, shoot

regeneration in the arf mutants, nph4-1 and mpG12, was lower than that in wild type. Thus, 33%

(22 out of 67) nph4-1 calluses, and only 16% (11 out of 68) mpG12 calluses regenerated shoots

(Fig. 4.5 A). In dMP, the frequency of shoot regenerating calluses was also lower than that in

wild type, and only 25% (18 out of 73 calluses) formed shoots (Fig. 4.5 A).

The number of shoots per shoot regenerating callus differed slightly among the genotypes, where

the only significant difference was detected between wild type and mpG12. In wild type, the

mean number of shoots per shoot regenerating callus was 2.55±0.28, range 1 to 9 in 46 calluses.

In dMP, the mean number of shoots was 4.38±1.3, range 1 to 20 in 18 calluses. In nph4-1, the

mean number of shoots was 2.9±0.54, range 1 to 12 in 22 shoot regenerating calluses (Fig. 4.5

B). In mpG12, the mean number of shoots was significantly less than that in wild type (1.64±0.3,

range 1 to 4 in 11 calluses, P<0.05) (Fig. 4.5 B).

When the time on CIM was extended to 9 days, the shoot regeneration profile changed

significantly. Generally, shoot regeneration increased, and became relatively similar in all the

genotypes. An exception was mpG12, where shoot regeneration stayed consistently low

(Fig.4.5). At 9 days, 86%, or 30 out of 35 wild type calluses; 85%, or 29 out of 34 dMP calluses;

81%, or 29 out of 36 nph4-1 calluses; and only 16%, or 5 out of 32 mpG12 calluses, regenerated

shoots (Fig. 4.5 A). Also, in this experimental condition, the mean number of shoots per shoot

regenerating callus generally increased in all the genotypes, except for mpG12. In wild type, the

mean number of shoots per shoot regenerating callus was 4.87±0.5, range 1 to 12 in 31 calluses.

In dMP, the mean number of shoots was 13.31±1.23, range 1 to 26 in 29 calluses (P<0.001). In

nph4-1, the mean number of shoots was 7.1±0.78, range 1 to 16 in 29 calluses (P<0.05) (Fig.4.5

B). In mpG12, the mean number of shoots per shoot regenerating callus was significantly less

than that in wild type (1.2±0.19, range 1 to 2 in 5 calluses, P<0.001) (Fig. 4.5 B).

66

Extension of the time on CIM to 12 days affected shoot regeneration in the genotypes differently.

In wild type, shoot regeneration was strongly reduced compared to that at 9 days on CIM, and

only 27% (6 out of 22) wild type calluses regenerated shoots. Shoot regeneration was also

reduced in the dMP and nph4-1 calluses. However, these reductions were not as strong as in the

wild type calluses. Specifically, 54% (13 out of 24) dMP calluses, and 68% (17 out of 25) nph4-

1 calluses regenerated shoots (Fig. 4.5 A). Shoot regeneration in mpG12 calluses was also lower,

with only 8%, or 2 out of 24 calluses, regenerating shoots (Fig. 4.5 A).

The mean number of shoots per shoot regenerating callus was also reduced compared to that of

the previous experimental settings (9 days on CIM) in all the genotypes, except mpG12. In wild

type, the mean number of shoots per shoot regenerating callus was 3±1, range 1 to 8 in 6

calluses. The mean number of shoots per shoot regenerating callus was significantly higher in

dMP compared to that in wild type (9.85±2.27, range 1 to 30 in 13 calluses, P<0.05) (Fig. 4.5 B).

The mean number of shoots in nph4-1 was insignificantly higher than that in wild type

(4.88±0.7, range 1 to 10 in 17 calluses) (Fig. 4.5 B). The number of shoots scored in 2 out of 24

mpG12 shoot regenerating calluses was 1 and 2, which was consistent with all the data for shoot

regeneration in this genotype (Fig. 4.5 B).

A further drop in shoot regeneration was observed when the time on CIM was extended to 16

days (Fig. 4.5). Only 8%, or 1 out of 12 wild type calluses, regenerated shoots. A decrease in

shoot regeneration was also detected in dMP and nph4-1. However, shoot regeneration in these

genotypes was still higher than that in wild type. Only 36%, or 5 out of 14 dMP calluses, and

33%, or 4 out of 12 nph4-1 calluses, were still capable of shoot regeneration. Shoot regeneration

ability in mpG12 did not change, and 8%, or 1 out of 12 calluses, regenerated shoots.

Because of a generally low number of shoot regenerating calluses, a statistical analysis could not

be performed to discriminate the differences in the mean number of shoots per shoot

regenerating callus in this category. In wild type, the single shoot regenerating callus produced 2

shoots. In dMP, the mean number of shoots was 5.4±2.4, range 1 to 10 in 5 calluses (Fig. 4.5 B).

In nph4-1, the mean number was 2±0.4, range 1 to 3 in 4 calluses (Fig. 4.5 B). In mpG12, the

single shoot regenerating callus produced one shoot (Fig. 4.5 B).

67

Taken together, these data indicate that the increased time on CIM affected shoot regeneration in

wild type, dMP, and nph4-1, but not in mpG12. While the extension of CIM preculture from 6 to

9 days generally had a stimulating effect on shoot regeneration in wild type, nph4-1, and dMP,

the longer extension (12 or more days on CIM) inhibited shoot regeneration on SIM.

Interestingly, dMP and nph4-1 were less sensitive to the increased time of CIM preculture, while

shoot regeneration in wild type dropped sharply after 9 days on CIM. Thus, the efficiency of

shoot regeneration underwent dynamic changes in response to the duration of CIM preculture in

wild type, dMP, and nph4-1.However, shoot regeneration was consistently low and relatively

independent of the duration of CIM preincubation in mpG12 calluses derived from adventitious

root explants.

4.3.7 Greening of calluses derived from adventitious root explants on SIM

Genotype related differences were detected in the timing of greening--when adventitious root

calluses, originally whitish after CIM, turned green on SIM. After transfer to SIM, the calluses

increased in size, and turned green. The period of time required for greening was clearly

different in the genotypes. In wild type, the first evidence of greening appeared after 5 days on

SIM. This was visualized as green spots embedded in the whitish color of the calluses. Later,

these green areas increased in size, and, after 10 days on SIM, the whole callus turned green

(Fig. 4.6). In the intermediate stage (8 days on SIM), wild type calluses usually had a mixed

color (green and whitish). About 56% of wild type calluses (29 out of 52) were mostly whitish

(the whitish color covered more than 50% of the area), while the rest, 44% (23 out of 52), were

mostly green (more than 50% of the callus area was green) (Fig. 4.7). A similar picture was

observed for the nph4-1 calluses, with different proportions between the whitish and greenish

calluses. 81% of the nph4-1 calluses (42 out of 52) were mostly whitish, while the rest (19%, or

10 out of 52) were mostly green (Fig. 4.7). The dMP and mpG12 calluses formed two distinct

phenotypic groups. Most of the dMP calluses were clearly whitish (98%, or 55 out of 56 calluses

in total). However, most of the mpG12 calluses (98%, or 50 out of 51 calluses) were bright green

in color (Fig. 4.6; 4.7).

Thus, the mpG12 calluses were the ones which turned green on SIM faster, more evenly and

intensively than the calluses of wild type and other genotypes. Greening of the dMP calluses was

delayed, and it was less intensive compared to those in wild type. Greening of the nph4-1

calluses was slightly less intensive compared to that of the wild type ones. No genotype related

68

dMPwt

mpG12 nph4-1

A

adventitious roots, 8 d SIM

mpG12

dMP

nph4-1

wt

dMP

mpG12nph4-1

wt

adventitious roots, 13 d SIM

B

Figure 4.6 Greening of calluses and root outgrowth in calluses derived from adventitious root

explants on SIM after 8 (A), and 13 (B) days on SIM (following 6 days on CIM to produce the

calli). Genotypes indicated within plate sectors.

69

0102030405060708090

100

wt n=52 dMP n=56 nph4-1 n=52 mpG12 n=51

frequ

ency

, %

lightgreen

wtn=52

dMPn=56

nph4-1n=52

mpG12n=51

roots, 8 days on SIM

perc

enta

ge, %

Figure 4.7 Percentage of green versus whitish calluses after 8 days on SIM in adventitious root

derived calluses; n, number of explants.

70

difference in greening was established for calluses derived from other organs than roots, because

they were already greenish at the time of their transfer on SIM.

4.4. Discussion and Conclusions

The results in this chapter indicate that the efficiency of shoot regeneration depends on the

genotype, the type of organ used as the explant, and the duration of preculture on CIM. As

shown earlier (Akama et al., 1992; Chatfield and Raizada, 2008), the Columbia ecotype of

Arabidopsis thaliana is relatively recalcitrant to shoot regeneration, especially when cotyledons

were used to induce shoot regeneration in vitro. In the experiments here, shoot regeneration in

wild type varied dramatically depending on the type of organ used as the explant. In wild type,

the frequency of shoot regenerating calluses was negligible in cotyledon derived calluses,

moderate in leaf and petiole derived calluses, and relatively high in hypocotyl and adventitious

root derived calluses. A similar dependence of shoot regeneration efficiency on the type of

explant was clearly detected in nph4-1 calluses. However, shoot regeneration in dMP and

mpG12 calluses derived from these explants differed from those in wild type. Shoot regeneration

in mpG12 was very low regardless of the organ derivation of the callus. In contrast, shoot

regeneration in dMP was strong in calluses derived from aerial organs (cotyledons, leaves,

petioles, and hypocotyls), where it was significantly and consistently higher than that in wild

type. No elevation of shoot regeneration was recorded in the dMP calluses derived from

adventitious root explants.

These data indicate that the presumably elevated MP function in dMP promotes shoot

regeneration in calluses derived from aerial explants. Consistent with this, loss of MP function in

mpG12 mutants lead to greatly reduced shoot regeneration. This was indicated by the low

frequency of shoot regenerating calluses, and by the low number of shoots per callus in mpG12.

Loss of NPH4 function in nph4-1 mutants had a relatively modest effect on shoot regeneration.

The frequency of shoot regenerating calluses and the number of shoots per callus were only

slightly lower in nph4-1 compared to those in wild type.

MP/ARF5 and NPH4/ARF7 have a sequence similarity, interact with each other, and have

partially redundant functions (Liscum and Reed, 2002; Hardtke et al., 2004; discussed in General

Introduction, Chapter 1.2.5). As shown here, during de novo shoot initiation, the NPH4 gene

71

could not completely substitute for the loss of MP function. The MP gene is implicated in the

regulation of several key shoot meristem genes, such as CUC2 and STM (Aida et al., 2002), and

the ESR1/DRN gene (Cole et al., 2009). Moreover, MP is involved in the control of cytokinin

signaling through the regulation of activity A-type ARR15 and ARR7 (Zhao et al., 2010). The

meristem genes, such as CUC, STM, and ESR1/DRN, and some of the ARR genes are implicated

in de novo shoot regeneration (reviewed in Chapter 1.6.1; Hwang and Sheen, 2001; Osakabe et

al., 2002; To et al., 2004; Zhang and Lemaux, 2005). Thus, the observed strong impairment of

shoot regeneration in mpG12 mutants, as well as the consistently elevated shoot regeneration in

dMP calluses derived from aerial explants, may be explained by changes in the expression levels

of these genes in the mpG12 and dMP genotypes. The failure of dMP to promote shoot

regeneration from root derived calluses may be explained by the fact that MP, and hence dMP,

are not normally expressed in mature roots (Dr. W. Ckurshumova, personal communication).

However, low shoot regeneration in mpG12 may contradict the suggestion of the irrelevance of

MP to shoot regeneration in root derived calluses. Future studies are required to determine

whether dMP expression in root derived calluses is similar to that in calluses derived from other

organs.

No data demonstrating the involvement of NPH4 in the regulation of shoot meristem genes were

found in the literature. Instead, it has been shown that NPH4/ARF7 plays a central role in the

modulation of auxin-dependent differential growth in the aerial parts of Arabidopsis plants

(Stowe-Evans et al., 1998, Watahiki et al., 1999; reviewed in Chapter 1.2.5). Auxin insensitivity

has been shown for aerial organs, but not for the roots of nph4 mutants (Watahiki and

Yamamoto, 1997). However, here shoot regeneration in nph4-1 calluses derived from

adventitious root explants, with presumably normal auxin sensitivity and callusogenesis (Chapter

3.3.6), was significantly lower than that in the wild type calluses. This could reflect the partially

redundant function that NPH4 has with MP in forming the apical part of embryos, including the

shoot meristem (Hardtke et al., 2004). The contribution of NPH4 could be subtle, and completely

dispensable, in calluses cultured for more than 9 days.

In summary, decreased MP/ARF5 and NPH4/ARF7 levels in mpG12 and nph4-1 mutants had a

different impact on de novo shoot regeneration in these genotypes. While shoot regeneration in

nph4-1 was only slightly decreased compared to that in wild type, shoot regeneration in mpG12

was consistently, and markedly, reduced in calluses derived from any type of explant in all sets 72

of experiments. In contrast, elevated MP activity in dMP had a strong stimulating effect on shoot

regeneration when aerial organs were used as explants. Thus, MP played an important role in de

novo shoot meristem initiation.

73

Chapter 5

Root regeneration under conditions of altered auxin signal

transduction


In tissue culture, regeneration of adventitious roots occurs in response to a high auxin to low

cytokinin ratio, or when auxin is present as a sole hormone in the root inducing medium (RIM)

(Miller and Skoog, 1957; Gaba, 2005). Rooting is often a critical step in the plant propagation

industry, because many species are extremely recalcitrant regarding adventitious root formation

(de Klerk, 2002; Schwarz et al., 2005). Although adventitious roots have been routinely induced

in response to auxin (Callis, 2005; Schwarz et al., 2005), the molecular mechanisms of this event

are unclear (Schwarz et al., 2005; Rose et al, 2006).

In tissue culture, the acquisition of competence by cells for either shoot or root formation occurs

during the preincubation stage on CIM (Christian and Warnick, 1985; Che et al., 2007) - the

stage when promeristemoid structures are formed (Schwarz et al., 2005). The specification of

organ development depends on subsequent hormonal stimuli on SIM or RIM. Ozawa et al.

(1998) - using the temperature-sensitive mutants, srd1, srd2 and srd3 -demonstrated that the

competence to form organs is acquired progressively, first roots and then shoots (Ozawa et al.,

1998).

In vitro experiments with leaf explants have shown that root primordia are formed from the

procambial-like cells of veins. These function as pluripotent stem cells, with the ability to form

either root primordia or vascular tissues in response to added auxin (Attfield and Evans, 1991;

Rose et al., 2006). Unlike the hormone auxin, cytokinin inhibits root formation in planta (Aloni

et al., 2006) and in vitro (Sugiyama 1999; Kubo and Kakimoto, 2000). Auxin insensitivity causes

impairment in root regeneration in culture. The auxin-tolerant rac mutants in tobacco are

defective in primary root formation and the regeneration of adventitious roots (Sugiyama, 1999,

and references therein). By contrast, cytokinin insensitivity promotes root and root primordia

formation even on cytokinin containing media (Kubo and Kakimoto, 2000; Kakani et al., 2009).

Che et al. (2006) demonstrated that molecular signatures for callus formation and root

74

regeneration are quite similar, since both are triggered by auxin-rich media. They also showed

that competence for root regeneration correlates with the increased expression of IAA genes on

CIM.

Besides their other features (for details see Chapter 1.2.5), ARF5/MP and ARF7/NPH4 are

implicated in root formation. However, mp seedlings lack primary roots (Berleth and Jurgens,

1993). Increased auxin responses and increased adventitious root formation have been shown for

cotyledon explants of the GOF MP line, dMP (Krogan et al., 2006). However, root formation

from cultured cotyledons of mp mutants is strongly decreased (Mattsson et al., 2003). Also, mp

mutants show auxin insensitivity and reduction of DR5 expression in leaf primordia (Mattsson et

al., 2003). The nph4 mutants exhibit auxin resistance in the aerial parts of plants. They have

severely impaired expression of many primary auxin-dependent genes and the auxin-responsive

reporter DR5 (Watahiki and Yamamoto, 1997; Stowe-Evans et al., 1998; Wang et al., 2005).

Also, NPH4, together with ARF19, regulates lateral and adventitious root formation (Tatematsu

et al., 2004; Wilmoth et al, 2005; Okushima et al., 2007). Wilmoth et al. (2005) showed that, in

the presence of exogenous IAA, the excised nph4 roots formed fewer lateral roots. The nph4

excised hypocotyls formed fewer adventitious roots than did wild type roots and hypocotyls. By

contrast, nph4 arf19 double mutants mainly lacked lateral roots.

In this chapter, the roles of the MP and NPH4 genes in de novo root regeneration in tissue culture

are explored. The extent to which root regeneration in each of the investigated genotypes is

further influenced by the origin of the respective tissue sample is also explored.


This project investigates the possible influence on de novo root regeneration of auxin signaling

through ARF5/MP and ARF7/NPH4. To this end, the loss-of-function mutants mpG12 (Hardtke

and Berleth, 1998), and nph4-1 (Harper et al., 2000), and the gain of function transgene line dMP

with truncated dimerization domains III and IV (reviewed in Chapter 1.2.5; Fig. 1.2; Krogan,

2006) were tested to determine the influence of these genotypes on adventitious root

regeneration in vitro. All the genotypes used in the experiments were in the Col-0 background

and were compared to the Col-0 wild type. Explants of different organs (cotyledons, leaves, and

adventitious roots) were first cultured on CIM for 6 days and then transferred to RIM (reviewed

75

in Chapter 2.8; Valvekens et al., 1988). Additionally, root formation in calluses derived from

adventitious root explants, and cultured on SIM after 6 days on CIM, was compared among the

genotypes. The frequency of root regeneration, a calculation based on the pooled data of all

replicas of the experiments, was compared among the tested genotypes after 21 days on RIM or

on SIM, as was the mean number of roots per root regenerating callus (mean +/- SE).

5.3 Results

5.3.1 Root regeneration in cotyledon derived calluses on RIM

There were some genotype related differences in root regeneration frequency, and in the mean

number of roots per root regenerating callus (Fig. 5.1; 5.2 A; 5.3). In wild type, only 67% of

calluses (63 out of 89) regenerated roots (Fig. 5.1 A). Among the rest of the genotypes, the

proportion of root regenerating calluses was higher than that in wild type, and relatively similar:

96% of dMP calluses (84 out of 88); 91% of nph4-1 calluses (76 out of 85); and, 93% of mpG12

calluses (71 out of 77) regenerated roots (Fig. 5.1 A).

The mean number of roots per root regenerating callus was quite similar among wild type, nph4-

1, and mpG12 calluses. It was significantly higher only in dMP (Fig. 5.1 B). In wild type, the

mean number of roots was 8.6±0.69, range 2 to 25 roots per calluses in 63 root regenerating

calluses. In nph4-1, the mean number of roots was 8.72±0.53, range 1 to 22 roots in 76 calluses.

In mpG12, this number was 8±0.5, range 1 to 20 in 71 calluses (Fig. 5.1 B). The mean number of

roots was significantly higher in dMP, equal to 17.3±0.82, range 3 to 35 in 84 root regenerating

calluses (P<0.001) (Fig. 5.1 B).

To summarize, in calluses derived from cotyledon explants, root regeneration in nph4-1 and

mpG12 was not impaired. It was either equal to that in wild type (in the mean number of roots

per root regenerating callus), or even exceeded that in wild type (in the frequency of root

regenerating calluses). Moreover, root regeneration in dMP calluses was significantly increased

compared to that in wild type.

76

0 10 20 30 40 50 60 70 80 90 100

adv.roots

leaves

cotyledons

Frequency of root regenerating calluses, RIM, %

w t

dMP

nph4-1

mpG12

0 5 10 15 20 25

roots

leaves

cotyledons

# roots per root regenerating callus, RIM

wtdMPnph4-1mpG12

B

A n=89n=88

n=85n=77

n=62n=67

n=69n=68

n=111n=99n=100

n=103

n=63n=84

n=76n=71

n=30n=63

n=40n=48

n=111n=99

n=100n=103

Figure 5.1 Percentage of root forming calluses (A), the mean numbers of roots per root forming

callus (B) in calluses derived from cotyledon, leaf and adventitious root explants after 21 days on

RIM (following 6 days on CIM to produce the calli);


root regenerating calluses. Bars represent SE.

77

A

cotyledons, RIM

dMP

mpG12

wt

nph4-1

dMP

mpG12

wt

nph4-1

leaves, RIM

B

Figure 5.2 Cotyledon (A) and leaf (B) derived calluses after 21 days on RIM (following 6 days

on CIM to produce the calli). Genotypes indicated within plate sectors; arrow in A points at a

callus without roots

78

A B

C D

wt wt

dMP dMP

E F

G H

nph4-1 nph4-1

mpG12 mpG12

Figure 5.3 Root initiation in cotyledon explants after 6 days on RIM (following 6 days on CIM

to produce the calli).

Genotypes, bottom right, arrows indicate root outgrowth.

79

5.3.2 Root regeneration in leaf derived calluses on RIM

Root regeneration in calluses derived from leaf explants was generally lower than that in

cotyledon derived calluses (Fig. 5.1; 5.2 B). In wild type, only 48% (30 out of 62) calluses

regenerated roots. Once again, this frequency was the lowest one among the genotypes (Fig. 5.1

A). In the arf mutants, nph4-1 and mpG12, the frequency of root regenerating calluses was

higher than that in wild type: 58% of nph4-1 calluses (40 out of 69); and, 71% of mpG12

calluses (48 out of 68) regenerated roots (Fig. 5.1 A). The highest frequency of root regenerating

calluses was scored again in dMP, where 94% of calluses (63 out of 67) formed roots (Fig.

5.1A).

The mean number of roots per root regenerating callus was similar among wild type, nph4-1, and

mpG12 calluses (Fig. 5.1B). However, the mean number of roots was significantly higher in

dMP (Fig. 5.1 B). In wild type, the mean number of roots per root regenerating callus was

4.35±0.35, range 2 to 10 per callus in 30 calluses. In nph4-1, the mean number of roots was

4.3±0.31, range 2 to 8 in 40 calluses. In mpG12 the mean number was 4.2±0.26, range 2 to 9 in

48 calluses (Fig. 5.1 B). The mean number of roots was significantly higher in dMP, equal to

7.3±0.39, range 2 to 13 in 63 calluses (P<0.001) (Fig. 5.1 B).

To summarize, in leaf derived calluses, root regeneration in the arf mutants, nph4-1 and mpG12,

was not lower than that in wild type. However, in dMP, root regeneration was significantly

greater than that in wild type.

5.3.3 Root regeneration in calluses derived from adventitious roots on RIM

Since root explants are able to produce roots on RIM without CIM preincubation (Ozawa et. al,

1988), all explants in this experiment formed roots. The genotype related differences were

manifested only in the number of roots per root regenerating callus (Fig. 5.1; 5.4; 5.5). In wild

type, the mean number of roots was 22.12±1, range 2 to 50 in 111 calluses. The mean number of

roots in dMP calluses was comparable to that in wild type (19.86±0.95, range 1 to 45 in 99

calluses) (Fig. 5.1 B). By contrast, the mean number of roots per callus was significantly lower in

nph4-1 (11.57±0.59, range 2 to 30 in 100 calluses, P<0.001); and especially in mpG12 calluses

(7.99±0.42, range 1 to 18 in 103 calluses, P<0.001). The difference in the mean number of roots

per callus between nph4-1 and mpG12 was also statistically significant (P<0.001) (Fig. 5.1 B). 80

dMP

mpG12

nph4-1

wt

adventitious roots, RIM

Figure 5.4 Calluses derived from adventitious root explants after 21 days on RIM (following 6

days on CIM to produce the calli). Genotypes indicated within plate sectors.

81

In summary, quantitative differences in root formation in this category of explants occurred only

with regard to the number of roots per callus. In the arf mutant genotypes, the number of roots

per callus in nph4-1 was only half that of wild type; and in mpG12 it was even less. In dMP root

calluses--in contrast to dMP cotyledon and leaf derived calluses--no increase in root regeneration

was observed. The number of roots per callus was comparable in wild type and dMP.

5.3.4 Root formation in adventitious root explants on SIM

Genotype related differences in root initiation on RIM in adventitious root calluses were

expressed only in the number of roots per callus (Fig.5.1). In light of that finding, this study

explored whether greater distinctions would become apparent in the presence of a root

suppressing stimulus (Fig. 4.4; 5.6; 5.7). To that end, root formation in adventitious root calluses

on cytokinin-rich SIM was investigated. As one might expect, rooting on SIM was much less

than on RIM in all genotypes and was particularly low in mpG12 calluses (Fig. 5.1; 5.6). After

21 days on SIM, 87% of wild type calluses (139 out of 159), 98% of dMP calluses (157 out of

159), and 82% of nph4-1 calluses (124 out of 143) had formed roots (Fig. 5.6 A). However,

roots formed in only 24% of mpG12 calluses (39 out of 150) (Fig. 5.6 A).

The mean number of roots per root regenerating callus also differed among the genotypes. In

wild type, the mean number of roots per callus was 6.98±0.22, range 1 to 30 per callus in 139

calluses (Fig. 5.6 B). In dMP, the mean number was slightly higher than that in wild type

(8.75±0.3, range 1 to 32 in 157 calluses, P<0.1) (Fig. 5.6 B). In nph4-1, the mean number of

roots per callus (6.06±0.45, range 1 to 20 in 124 calluses) was similar to that in wild type (Fig.

5.6 B). In mpG12, the mean number of roots was the lowest compared to that in wild type

(1.9±0.42, range 1 to 5 in 39 mpG12 calluses, P<0.001, and to that in the other genotypes

(P<0.001) (Fig. 5.6 B).

In summary, compared to root formation on RIM, fewer roots were formed in all the genotypes

on SIM. However, root development was particularly suppressed in the mpG12 calluses, which

barely produced any roots on the cytokinin-rich SIM.

82

A

wt, 12 d RIM

C

dMP, 12 d RIM

B

D

wt, 21 d RIM

dMP, 21 d RIM

E

nph4-1, 12 d RIM

G

mpG12, 12 d RIM

F

H

nph4-1, 21 d RIM

mpG12, 21 d RIM

Figure 5.5 Calluses derived from adventitious root explants after 12 (A, C, E, G) and 21 (B, D,

F, H) days on RIM (following 6 days on CIM to produce the calli). Genotypes, and days on RIM,

bottom right.

83

0 2 4 6 8 10

# roots per callus, SIM

wtdMPnph4-1mpG12

0 20 40 60 80 100

Frequency of root regenerating calluses, SIM, %

wtdMPnph4-1mpG12

A

B

n=159

n=159

n=143

n=150

n=139

n=157n=124

n=39

Figure 5.6 Percentage of root regenerating calluses (A), and the mean numbers of roots per root

regenerating callus (B) in calluses derived from adventitious root explants after 21 days on SIM

(following 6 days on CIM to produce the calli)


root regenerating calluses. Bars represent SE.

84

A B

C D

wt, 8 d SIM wt, 13 d SIM

dMP, 8 d SIM dMP, 13 d SIM

E F

G H

nph4-1, 8 d SIM nph4-1, 13 d SIM

mpG12, 8 d SIM mpG12, 13 d SIM

Figure 5.7 Root outgrowth in calluses derived from adventitious root explants after 8 (A, C, E,

G) and 13 (B, D, F, H) days on SIM (following 6 days on CIM to produce the calli). Genotypes,

and days on SIM as indicated on bottom right.

85

5.4 Discussion and Conclusions

The results in this chapter show that root regeneration differed among the auxin signal

transduction genotypes, and was also influenced by the origin of the explant. Quite unexpectedly,

root regeneration responses were only partially correlated with the assumed level of auxin

sensitivity in the explants.

Root regeneration on RIM in cotyledon and leaf derived calluses was elevated only in the dMP

explants, yet undiminished in the arf mutant genotypes, nph4-1 and mpG12. Earlier Krogan

(2006) showed that dissected dMP cotyledons were highly competent for adventitious root

formation when they were cultured in liquid RIM containing IBA. By contrast, rooting of mp

cotyledons in similar conditions was greatly decreased compared to that in wild type (Mattsson

et al., 2003). The discrepancy between the results of this study and the previous data on root

regeneration in mpG12 cotyledon explants can possibly be explained by a difference in the

experimental methods of rooting. Culture on liquid RIM may trigger different developmental

processes compared to those after pre-incubation on CIM, during which competence for

organogenesis stimuli is acquired (Christian and Warnick, 1985; Che et al., 2007; see Chapter

2.8 for details).

Another possible explanation for the normal root regeneration here in the aerial explants of the

arf mutant genotypes, particularly mpG12, may be related to the different molecular properties of

the two auxins, 2,4-D in CIM, and IBA in RIM. IBA is one of the natural forms of auxin with

physiological properties similar to IAA (Rashotte et al., 2003; Gaba, 2005). 2,4-D is a powerful

synthetic auxin, used for callus initiation (Gaba, 2005), which is known to be a good influx and a

bad efflux substrate - the qualities which allow 2,4-D to accumulate intracellularly in high

concentrations (Delbarre et al., 1996; Estelle, 1998). Possibly, the high intracellular auxin (2,4-

D) concentration during CIM incubation diminished differences in the relative auxin responses

among the tested genotypes. Also, the genotype related sensitivity of the explants to these two

auxins can differ (Estelle, 1998).

All the calluses derived from the adventitious root explants produced roots on RIM. The calluses

of the arf mutant genotypes (nph4-1 and mpG12) had a lower mean number of roots per callus.

Differences in the mean number of roots per callus were the only discriminative phenotypical

86

trait found in this category of explants. Also, lower root numbers in the arf mutants were only in

partial agreement with the original levels of auxin sensitivity in root explants, where low auxin

sensitivity is known for mpG12 (Mattsson et al., 2003), but not for nph4-1 roots (Watahiki and

Yamamoto, 1997; Okushima et all., 2005).

Another explanation for the lower root formation on RIM in the arf root derived calluses may be

their genotype related ability for lateral root formation. Root development in root derived

calluses is initiated from the pericycle of the root explants, i.e., it is similar to lateral root

formation (Casimiro et al., 2003; Atta et al., 2009; Sugimoto et al., 2010 ). Auxin stimulates the

development of lateral roots (Casimiro et al., 2003; De Smet et al., 2010). Because of that, the

lower root numbers on RIM found in nph4-1 and mpG12 could reflect their ability for lateral root

formation in response to auxin. As shown recently, ARF7/NPH4 and ARF5/MP are components

of a bimodular organogenesis system, and both ARFs are required for proper organogenesis and

lateral root formation in response to auxin (De Smet et al., 2010). LOF of these ARFs can lead to

reduced lateral root formation; and possibly, to lower number of roots in the mpG12 and nph4-1

adventitious root explants on RIM.

In this study, calluses derived from adventitious roots on SIM, had abundant root formation in

most of the wild type, nph4-1, and dMP calluses--although in lesser numbers compared to that

on RIM. However, root formation was dramatically reduced in mpG12. This was in broad

correlation with known root auxin sensitivities, where mpG12, the only genotype with reduced

auxin sensitivity in roots, also had greatly reduced root formation (Watahiki and Yamamoto,

1997; Mattsson et al., 2003; Krogan, 2006). Cytokinin inhibits root formation (Aloni et al.,

2006). When roots appear on cytokinin-rich SIM after CIM, this occurs against the external

hormonal signal. The latter is overcome by the inherent ability of roots to produce lateral roots,

which is normally an auxin-dependent process (Casimiro et al., 2003; Hardtke, 2006). However,

in the results here, the difference in genotype related auxin sensitivity comes out more clearly on

SIM, rather than on RIM. The reasons for this are as yet unclear. To attempt to clarify them,

further study with a wide range of mutants with altered auxin sensitivity would be required.

Also in this study, cytokinin counter activity on SIM was inversely correlated to auxin

sensitivity in the genotypes. In these results, low auxin sensitivity in mpG12 can also be paired

with elevated cytokinin sensitivity. The latter was also indicated by early and intensive greening

of mpG12 roots on SIM (discussed in Chapter 4.3.7). The lack of roots, or root primordia, and 87

the intensive greening of calluses on SIM are known as cytokinin responses (Sugiyama 1999;

Kubo and Kakimoto, 2000; Gaba, 2005). As shown recently, ARF5/MP is implicated in crosstalk

between auxin and cytokinin transduction pathways (Zhao et al., 2010; see for details Chapter

1.2.5). Thus, auxin/cytokinin balance and cytokinin sensitivity may be changed in the mpG12

explants due to the implication of MP in auxin/cytokinin crosstalk, and due to the interrelated

and antagonistic activity of these hormones in organogenesis (Coenen and Lomax, 1997;

Nordstrom et al., 2003; Zhao et al., 2010). These changes can result in visually noticeable

elevated cytokinin responses, and a lack of roots in the mpG12 root derived calluses on SIM.

88

Chapter 6

General Discussion and Conclusions

6.1 Effects of altered MP/ARF5 and NPH4/ARF7 activities on

callusogenesis and de novo generation of shoot and root

Plant cells have the ability to generate a whole plant from a wide variety of tissues and organs,

or even from a single cell (Skoog and Miller, 1957; Vogel, 2005; Sugimoto et al., 2010).

Several researchers have shown that auxin signaling genes may be involved in de novo shoot

and root formation (Coenen and Lomax, 1998; Gordon et al., 2007; Chatfield and Raizada,

2008; Zhao et al., 2008; Kakani et al., 2009). Auxin signaling operates through auxin response

factors (ARFs), a family of transcription factors that regulate the expression of auxin

responsive genes (reviewed in Teale, 2006; Guilfoyle and Hagen, 2007). In this research, the

LOF arf mutants, mpG12 and nph4-1, and the GOF transgene line dMP with truncated

regulatory domains III and IV (Krogan, 2006; Fig. 1.2) were tested to determine the roles of

altered MP and NPH4 activity on callus formation and de novo organogenesis in vitro. The

results of the study indicated genotype, and organ related differences during developmental

processes in plant regeneration.

Detailed discussion of the results related to each type of organogenesis is presented at the end

of each result chapter (3.4; 4.4; 5.4). The main focus of this section is the discussion and

general conclusions of the main results.

6.1.1 NPH4/ARF7 in developmental processes in tissue culture

The results of this study indicated that callus proliferation, root formation, and shoot

regeneration were diminished in some of the nph4-1 explants, though relatively normal in others

(Fig. 3.1; 4.1; 5.1). The observed variations in the organogenesis responses correlated with the

level of auxin sensitivity in the tested genotypes only during callus proliferation, but not always

during root and shoot formation.

Callusogenesis and root formation - developmental processes induced by auxin--have been

considered to share similar molecular pathways (Gaba, 2005; Che et al., 2006; Atta et al., 2009; 89

Sugimoto et al., 2010). However, in this study, callusogenesis and root regeneration in nph4-1

mutants were affected differently. For example, only nph4-1 aerial organs were defective in

callusogenesis, and only calluses derived from adventitious roots had fewer roots compared to

those in wild type. Interestingly, only aerial parts of nph4-1 plants have been known to be auxin

insensitive (Watahiki and Yamamoto, 1997; Stowe-Evans et al., 1998; Okushima et al., 2005).

The diminished level of auxin sensitivity in the nph4-1 aerial explants correlated with the

diminished callus proliferation, but it did not correlate with the ability for root regeneration (Fig.

3.1).

Callus proliferation depends on the activation of cell cycle genes in response to auxin and

cytokinin signals (Hemerly et al., 1993; Soni et al., 1995; Shaul et al., 1996; Riou-Khamlichi et

al., 1999; Sugiyama, 1999; Richard et al., 2002; Che et al., 2007). AuxREs elements have been

found in the promoters of various cyclin genes and some CDKs (Richard et al., 2002; Perrot-

Rechenmann, 2010). In that light, NPH4/ARF7 function in aerial organs might be related to the

auxin mediated activation of cell cycle genes.

The most straightforward interpretation of the apparently normal callusogenesis in nph4-1

adventitious roots is that auxin signal transduction is not defective in nph4-1 roots – because of

the presence of other ARFs, particularly ARF19, which can compensate for NPH4 function

specifically in this organ (Okushima et al., 2005). This interpretation is plausible, because it is

consistent with the observed absence of root defects, and normal root auxin sensitivity in nph4

single mutants (Watahiki and Yamamoto, 1997; Stowe-Evans et al., 1998; Okushima et al.,

2005). This is contrasted by the appearance of root specific defects in the nph4 arf19 double

mutants (Tatematsu et al., 2004; Okushima et al., 2005; Wilmoth et al, 2005; De Smet et al.,

2010).

Similarly, apparently normal de novo root development in nph4-1 aerial explants (cotyledons

and leaves) (Fig. 5.1) reflects the lack of an NPH4 requirement for de novo rooting in aerial

organs. This may be due to ARF redundancy in these organs (Hardtke et al., 2004; Tatematsu

et al., 2004; Wilmoth et al., 2005). By contrast, the results suggest that the molecular

mechanisms of root initiation in adventitious root explants are different from those in aerial

explants, as the latter seem to selectively require NPH4.

90

Shoot regeneration in nph4-1 calluses also reflected the origin of the explants. Shoot

regeneration was relatively mildly, or not at all, affected in explants from aerial organs. By

contrast, shoot regeneration in nph4-1 adventitious root explants was significantly less than that

in wild type (Fig.4.1). Thus, regarding the NPH4 requirement in organogenesis, the organ origin

of the nph4-1 explants determines the efficiency of de novo shoot, and root formation.

These organ related similarities in the efficiency of root and shoot regeneration may indicate that

these processes share a common developmental pathway. This statement accords with the model

in which shoot or root organogenesis is preceded by a state of tissue competence for either type

of organogenesis (Christianson and Warnick, 1985; Schwarz et al., 2005).

For example, tissue structures similar to lateral root meristems (LRMs) develop during callus

proliferation on CIM (Che et al., 2007; Atta et al., 2009; Sugimoto et al., 2010). Interestingly,

similar structures were also found in aerial explants when they were cultured on CIM (Sugimoto

et al., 2010). When transferred on RIM or SIM, LRM-like structures convert to either root or

shoot meristems (Che et al., 2007; Atta et al., 2009). Taking into consideration that lateral root

formation is slightly diminished in nph4-1 roots (Okushima et al., 2005; Wilmoth et al., 2005),

nph4-1 adventitious root explants may have fewer organogenesis-competent cells that are the

precursors of lateral root meristems. This may explain reduced shoot and root regeneration in

nph4-1 adventitious root explants. The organogenesis aspect of the NPH4 function was almost

irrelevant to the auxin sensitivity of nph4-1 organs, as measured by auxin sensitivity assays

(Watahiki and Yamamoto, 1997; Stowe-Evans et al., 1998; Okushima et al., 2005).

The variation in organogenesis productivity among the different organs and types of explants of

the same genotype is a well known fact in tissue culture research (Akama et al., 1992; Zhao et

al., 2002; Gaba, 2005). The question remains, however, of how calluses are able to maintain their

organ related identity, which causes the variations in organogenesis activity among calluses from

different explants. One explanation is the variation in endogenous hormone levels among plant

organs, which can affect their organogenesis responses in vitro (Davies, 2004; Gordon et al.,

2007). Also, the difference in epigenetic regulation may be a factor contributing to the variation

in organogenesis activity among plant organs (Zhao et al., 2008). Calluses generally comprise

heterogeneous mixtures of mostly undefined cell types (Schwarz et al., 2005). A further analysis

91

of calluses derived from various organs is necessary to understand the causes of organ related

differences in organogenesis responses.

6.1.2 MP/ARF5 in auxin and cytokinin mediated developmental processes

The MP gene together with NPH4 plays a key role in embryonic, vascular, and phyllotactic

responses (Hardtke et al., 2004). However, the overexpression of NPH4 can not rescue the

phenotype of mp mutants, which indicates only unilateral redundancy of their developmental

roles (Hardtke et al., 2004; Wilmoth et al., 2005). The results in this study showed that

callusogenesis and root regeneration had relatively similar features in mpG12 and nph4-1

mutants. The quantitative differences that distinguish callusogenesis and rooting responses in

mpG12 from those in nph4-1 possibly reflect the levels of contribution of NPH4 and MP in a

particular process (Fig. 3.1; 5.1). However, the ability for shoot regeneration strongly

distinguished mpG12 from nph4-1 mutants (Fig. 4.1).

In particular, the impairment of callus proliferation in mpG12 aerial explants was less

pronounced than that in nph4-1. However, dissimilar to nph4-1, the fresh weight of the mpG12

adventitious root derived calluses was below that of the wild type ones. This may indicate that

some level of auxin insensitivity exists in all mpG12 organs. The generally milder phenotype of

mpG12 aerial explants on CIM can possibly be explained by the presence of the functional

NPH4 gene. NPH4 contribution to callusogenesis seems stronger than that of the MP gene.

Based on the callusogenesis phenotype of mpG12 mutants, the possible contribution of the MP

gene to the regulation of cell cycle genes may also be a consideration (Richard et al., 2002;

Perrot-Rechenmann, 2010).

Root regeneration in mpG12 and nph4-1 explants had rather similar features. Based on normal

root regeneration in the mpG12 aerial explants, two suggestions can be made. Either MP is not

involved in de novo root formation in aerial explants; or, other ARFs substitute for the lack of

MP function (Hardtke et al., 2004; Wilmoth et al., 2005). This might be surprising, given MP

requirement for embryonic root formation, but it has previously been noted that this does not

imply a similar role in other cell types (Berleth and Jurgens, 1993). Similarly to NPH4, MP can

facilitate root formation in calluses derived from adventitious root explants, as indicated by the

result that fewer roots were formed in the mpG12 adventitious root calluses. The low root

92

formation in mpG12 root explants may reflect the regulation of lateral root formation by the

MP gene, shown recently by De Smet et al. (2010).

Normally, the MP gene is not expressed in mature root tissue (Dr. W. Ckurshumova, personal

communication), although MP expression is reactivated during callus proliferation (Dr. W.

Ckurshumova, personal communication), and during lateral root initiation (De Smet et al.,

2010). This concords with the data presented in this thesis, which indicate that the MP gene

might be involved in callusogenesis and root formation in adventitious root explants based on

the phenotype of mpG12 mutants.

The major difference between mpG12 and nph4-1 mutants was in their ability for shoot

regeneration (Fig. 4.1). Unlike in nph4-1, shoot regeneration was consistently very low in all the

types of mpG12 explants. Hardtke et al. (2004) demonstrated that the functions of NPH4 and MP

during SAM formation in embryos are only partially redundant, and the NPH4 gene could not

rescue the loss of MP functions in mp mutants. Likewise, the low rate of de novo shoot

regeneration in mpG12 may indicate one of the major differences between MP and NPH4

functions in de novo SAM formation.

As previously reported (Przemeck et al., 1996), the overall morphology of the developing

mpG12 shoots was relatively normal during vegetative stages. Similarly, in this study, no major

abnormalities were detected in regenerated shoots on SIM. Taking into consideration that the

competence to form organs is acquired sequentially, first roots and then shoots (Ozawa et al.,

1998), and that de novo root regeneration in mpG12 explants was not impaired, it is possible to

suggest that MP function is somehow related to the conversion of LRM-like structures to shoot

meristems in response to cytokinin (Che et al., 2007; Atta et al., 2009; Sugimoto et al., 2010).

A growing body of research shows that MP activity is linked to regulation of shoot meristem size

and formation (Vidaurre et al., 2007; Schuetz et al., 2008; Zhao et al., 2010), and to the activity

of several key meristem genes (Aida et al., 2002; Cole et al., 2009; Chapter 1.2.5). Moreover, the

MP gene facilitates auxin-cytokinin crosstalk via the auxin mediated regulation of two A-type

ARRs (Zhao et al., 2010). The strong phenotype of mpG12 mutants in shoot regeneration also

demonstrates the key role of the MP/ARF5 gene in de novo SAM initiation.

93

6.1.3 Enhancement of de novo organogenesis in the dMP genotype

MP function is elevated in the dMP genotype due to a constitutive MP activity in the truncated

dMP transgene (Fig. 1.2; Krogan, 2006). Krogan (2006) demonstrated that auxin responses are

strongly elevated in the dMP genotype. His results also showed a high capacity of cotyledon

explants for rooting in liquid RIM (Krogan, 2006). The data of this study further indicated an

enhancement of organogenesis responses in dMP, such as callusogenesis, root regeneration,

and particularly strongly, shoot regeneration. This enhancement, however, was detected only in

the aerial, but not in the adventitious root explants, indicating the organ dependent character of

these responses in the dMP genotype.

The dependence of organogenesis responses on MP activity in the GOF and LOF genotypes

(dMP and mpG12) turned out to be complex. Elevated organogenesis responses in dMP were

not always paired with a reduction of the same responses in LOF mpG12, as was observed

during shoot regeneration in aerial explants (Fig. 4.1). However, that was not always the case

in other organogenesis responses and explant types. Moreover - though to a different degree -

shoot organogenesis was diminished in the adventitious root explants of dMP and mpG12 (Fig.

4.1). Thus, the data of this study indicated that the effect of MP on organogenesis processes

depended on the level of gene activity only in particular processes, such as shoot regeneration

in aerial explants. In other cases, the increase in organogenesis responses by dMP was not

significant, or it may have been prevented by some unknown limiting factors. For example,

low shoot regeneration in dMP adventitious roots was greatly elevated when the time of CIM

preculture was increased, although the latter had no effect on mpG12 explants (Fig. 4.5). This

may indicate that some factors suppressing shoot regeneration in dMP were eliminated by the

prolonged time on CIM. More experiments would need to be done to be able to characterize

the factors and their molecular bases that possibly counteract MP activity in organogenesis

responses.

6.2 General Conclusion

According to the results of this study, callusogenesis, and root and shoot regeneration showed

genotype and organ related differences. Loss of MP and NPH4 functions affected callus and root

regeneration in mpG12 and nph4-1 mutants in a relatively similar manner. However, a major

94

difference in organogenesis responses between these arf mutants was indicated in their shoot

regeneration. Putatively elevated MP activity levels in the dMP genotype seem to have resulted

in the promotion of root regeneration, callus growth, and shoot regeneration in aerial, but not in

adventitious root explants. The different levels of gene redundancy, and the joint functions of

MP and NPH4 (Hardtke et al., 2004) during de novo organogenesis can be suggested based on

the results of this study. Overall, screening arf mutant genotypes in tissue culture assays can be a

powerful tool to elucidate unknown functions of ARF genes, as demonstrated in this thesis. The

possibly dominant effects of other ARF gene transgenes on organogenesis processes in vitro can

be of significant interest for plant biotechnology, if they can be reproduced in suitable plant

species - ones with normally poor regeneration properties.

95

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