Verticordia Micropropagation Through Direct Ex Vitro Rooting

Home , Chamelaucieae, Homoranthus, Verticordia, Verticordia densiflora, Verticordia eriocephala, Verticordia grandis

Edith Cowan University Research Online

Theses: Doctorates and Masters Theses

2013

Verticordia micropropagation through direct ex vitro rooting

Belinda Delaney Edith Cowan University

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Recommended Citation Delaney, B. (2013). Verticordia micropropagation through direct ex vitro rooting. https://ro.ecu.edu.au/ theses/615

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The Use of Thesis statement is not included in this version of the thesis. Verticordia micropropagation through direct ex vitro

rooting

Belinda Delaney

BSc (Biological Sciences) School of Natural Sciences Edith Cowan University Abstract

The objective of this study was to improve the existing shoot multiplication protocol for

Verticordia grandis (McComb, Arthur & Newll, 1986; Newell, Growns & McComb, 2005) and to investigate and establish reliable root induction and acclimatisation protocols to enhance survival of micropropagated plantlets. It was envisaged that these protocols would be successful in micropropagation, growth and survival of different V. grandis clones and possibly applicable to other Verticordia species.

The elongation of in vitro Verticordia shoots on multiplication media was improved by reducing the concentration of BAP from 1µM to 0.25 µM, which resulted in a more uniform shoot length of 4.5 – 5 cm; necessary for root induction experiments. The root induction protocol was optimised by determining the appropriate auxin concentration (80µM indole butyric acid; IBA) with an exposure time of 6 days. Acclimatisation and survival was greatly improved by transferring the IBA pulsed shoots to ex vitro conditions consisting of a free draining and aerated substrate (a mixture of peat and perlite 1:3) in crack pots. These were initially placed into a greenhouse (with controlled temperature & light conditions) in order to maintain high humidity.

Over time humidity was reduced and after 112 days the plantlets were transferred to larger pots, containing fresh soil (peat/perlite/sand = 1:1:1) and placed in a shade house with a regular watering regime. Long-term survival was monitored and after 252 days survival was over 70%.

The declining survival rates after this time has made it evident that field performance and long- term survival needs further investigation. The application of the improved shoot multiplication and root induction protocols on other V. grandis clones produced survival rates of 0 to 62.5%

(depending upon clone) over 252 days.

I certify that this thesis does not, to the best of my knowledge and belief:

(i) incorporate without acknowledgment any material previously submitted for a degree

or diploma in any institution of higher education;

(ii) contain any material previously published or written by another person except where due reference is made in the text of the thesis; or

(iii) contain any defamatory material

______

Acknowledgements

I dedicate this research to the memory of Edward ‘Ted’ Williams and

Diane Forsythe

I hereby gratefully acknowledge the people who supported my research for this masters project,

especially my supervisors Dr Ian Bennett and Dr Mary Boyce. I thank them for their support,

advice, time, encouragement and influence, as these were greatly appreciated. Thanks are also due to Clay Millar and his laboratory staff who devoted time and energy into all practical aspects of my working in a laboratory, whilst I was researching for my project. I consider myself

fortunate to have had the financial support for my research provided by the School of Natural

Sciences at ECU. I also wish to thank my fellow students, volunteers and friends that I have met over the years at ECU, especially those who offered encouragement and were always ready to lend a helping hand.

This project would have not been possible without the dedication of Ted Williams who helped

me discover Eneabba’s wildflower treasures, Alan Tinker who shared his vast knowledge of

Beekeeper’s Reserve and his amazing botanical skills with me, and Diane Forsythe, who always

encouraged my endeavours in any way possible.

I hope the research presented here may contribute to the conservation of the amazing scarlet

feather flower – Verticordia grandis.

TABLE OF CONTENTS

Use of Thesis Declaration 1

Abstract 2

Copyrights and Access Declaration 3

Acknowledgements 4

1. Introduction

1.1 Background 9

1.2. Myrtaceae and Genus Verticordia 13

1.2.1 Verticordia Habitat 13

1.2.2 Verticordia Morphology 14

1.2.3 Verticordia Seed Germination and Reproduction 14

1.2.4 Verticordia Conservation & Cultivation Value 16

1.2.5 Verticordia Propagation 16

1.2.6 Micropropagation of Verticordia grandis 17

1.3 Micropropagation 18

1.3.1 Principles of in vitro Propagation 18

1.3.2 Composition of in vitro Culture Media 20

1.3.3 Physical environment for micropropagation 21

1.3.4 Selection of Source Plants for Micropropagtion 23

1.3.5 Juvenility & Rejuvenation in Woody Plants in Tissue Culture 24

1.3.6 Morphological Characteristics of Micropropagated Plantlets 25

1.3.7 Root Induction 27

1.3.8 Acclimatisation of in vitro Microcuttings to ex vitro Conditions 28

1.3.9 Acclimatisation 30

1.4 Aims 31

1.4.1 Limitations of conventional propagation methods 31

1.4.2 Research aims 32

1.4.3 Benefits of this research 33

2. Materials & Methods

2.1 Plant material 34

2.2 Process of initiation of new clones into culture 34

2.3 Media preparation 35

2.4 Culture conditions 36

2.5 Changes to shoot multiplication media composition 37

2.5.1 Testing various BAP concentrations to improve elongation in clones 37

2.6 Original root induction protocol 38

2.6.1 Optimisation of duration of IBA treatment for root induction 39

2.6.2 Optimisation of IBA concentration for root induction 39

2.6.3 Trials with auxin combinations for root induction 40

2.7 Trials with different in vitro substrates for root induction 41

2.7.1 Comparison of IVS with different nutrients 42

2.7.2 Comparison of in vitro to ex vitro on IVS media 43

2.8 Survival in greenhouse during acclimatisation phase 43

2.9 In vitro shoot multiplication of new GRD clones 45

2.9.1 Root induction of successful new clones 45

2.10 Statistical analysis 46

3. Results

3.1 Survival of new clones initiated into culture 47

3.2 Changes to shoot multiplication protocol 48

3.3 Changes to root induction media composition 50

3.3.1 Optimisation of duration of IBA pulsing 50

3.3.2 Optimisation of IBA concentration 52

3.3.3 Auxin combinations for in vitro root induction 54

3.4 Different substrates 56

3.4.1 Comparison of Agar, black sand and IVS 58

3.5 Comparison of IVS with different nutrients 60

3.6 Ex vitro root formation and acclimatization 63

3.7 Survival 65

3.8 Effectiveness of improved protocols on new clones 67

3.8.1 Shoot growth and maintenance 67

3.8.2 Roots induction, growth and acclimatisation 68

4. Discussion

4.1 Collection and survival of new clones initiated in to culture 69

4.2 Changes to shoot multiplication protocol 70

4.3 Changes to root induction media composition 71

4.3.1 Optimisation of duration of IBA pulsing 72

4.3.2 IBA concentration 73

4.3.3 Auxin combinations for in vitro root induction 73

4.4 Different substrates 75

4.4.1 Comparison of Agar, black sand and IVS 77

4.5 Comparison of IVS with different nutrients 78

4.6 Ex vitro root formation and acclimatization 80

4.7 Survival 81

4.8 Effectiveness of improved protocols on new clones 83

4.8.1 Survival of new clones 84

5. Conclusion 85

6. References 87

TABLES

Table 2.1: Origin of the explant material for new clones 35 Table 3.1: Survival rates of new microcuttings 47 Table 3.2: Periodical assessment of in vitro root induced GRD1 shoots 67

FIGURES

Figure 1.1 Map of the Southwest Botanical Region in Western Australia. 10 Figure 3.2 Means for growth of shoot length 49 Figure 3.3 Optimisation of IBA pulsing time 51 Figure 3.4 Optimisation of IBA concentrations for root induction 53 Figure 3.5 The effect of different auxin combinations and concentrations 55 Figure 3.6 Comparison of different substrates and root growth 57 Figure 3.7 Comparison of different substrates and root growth 59 Figure 3.8 Comparison of root appearance from different substrates 60 Figure 3.9 Comparison of different nutrients and root growth 62 Figure 3.10 Comparison of sterile soil vs direct soil 64 Figure 3.11 Comparison of in vitro & ex vitro root growth and survival 66 Figure 3.12 Survival of all clones 69

ABBREVIATIONS

BAP – 6 benzylaminopurine DDI – double de-ionised water CO2 – carbon dioxide

GF – Verticordia grandis from lignotuber regrowth after fire GRD – Verticordia grandis clone IAA – indole-3-acetic-acid IBA – indole-3-butyric-acid IVS – in vitro soil MilliQ water – de-ionised water MS – Murashige & Skoog basal salt mixture NAA – naphthalene acetic acid NaOH _ sodium hydroxide

O2 - oxygen VOV – Verticordia ovalifolia clone VFR – Verticordia fragrans clone VDF - Verticordia densiflora clone

1. Introduction

1.1 Background

The Southwest Botanical Province of Western Australia is situated between latitudes 27° S and

35° S and longitudes 112° E and 128° E‘ and extends from Shark Bay in the northwest to

Esperance in the southeast. This region includes the narrow coastal land from east of Esperance stretching to the Western and South Australian border and is confined by oceans in the west and south and by arid landscapes in the north and east (Cole, 2006).

Figure 1.1: Map of the Southwest Botanical Region in Western Australia. The green area shows the ‘eco-region’ boundary; the dark yellow area shows the ‘transitional zone’ from Shark Bay in the northwest continuing to the SA border in the southeast (Cole, 2006).

It is recognized as one of the world’s 34 major biodiversity hotspots and is the only one in

Australia; it consists of more than 480 000 km2 (48.0 million hectares) with over 52% of vascular plants and 12.5% of genera being endemic to the region. This region is also classified as a ‘centre for plant diversity’ by the WWF and the International Union for Conservation of

Nature and Natural Resources (IUCNN) because of its plant species richness (Hopper & Gioia,

2004). The southwest of Australia is one of the oldest landscapes in the world with very diverse vegetation communities that have adapted to the leached and nutrient-poor soils. The species richness and diversity of flora in this region is probably due to the combination of ecological and phylogenetic processes that have occurred over long periods of time. This could explain the diverse plant communities; ranging from kwongan (sclerophyllous shrubs) heath- and shrub lands on the coastal sand plains to temperate forests in the far southwest, and wood- and shrub lands further east. All these plant communities vary greatly over short distances and because of these varying features the region has become popular for its wild flower displays in spring, and eco-tourism has become an important industry (Cole, 2006). Many of these plants have become desirable for the domestic and export cut flower trade, ornamental horticulture and land restoration (Seaton, 2002).

The world’s plant biodiversity is in sharp decline with many species being listed as ‘critically endangered, endangered or vulnerable’ and requiring conservation (Sarasan, Cripps, Ramsay,

Atherton, McMichen, Prendergast & Rowntree, 2006); this includes many taxa of the Southwest

Botanical Province (Cole, 2006). Over the last century this region has been extensively degraded, resulting in landscape modification, fragmentation, biodiversity and habitat loss (Hobbs &

Saunders, 1991). Human induced processes have altered over 90% of the original vegetation

(Beard, 1980), which has been reduced to mostly scattered, fragmented vegetation remnants in

various conditions. Land clearing for agricultural purposes, urban development, domestic stock

grazing, logging and changes in fire regimes account for most of the native vegetation loss and

habitat destruction (Cole, 2006). Salinisation of groundwater, due to altered landscape hydrology

(Hobbs, 1992), susceptibility of remnant vegetation to weed invasion (Pigott, 2000) and (feral)

grazing animals are other threats to the remaining bio-diversity of the Southwest Botanical

Province (Denham & Auld, 2004). Another major threat to the flora of this region is a disease caused by Phytophthora cinnamomi, a soil-borne pathogen (water mould), which can cause irreversible damage to affected plant communities. The pathogen attacks the roots of susceptible host plants, thus significantly reducing the host plant’s ability for nutrient and water transport, and eventually kills the host (Shearer, Crane, Barrett & Cochrane, 2007).

The conservation significance of the highly diverse and endemic flora of the Southwest

Botanical Region has a high priority with government conservation agencies, as many critically endangered taxa have been identified. Many of these endangered species require immediate remedial action to prevent extinction (Coates & Atkins, 2000), and in situ conservation by protecting natural habitats and efficient management of wild plant populations is very efficient with many species. But these sometimes need to be complimented with ex situ methods

(Sarasan, et al., 2006), as numerous species have recalcitrant seeds or produce poor quality seeds; so successful seed propagation and plant establishment is often very difficult or impossible (Bell, 1999). Many taxa from the Myrtaceae, including some Verticordia, are

threatened by habitat loss.

1.2. Myrtaceae and Genus Verticordia

Myrtaceae is one of the ten largest plant families in the Southwest Botanical Province and of the

785 species 92% are endemic (Beard, Chapman & Gioia, 2000). The genus Verticordia, belongs

to this family and is a member of the Australian sub-family Leptospermoideae, characterised by

their dry capsular fruit (Cochrane, Brown, Cunneen & Kelly, 2001; Clark & Lee, 2002). It was

named by Augustin Pyramus de Candolle in 1821, and is endemic to Australia (George, 1991).

Verticordia belong to the Chamelaucium alliance with its closest relatives being Chamelaucium,

Darwinia, Homoranthus and Rylstonea (George, 1991). These are commonly associated with

many other myrtaceaous genera in Western Australia’s shrub and heath land communities

(Cochrane, et al., 2001).

1.2.1 Verticordia Habitat

All but three of the species, subspecies and varieties of verticordias are endemic to Western

Australia, and the distribution largely depends on the annual rainfall, with many adapted to 350

mm to 850 mm rainfall zones (George, 1991; George, 2002; Seaton, 2002). They grow in a

diversity of habitats and soils ranging from coastal regions to far inland arid regions. They occur

as woody perennial understorey shrubs in open forests or as heath shrubs on sand plains and even

as saline tolerant shrubs growing on the edges of salt lakes (George, 1991; Seaton, 2002).

Although verticordias grow in this diversity of habitats and soils, some only occur in very

particular locations and/or specific conditions, many have already been declared rare and endangered (George, 1991; George, 2002; Yates & Ladd, 2004).

1.2.2 Verticordia Morphology

Verticordias generally grow up to 2 m tall, with a few species growing taller, and have very diverse forms with considerable variation depending on species, although the bushy shrub is the most common form. Their shrub form may be dwarf, prostrate, erect or widely spreading with a compact, straggly or horizontally layered appearance (George, 1991). All verticordias, however, share one prominent characterising feature; the sepals are deeply divided in various ways, with most species having petals that are often dentated, divided, fimbriated or lobed and most flowers having a hairy hypanthium (Marchant, Wheeler, Rye, Bennett, Lander, MacFarlane, 1987), hence, the common name - feather flowers.

1.2.3 Verticordia Seed Germination and Reproduction

Seed production and germination processes are critical for the survival of most plant species

(Bell, 1999), but poorly understood for Verticordia species. Bell (1999) emphasised that

Australian plant species rely on several germination mechanisms that interact to interrupt seed dormancy and initiate germination when environmental conditions are suitable. Cochrane et al.

(2001) reported that for many Verticordia species there is no adequate knowledge of the reproductive potential, although some germination trials with a few species have been successful

(Cochrane, et al., 2001). Generally, annual reproductive capacity in the form of seed production and viability in many Verticordia species is exceedingly low (Cochrane, et al., 2001).

Many Verticordia species are capable of superfluous flower production and the low quantity of seeds produced is disproportionate to the number of flowers. This low seed to flower ratio is common in the genus (Housten, Lamont, Radford & Errington, 1993; Cochrane, et al., 2001) and

Cochrane et al. (2001) concluded that the low seed set could be a result of the self- incompatibility common in Myrtaceae. However, as Tyagi, McComb & Considine, (1991) and others (Bellairs & Bell, 1990; Speer, 1993; McEvoy & True, 1995; Bell, 1999) suggest an array of environmental and physiological factors could contribute to the low seed set and viability in the genus (Rye & James, 1992; George, 2002; Yates & Ladd, 2004). It is not unusual for

Australian native species to have limited seed production as many of these plants reproduce vegetatively (Johnson & Burchett, 1996; Kimpton, James & Drinnan, 2002). Cochrane, et al

(2001) also proposed that another biological constraint that could play a part in the reproductive success is that protein and lipid creation required for seed production requires more energy than flower production. This may be a survival strategy for taxa during adverse environmental conditions, e.g. changes in pollinator activity, predation, competition and other unpredictable events (Cochrane, et al., 2001).

Tyagi, Considine & McComb (1992) researched the remarkable longevity of some verticordia pollen at different temperatures for species both with and without a pollen presenter, but did not discover the physiological basis for this. It was suggested that this long viability could facilitate artificial hybridisation of species with breeding barriers such as different flowering times, but it was not determined if in vitro germination levels could match levels of in vivo germination of the

Verticordia species examined (Tyagi, et al., 1992).

While verticordias have limited reproductive success from seed germination, Bell (1999) states, that the secondary dormancy mechanisms, which can prevent germination of many Australian native species are not fully understood. Several Western Australian recalcitrant plant seeds have

been investigated on with seed dormancy-relieving treatments, including smoke treatments, to promote and increase germination (Dixon, Roche & Pate, 1995; Roche, Dixon & Pate, 1997;

Roche, Koch & Dixon, 1997), and it was suggested that research on seed germination and recalcitrance needs to be further investigated. Bell (1999) concluded that it was vital to understand the eco-physiology of seed germination processes and that further research would be needed in order to improve conservation of Australian plant species for improved land management, rehabilitation and for progress in horticultural practices.

1.2.4 Verticordia Conservation & Cultivation Value

The long lasting feather flowers are highly sought after for the commercial flower markets and many species have been bush-picked to the extent that this had adversely impacted on wild populations (e.g. Verticordia eriocephala). Commercial bush-picking, therefore, has become unsustainable for most native populations (Tyagi et al., 1991; Atkins, 1998; Cochrane, et al.,

2001) and the harvesting from natural bush populations is now restricted (George, 2002).

Verticordias are not widely cultivated for the cut flower trade (Seaton, 2002), but some success

has provided an alternative to bush-picked plants. However, this has been limited to only a few

species, such as V. plumosa (Seaton, 2002).

1.2.5 Verticordia Propagation

In the past, asexual propagation techniques have been successfully used on some Verticordia

species (e.g. V. plumosa, V. chrysanthella), with the focus mainly on cuttings and some success

16 with tissue culture (George, 2002). In addition, Egerton-Warburton, Ghisalberti, & Burton,

(1998) found that intergeneric hybridisation facilitated by synchronous flowering can be useful

(e.g. C. floriferum x V. plumosa), and grafting on to Chamelaucium root stock (Yan, 2001) has also produced some useful results. Grafting of several Verticordia species, for commercial applications in field conditions, however, has not been successful (Ben-Jaacov, Ackerman &

Evenor, 2000). Grafting is also considered uneconomical due to its slower production rate and additional costs (Lullfitz, 2001).

1.2.6 Micropropagation of Verticordia grandis

The application of ex situ conservation practices for plant species can compliment in situ methods, or replace them all together with species that are too difficult to propagate by conventional methods (Sarasan, et al., 2006). Verticordia grandis, with its large scarlet red flowers, is an attractive species for the cut flower industry, but has proven to be difficult to propagate by conventional methods (Speer, 1993), due to recalcitrance and low viability of its seeds (Bell, 1999). Pollination of this species is poorly understood (Bell, 1999), but it can regenerate from a lignotuber; it is one of the few verticordias that possess this capacity (George

& George, 2003). A reliable alternative to conventional propagation methods is tissue culture, which can successfully produce viable clones of difficult to propagate species (Ilan & Khayat,

1997). Micropropagation of V. grandis could produce numerous, disease free plants for horticultural purposes, the cut flower trade and for the restoration of degraded habitats.

1.3 Micropropagation

1.3.1 Principles of in vitro Propagation

The development of the elementary methods of plant tissue culture took place in the 1950s and

1960s and has since advanced to commercial plant biotechnology, which has become an invaluable tool in the plant propagation industry; e.g. ornamental horticulture and agriculture

(DeKlerk, 2002). One distinct advantage of producing plants by tissue culture is the raising of disease free true-to-type plantlets and minimising diseases and pests, which include internal pathogens, phytoplasma, viruses, viroids, parthogenic fungi, moulds, yeasts, bacteria and insects.

(Bandyopadhyay, Cane, Rasmussen & Hamill, 1999; De Klerk, 2002). In general it is based on the principle of establishing and maintaining healthy micro-shoots in culture for multiplication so that these produce rooted micro-cuttings and plantlets (DeKlerk, 2002; Liu & Bao, 2003).

Conventional propagation methods are often inefficient for multiplying many mature and/or difficult-to-propagate plant species, but tissue culture can successfully produce viable clones

(Ilan & Khayat, 1997). However, it has also been well documented that root induction can be problematic in woody plants (DeKlerk, 2002; Liu & Bao, 2003; Seth, Kendurkar, & Nadgauda,

2007). A major benefit of micropropagation applied to slow growing woody perennials is the propagation and multiplication of uniform and numerous disease free plantlets in a short time, as there are no effectual vegetative propagation techniques for reproducing mature plants that can equal this. Microproagation also allows for faster and greater production of numerous plantlets compared to the slower, conventional seed or vegetative propagation methods (Franclet, 1991;

Haapala, Pakkanen, Pulkinnen, 2004).

The basic principle of plant micropropagation of meristematic shoot tissues is to grow, manipulate and multiply identical plant cells, tissues and organs, which have been isolated from the mother plant. This approach involves techniques that are conducted in a controlled and aseptic environment; the cells, tissues and organs of a selected plant are isolated, surface sterilised (Teng, Sin & Teng, 2002) and cultured in a growth-promoting environment (Franclet,

1991; van Acker & Scholten, 1995; George, Hall & de Klerk, 2007). There are generally four distinct phases applied in micropropagation; 1. Establishment, 2. Multiplication, 3. Root

Induction and 4. Acclimatisation. In the ‘Establishment Phase’ a plant part is surface sterilised and then introduced to a sterile culture media. This phase determines the suitability of the explants; e.g. being free of exo- and endogenous contamination. In the ‘Multiplication Phase’ the explants are aspectically transferred to media containing growth hormones that promote shoot proliferation (usually) and elongation. This phase can generate numerous, stabilised explants in a very short time, which can be kept in culture (Multiplication Phase) as long as needed by periodic sub-culturing on to new ‘multiplication media, or the explants can be aseptically transferred to root induction media (Root Induction Phase). In the ‘Root Induction Phase’ elongated explants are aseptically transferred to a growth media that usually contains auxin/s to initiate and promote root growth in preparation for autotrophic growth of explants. In the

‘Acclimatisation Phase’ rooted explants are transferred to a suitable growth substrate and placed in the controlled environment of a glasshouse for gradual acclimatisation to ex vitro conditions.

The plants are initially kept in high humidity, which is gradually decreased as plantlets acclimatise; they become more capable of controlling water loss.

1.3.2 Composition of in vitro Culture Media

An optimised culture media is essential for shoot growth, multiplication and root induction in tissue culture, as the media has to provide all nutrients and elements for in vitro growth of plants.

Most culture media contain the following:

• Minerals (as macro- and micro salts): Most plants respond favourably to the widely

applied Murashige & Skoog (1962) MS salts. Lloyd & McCowan’s (1980) Woody Plant

Medium is also widely used. The application quantity depends on the in vitro cultivation

objectives; e.g. high formulations (full strength) are better suited for shoot multiplication,

1 1 whilst lower concentrations ( /2 or /4 strength) are generally used to promote root

induction depending on plant species (Kim, Oh, Jee & Chung, 2003).

• Carbon (energy) source: Sucrose in the form of commercial sugar is the most preferred

carbon source in tissue culture because it is cheap and readily available. Carbon (e.g.

sucrose, glucose, maltose or galactose) for in vitro plant metabolism has to be added to

many culture media as the plantlets are not fully autotrophic and photosynthesis is not

adequate (Rahman, Islam, Hossain & Islam, 2010).

• Plant growth regulators (auxins, cytokinins, gibberellins and others): Several growth

regulators consist of natural and/or synthetic compounds that manipulate development

and growth of in vitro plants. The application and quantities of growth regulators

depends on the in vitro cultivation objectives; e.g. callus formation, shoot multiplication

and elongation or root induction. Auxins regulate several physiological process; e.g.

formation of shoots and adventitious and lateral roots (Ibrahim & DeBergh, 2001).

• Other organics: Vitamins are not necessarily added to tissue culture media, but most

formulations contain vitamins; e.g. in MS salts the included vitamins are thiamine,

pyridoxine, nicotinic acid, with myo-inositol (sugar alcohol) and thiamine considered to

be essential to plant growth (Abrahamian & Kantharajah, 2011).

• pH; the pH is adjusted to 5.8 using 1M NaOH for shoot multiplication to approximate

what would be suitable for plant metabolism and to ensure agar solidification.

• Gelling agents (purified agar or gellan gum products): In micropropagation traditionally

solid media is favoured over liquid media with agar often being the preferred gelling

agent. The concentration of the gelling agent is associated with the water potential from

the medium to the cultured plants, depending on the objectives of cultivation; e.g. shoot

multiplication or root induction (DeBergh, 1983). Gellan gum (Gelrite) produces a clear

gel, which allows for accurate observation of root induction, and it has been observed that

many Australian woody plants have a greater survival rate when cultured in a Gelrite

medium (Williams & Taji, 1987).

• De-ionised water; water is the universal solvent for all solutes required for

micropropagation culture media (DeBergh, 1983).

• The media needs to then be dispensed into appropriate culture vessels and sterilised in an

autoclave generally at 121oC and 1.2kg/cm-2 for 20 minutes.

1.3.3 Physical environment for micropropagation

The physical environment for micropropagation is as important as the appropriate culture media formulation. Light and temperature cycles, necessary for optimum growth of cultured plantlets, are programmable and controllable according to the individual plant species’ requirements

(Johnson, 1996).

• Light: Plant development and growth is influenced by wavelength, intensity and the

duration of light and this can be controlled in micropropagation according to plant

species and the in vitro culture objectives. Light is also an important factor for

phototropism, morphogenesis and photosyntehsis (Read & Preece, 2003). It has been

observed that light can enhance shoot growth and root formation in some species, whilst

other species preferred darkness for root induction (Kumar, Palni & Nandi, 2003).

• Temperature: Plant development in micropropagation also depends on the optimum

temperature range for the physiological process of respiration, but these optimum

temperatures vary according to species and genotypes. Culture temperatures between 20°

and 27° C are most commonly applied (Read & Preece, 2003).

• Gas exchange: In vitro plantlets are grown in culture vessels that present a closed system

to prevent microbial contamination. This can limit the inflow of CO2 and the outflow of

O2 and ethylene, although some gas exchange may occur depending on the type of

culture vessels used (Park, Jeon, Kim, Park, Aswath & Joung, 2004). The relative

humidity inside the culture vessel is usually very high, approximately 98-100%

(Gribaudo, Restagno & Novello, 2003), which can be problematic for initial

acclimatisation of plantlets from in vitro to ex vitro conditions.

1.3.4 Selection of Source Plants for Micropropagtion

Cuttings from different individual explants are likely to behave differently under culture

conditions (Geneve, 1989; Anthony, McLean & Lawrie, 2000). Nevertheless, it is important to

carefully select plant source material that is of vigorous, healthy appearance with the desired

phenotypic attributes (Kartsonas & Papafotiou, 2007). The initial explant source selection is vital

with regard to epigenetic and genetic characteristics of the source plants (Wiltshire, Potts &

Reid, 1998), so the careful selection of source plants is crucial (Idczak, & Brielmaier-Liebetanz,

2003). This can ensure the cloning of the desired ‘true-to-type’ phenotypic plant material, although this can only be confirmed by close molecular examination (Arora, Sharma, Srivastava,

Ranade & Sharma, 2011).

Epigenetic and genetic stability of micropropagated plants are an important pre-requisite and this has been investigated with various molecular techniques. Hop plants displayed epigenetic differences between the field grown mother plants and the micropropagated clones (Peredo,

Arroyo-Garcia & Revilla, 2009). Kaeppler, Kaeppler & Rhee (2000) observed that there was phenotypic variation and temporary alterations in the physiology of the shoots in micropropagated plants compared to seed grown plants. But it was also concluded that over time, with repeated subculture, the phenotypic variation would initially accumulate but then reduce; the micropropagated material would become more similar to the source plant with culture age (Rodriguez Lopez, Wetten & Wilkinson, 2010). The long-term genetic stability was also confirmed in a study for banana clones (Lakshmanan, Venkataramareddy & Neelwarne,

2007).

1.3.5 Juvenility & Rejuvenation in Woody Plants in Tissue Culture

Several studies of the different phases in plant development have verified that there are four phases; embryonic phase (establishment of shoot and root meristems), juvenile phase (no reproduction capabilities), adult vegetative phase (establishment of reproductive capacity) and the adult reproductive phase (Hackett & Murray, 1992; Greenwood, 1995; Wiltshire, et al.,

1998). These phases are stable and comparatively inconspicuous as the characteristics of one phase are usually replaced by traits that characterise the next phase (Conway & Poethig, 1993;

Kerstetter & Poethig, 1998).

As with other methods of propagation, the micropropagation success of woody plant species can depend largely on the source plant’s juvenility status (Hackett, 1985), as the more juvenile explant is easier to reproduce than a more mature plant (Lullfitz, 2001). A cutting from a woody plant in the juvenile phase has greater ability to root and this is regarded as evidence that the plant has not yet reached maturity (Hackett, 1985; Jones, 1999). The juvenile stage is defined by gradual changes, e.g. biochemical, morphological and physiological characteristics, that occur in the young plant after germination (Haapala, et al., 2004).

One of the greatest limitations in slow growing woody perennials is their recalcitrance due to complex seasonal and life cycles, which can make it difficult to establish stabilised shoot cultures (McCown, 2000). This could be overcome with the generation of shoot culture by micropropagation techniques that utilise the plant’s meristematic tissues (Irish & Karlen, 1998) that can renew and sustain growth, and produce new plants from the existing vegetative structures (Schmidt, 1997; Franclet, 1991; Hackett & Murray, 1992). Each step of

micropropagation can be manipulated and has become the preferred method for production of

woody plants that typically have long juvenility phases and slow breeding processes (Stokes,

1980; de Jeu & Cadic, 2000), as observed in many Verticordia species. Tissue culture can be a

method to restore the juvenile phase in large amounts of explant material, as studies suggest that

sub-culturing over time can re-establish juvenility in clones cultured from mature woody plants

(McCown, 2000; Andreu & Marín 2005).

Hackett (1985) stipulates that juvenility of a woody species ends with the onset of flowering.

Andreu & Marín (2005) discussed reinvigoration of mature woody plants during in vitro culture and established that the best multiplication results for shoot increase were gained from micropropagated plants that originated from cutting derived plants. These rejuvenated plants differ in physiology and often have a restored rooting capability that may only be temporary

(Hackett & Murray, 1992; Haapala, et al., 2004).

1.3.6 Morphological Characteristics of Micropropagated Plantlets

Most research suggests that genotypic and phenotypic variation seems to be a common occurrence observed in micropropagated plants. These variations can pre-exist in plant material from natural populations that has been used for tissue culture initiation or they can result from tissue culture conditions. They can be beneficial, as somaclonal variation can lead to the

development of new varieties and/or the development and utilisation of desired traits (Skirvin,

1978; Larkin & Scowcroft, 1981; Perez, Mbogholi, Sagarra, Aragon, Gonzalez, Isidron &

Lorenzo, 2011). But repeated sub-culturing of micropropagation can also result in observed

malformation of leaves, stems and roots; e.g. changes in leaf morphology resulting in thinner

leaves, undifferentiated mesophyll, flattened chloroplasts and other changes at sub-cellular level

which could affect some vital physiological processes like photosynthesis (Skirvin, 1978;

Sebastiani, Minnocci, Vitagliano, Gribaudo & Novello, 2001; Romano & Loucao, 2003). Most

micropropagated plantlets lack a well-developed and functional waxy cuticle, which acts as a

barrier to prevent water loss, although the cuticle is present. This is problematic for ex vitro

acclimatisation as in vitro plantlets cannot regulate their water balance and experience water

stress (Skirvin, 1978; Fila, Ghashghaie, Hoarau & Cornic, 1998). Morphological changes

observed in stems can include shorter internodes, thickened stems and stunted growth (Skirvin,

1978; Martin, 2003). Callus formation, deformed and brittle roots have been observed with in

vitro root induction of shoots which can prove to be problematic when transferring rooted shoots

from in vitro to ex vitro conditions, causing new roots to break (Kim, Klopfenstein & Cregg,

1998; Metivier, Yeung, Patel & Thorpe, 2007).

After successful acclimatisation most of these described morphological deficiencies and

malformations improve with plant age and the plant’s functions (leaf structure, water relations,

photosynthetic parameters) become comparable to the ‘stock’ plant in the field (Pospislova,

Ticha, Kadlecek, Haisel & Plzakova, 1999; Agnihotri, Mishra & Nandi, 2009). DNA analysis

can be used to confirm that in vitro cultured plants are genetically comparable and true-to-type of

the wild host plant (Arora, et al., 2011).

1.3.7 Root Induction

In micropropagation of woody plants, adventitious root induction and formation are fundamental

for successful production of viable plantlets. Many woody plants are recalcitrant and do not

propagate easily due to the lack of morphological plasticity in their cells and tissues and require

the application of exogenous auxins to induce adventitious root induction.

Auxins - The most commonly used auxins for root induction includes indole-3-butyric-acid

(IBA), indole-3-acetic-acid (IAA) and α-naphthalene-acetic-acid (NAA), with IBA being the most effective and most commonly used. The different effects depend on the plant species, the concentration and the purpose of application (Ibrahim & Debergh, 2001; De Klerk, Van Krieken

& De Jong, 1999). Different auxins, concentrations or combinations, may have to be applied depending on rooting treatments (in vitro or ex vitro) and on plant species (DeKlerk, terBrugge

& Marinova, 1997). The application methods vary greatly, depending on the culture methods; for in vitro root induction the auxin is added to the culture medium, for ex vitro root induction the stems can be dipped in a highly concentrated auxin formula (Ibrahim & Debergh, 2001).

Substrate - For in vitro root induction the microcuttings are maintained in a culture media with low mineral concentrations (½ MS or less) and low auxin concentrations to stimulate adventitious root formation (Kim, et al., 2003). For ex vitro root induction, microcuttings can be pulsed on high auxin concentrations in a culture media with low mineral concentrations (½ MS or less) for usually 7 days (Newell, Growns & McComb, 2005) or the cuttings are dipped in high concentrations of auxins before being transferred to an organic and aerobic growing substrate

(peat/perlite/sand) for root formation and simultaneous acclimatisation (Martin, 2003; Martin,

2003/04; Hazarika, 2003; Meiners, Schwab & Szankowski 2007).

1.3.8 Acclimatisation of in vitro Microcuttings to ex vitro Conditions

The transfer of microcuttings from in vitro to ex vitro greenhouse conditions has been problematic for a number of woody species including those related to Verticordia, e.g.

Eucalyptus, Chamelaucium and Scholtzia (Beardsell, 1996; Johnston, 1996; Ben-Jaacov, et al.,

2000; Lullfitz, 2001). Examination of the plantlet condition has resulted in the recognition that plants coming from tissue culture do not necessarily have normal physiological and morphological anatomy (Santamaria & Kertsins, 1994; Dami & Hughs, 1997). These functional differences include lack of junction between adventitious roots and stem vascular tissue, stomatal malfunction and the microcutting’s lack of a well-developed and functional cuticle that acts as a barrier to prevent water loss; this can cause plantlets to die of water stress (Skirvin, 1978; Fila, et al., 1998; Romano & Loucao, 2003). The survival rate of microcuttings to be acclimatised ex vitro can be increased significantly by maintaining high relative humidity (~ 90% - 98%) by using systems that apply water vapour, fog or mist periodically. In particular, the need for high humidity but low water droplet size has been emphasised by several authors (De Bergh, 1991;

Drew, Smith, Moisander & James, 1991; Read & Preece, 2003; Johnson & Armstrong, 2003).

This has been facilitated by the availability of fog generators that can deliver water droplets of less than 5µ, rather than misting systems.

The use of humidity tents can increase the retention of relative humidity and create a microclimate. This watering regime can be reduced as the microcuttings acclimatise (Offord &

Campbell, 1992; Aragon, Escalona, Capote, Pena, Cejas, Rodrigues, Canal, Sandoval, Roels,

DeBergh, & Gonzalez-Olmedo, 2005).

IBA pulsed in vitro shoots of woody plants have successfully been tested for their ability to root

under high humidity (Yan & Sedgley, 2006). New areas of micropropagation were being

investigated and in 2003 Newell and co-workers reported work on shoot microcuttings of a

Verticordia species in a porous agar medium that contained air filled porosity from 10% to 29%.

The researchers concluded that root induction and length was significantly increased compared

to standard agar solidified media (Newell, 2003). Newell et al. (2005) have proposed that an

aerobic medium base may be important in facilitating transfer of in vitro plants to ex vitro

conditions. It was concluded that for several woody perennials the rooting performance in In

Vitro Soil (IVS) was successful and this method could reduce the stress and plant loss during the acclimatisation stage of micro-cuttings from sterile laboratory conditions to field conditions

(Newell, et al., 2005).

Alternatives to the in vitro shoot multiplication (Phase 2) followed by in vitro root induction

(Phase 3) have also been examined. The simultaneous ex vitro root induction and acclimatisation

have been trialled as a cost and time reducing strategy (McClelland, Smith & Carthers, 1990;

Kim, et al., 1998; Martin, 2003; Thomas & Schiefelbein, 2005; Feyissa, Welander & Negash,

2007; Xu, Wang & Zhang, 2008). Under these circumstances root induction and acclimatisation

(Hazarika, 2003) occur simultaneously in a controlled environment such as a greenhouse, under

conditions similar to those used for acclimatisation of in vitro rooted plants. A major advantage

of ex vitro root induction is that root damage from de-flasking and transfer to soil, that can

preclude growth and vigour, is avoided (Yan & Sedgley, 2006).

1.3.9 Acclimatisation

The transfer of plants from culture to soil has, in the past, caused considerable problems and has

been the subject of discussion (Rohr, Iliev, Scaltsoyiannes & Tsoulpha, 2003) and even

conferences on the topic (e.g. International Society of Horticultural Science has organised three

international conferences on this topic in 2003, 2005 and again in 2007). In general, the

requirements are dependent upon two major aspects: plantlet condition and the environmental

conditions required for transfer. More recently, where these aspects have not been important or

overcome (Newell, et al., 2005), other aspects have been concentrated upon; such as the media

from which the plants have been derived and/or the number of roots produced per plantlet

(McCelland, et al., 1990; Bonal & Monteuuis, 1997; Kim, et al., 1998; Bennett, McDavid,

McComb, 2004).

The environmental conditions have received considerable attention and in recent years these

have become more standardised for many species. The ultimate requirement of propagated plants

is that they survive under the field condition for which they have been produced. To this end,

while there has been some research done on the survival of plants produced from different modes

of propagation, relatively little has been produced regarding the long-term growth and

production. Comparisons between micropropagated plants and seedlings (Bell, van der Moezel,

Bennett, McComb, Wilkins, Marshall & Morgan, 1993) and micrpropagated plants and cuttings

(Bergman, 1998; Mulataya, Wilson, Ong, Deans & Sprent, 2002) have shown that any initial

differences between plants are generally lost after long-term growth. This is particularly

30 important for forest species where it has been suggested that root development of micropropagated plants may lead to trees being susceptible to wind-throw (Bell, et al., 1993).

1.4 Aims

Research on verticordias has been fairly limited (McComb, Arthur & Newll, 1986; George,

1991; Tyagi, Considine & McComb, 1991; Tyagi, Considine & McComb, 1992; Housten,

Lamont, Radford & Errington, 1993; Speer, 1993; McEvoy & True, 1995; Stummer, Smith &

Langridge, 1995; Egerton-Warburton, Ghisalberti & Burton, 1998; George, 2002; George &

George, 2003; Yates & Ladd, 2004), when considering the diversity of this genus and the common threat of habitat loss. The main objective of this research was to investigate and establish reliable and appropriate micropropagation, root induction and acclimatisation protocols for V. grandis in order to increase the quality and survival ex vitro, as results varied in previous studies.

1.4.1 Limitations of conventional propagation methods

There are several limitations in producing verticordias for horticultural, land restoration and conservation purposes; e.g. slow growth, limited seed production, seed recalcitrance and poor germination rates (Bell, 1999; Cochrane, et al., 2001). Conventional means of asexual propagation (cuttings and grafting) have been useful for a limited number of species, but most are not amenable to these processes. This is related to the availability of suitable cutting material from wild populations and genetic and environmental factors (Johnson, 1996). Often it

is difficult to obtain an adequate amount of the appropriate material to establish well-replicated

and controlled experiments on the development protocols; e.g. it is very difficult, if not

impossible, to obtain a large number of shoots, at the same stage of development (seasonal or

ontogenetic age) and from a single genotype, for adequate replication.

Most available verticordia clones, such as V. chrysanthella, V. mitchelliana and V. plumosa have

limited genetic variation, and have probably been selected on the basis of ease of propagation. In contrast, Verticordia grandis is a difficult-to-root species and micropropagation with consequent in vitro subcultures can induce rejuvenation or increased re-invigoration, which can increase rooting capacity.

Micropropagation has become a popular technique to overcome many of these limitations; stock plant material from difficult to propagate plants can be used to micropropagate large numbers of progeny plants.

1.4.2 Research aims

This research investigated the capacity to produce numerous explants by improving the existing micropropagation protocol (McComb, Newell & Arthur, 1986) for Verticordia grandis, as well as investigating the most successful root induction protocol for acclimatisation and survival of plantlets. This was achieved by determining the best method to achieve optimal shoots for micropropagation from mature woody cuttings. Plantlets from the clones were observed for

juvenile growth phase traits in the growth of roots and shoots and the different modes of

propagation were compared for the development of parameters that can increase growth and

survival in field conditions.

For this study 2 clones of Verticordia grandis, which have been in culture for several years in the tissue culture laboratory at Edith Cowan University, were used, and 21 different Verticordia cuttings were introduced into culture.

Cuttings were taken from 4 different Verticordia grandis plants sprouting after a fire, 10 from new growth on different mature Verticordia grandis plants from various locations around

Eneabba in Western Australia and cuttings from new growth of 7 different verticordia plants (5 from V. ovalifolia, 1 from V. fragans and 1 from V. densiflora).

The aim was to initially experiment with the existing clones of Verticordia grandis to establish reliable shoot multiplication, root induction and acclimatisation protocols and then to test the effectiveness of these on the newly introduced clones of verticordias for comparison.

1.4.3 Benefits of this research

In particular, the findings of this research could be beneficial to the reconstruction and rehabilitation of plant communities on disused mine sites, drilling exploration sites and other land cleared sites in the northern sand plain kwongan vegetation around Eneabba, Western

Australia, the natural habitat of Verticordia grandis. The results of this research could also benefit growers of West Australian wild flowers, as especially Verticordia grandis is an attractive species for the export cut flower market (Seaton, 2002).

2. MATERIALS & METHODS

2.1 Plant material

All micro-cuttings (shoots) used initially for the optimisation of the existing protocol for shoot

multiplication of V. grandis were obtained from two V. grandis clones (GRD 1; GRD 3) that had

been in culture (at Edith Cowan University’s School of Natural Sciences Tissue Culture

Laboratory) for over 5 years before experiments began.

In July 2007 explants of V. grandis (GRD) were collected from the Eneabba area; locations were

recorded with GPS readings, host plant and site descriptions were noted and all material was cut

from new shoot growth. Explant material from different new shoots of V. grandis (GF) that had resprouted after a fire were collected, as well as explant cuttings from V. ovalifolia (VOV), V. densiflora (VDF) and V. fragrans (VFR) (Table 2.1).

For each explant, a minimum of 5 - 10 cuttings of ~12cm length were collected, depending on the size of the host plant; e.g. the new shoots of the VGF clones were small, and the lignotuber was exposed ensuring that cuttings were taken from the same root. The cuttings were placed in a zip lock bag, kept on ice or at 4 0C and processed in the laboratory the next day.

2.2 Process of initiation of new clones into culture

The cuttings were surface sterilised in a 2% benzalkonium chloride in 10% ethanol solution for

20 minutes, before being rinsed 3x in sterilized milliQ water, after which they were aseptically cut into ~0.5-1cm pieces at internodes. These were then transferred onto the 5 ml of shoot multiplication media in 50 ml flasks. Initially, a total of 548 flasks were prepared for each microcutting. Each uncontaminated, surviving microcutting was transferred daily on to fresh 5ml

of shoot multiplication media in 50 ml flasks in aseptic conditions for 28 days. After 4 weeks

the surviving shoots were transferred to 250 ml culture vessels.

Table 2.1: The explant material for clones that were collected at different locations and habitats are as follows: ______

Clone______Location______Habitat______

GRD4 S29° 58’ 19.2”, E115° 18’ 51.7” Sand over laterite, heath GRD5 S29° 58’ 19.2”, E115° 18’ 48.4” Sand over laterite, heath GRD6 S29° 58’ 13.8”, E115° 18’ 51.7” Sand over laterite, heath GRD7 S29° 54’ 22.7”, E115° 15’ 02.9” Sand over laterite, heath GRD8 S29° 43’ 12.9”, E115° 13’ 33.6” Sand, heath, banksia GRD9 S29° 43’ 23.4”, E115° 13’ 21.8” Sand, heath, banksia GRD10 S29° 43’ 15.5”, E115° 13’ 18.1” Sand, heath, banksia GRD11 S29° 42’ 12.1”, E115° 13’ 29.3” Sand, heath, banksia GRD12 S29° 42’ 10.5”, E115° 13’ 29.6” Sand, heath, banksia GRD13 S29° 42’ 09.8”, E115° 13’ 27.1” Sand, heath, banksia GF1 S29° 41’ 12.9”, E115° 12’ 33.6” Burnt heath, woodland* GF2 S29° 41’ 13.6”, E115° 12’ 32.9” Burnt heath, woodland* GF3 S29° 41’ 21.4”, E115° 12’ 31.6” Burnt heath, woodland* GF4 S29° 41’ 24.8”, E115° 12’ 30.7” Burnt heath, woodland* VOV1 S29° 57’ 13.1”, E115° 16’ 15.8” Sand, heath, verge VOV2 S29° 57’ 13.9”, E115° 16’ 16.7” Sand, heath, verge VOV3 S29° 57’ 14.3”, E115° 16’ 16.8” Sand, heath, verge VOV4 S29° 57’ 14.8”, E115° 16’ 17.1” Sand, heath, verge VOV5 S29° 57’ 16.4”, E115° 16’ 19.9” Sand, heath, verge VFR1 S29° 58’ 10.8”, E115° 17’ 12.2” Remnant heath VDF1 S29° 59’ 24.3”, E115° 18’ 23.1” Private Property ______Clones GRD1, 2 & 3 were in culture at ECU; GRD2 became contaminated and could not be used in experiments. *New shoot growth from roots after original plant was destroyed by fire. All others were from new seasonal shoots.

2.3 Media preparation

All micro-cuttings (shoots) used for the optimisation of the existing protocol for shoot

multiplication of V. grandis were obtained from two V. grandis clones (GRD1; GRD3). The

initial protocol was as follows: full strength Murashige & Skoog (MS) (Murashige & Skoog,

1962) medium supplemented with 1µM 6-benzylaminopurine (BAP), 1.5 µM Kinetin, 1.25 µM

naphthalene acetic acid (NAA), 20 g L-1 sucrose, pH adjusted to 5.8, solidified with 2.5 g L-1 35

agar and 2.5 g L-1 gelrite. The media was dispensed (50 ml) into 250 ml culture vessels and

autoclaved at 121°C for 20 minutes.

2.4 Culture conditions

All prepared liquid media, nutrient solutions and growth substrates were sterilised by autoclaving

at 121°C for 20 minutes prior to application for all in vitro experiments conducted in the

laboratory. All in vitro cultures were maintained in standardised conditions (under a 16h day with irradiance of 90µmol m-2 s-1 / 8h night photoperiod, with temperature at 25°C ± 1°C) for 28 days, before being aseptically sub-cultured for shoot multiplication or utilised for further experiments.

All ex vitro cultures (explants/plantlets) were maintained in standardised conditions (10 seconds

of water misting at 7000 lux accumulated light, with temperatures at 25°C ± 5°C) in enclosed tents for high humidity (~90% - 98%) in the greenhouse for 28 days. Then the ex vitro cultures were moved out of the tents, assessed for root growth, and continued to be maintained in standardised conditions without the enclosed tent (10 seconds of water misting at 7000 lux accumulated light, with temperatures constant at 25°C ± 1°C and reduced humidity). The plants were assessed every 28 days and at 84 days the surviving plants were moved to the shade house for further hardening. The shade house did not have temperature or light control, but had automated, periodical watering (morning and evening sprinklers for 10 minutes). At 112 days the surviving plants were re-potted in a soil mixture (pastuerised) consisting of peat, perlite and white sand (1:1:1) and survival was assessed every 28 days.

2.5 Changes to shoot multiplication media composition

It was observed that using the original media formulation resulted in shoot multiplication of

GRD1 clones being unvarying in shoot length and prolific, and in GRD 3 shoot multiplication

was prolific, but with no consistent elongation; most of the shoots were less than 1cm long.

Sub-cultured shoots of uniform length (~ 2 - 5cm) from both clones were needed for further experimentation and comparisons.

2.5.1 Testing various BAP concentrations to improve elongation in clones

The original shoot sterilised multiplication media was used as a control (1 µM, BAP) against media supplemented with 0.1 µM, 0.15 µM, 0.2 µM, 0.25 µM, 0.3 µM, 0.35 µM, 0.4 µM, 0.45

µM and 0. 5µM BAP. Twenty five randomly selected shoots of clones GRD1 and GRD3 were

kept on the various media for 28 days and then harvested, after which shoot growth was

measured; this experiment was repeated 2 times, with all the results combined. The mean growth

for each concentration was calculated and the variance was analysed. The shoot multiplication

media was changed, accordingly due to the results that were achieved, for all following media

preparations.

2.6 Original root induction protocol

The success of micropropagated plantlets depends on viable adventitious root formation on shoots (with the help of exogenous growth regulators such as auxins), as this determines the survival of the new plants (Martin, 2003), but root induction for micropropagated V. grandis has proven to be difficult. The shoot length could also determine the efficiency of root formation, as longer shoots are in a better physiological condition for root induction (Xu, et al., 2008).

For root induction experiments shoots of GRD1 (each 4.5 - 5 cm long) were randomly chosen from numerous culture vessels, leaves removed from the lower part of the stem (~1cm) as wounding of the stem tissue can affect the balance of growth regulators, which can enhance root induction (Marks & Simpson, 2000). Shoots were grown in sterile rooting media (50ml) modified from Bennett, et al., (2004); it contained ¼ strength MS macronutrients and full strength micronutrients supplemented with 10 µM indol-3-butyric acid (IBA concentration depending on experiment), 2% sucrose, pH adjusted to 5.5, solidified with 2.5 g L-1 agar and 2.5 g L-1 gelrite in 250 ml clear culture vessels. Each vessel contained 50 ml of sterilised medium.

The control rooting medium had no IBA. Each culture vessel contained 5 shoots and these were maintained in the growth room at standard conditions for 28 days, after which the results were analysed.

This protocol was used for all experiments – auxin pulsing duration and concentrations were changed in some of the following experiments. The shoots were always kept on the rooting maintenance media with no auxins for 28 days before being evaluated.

2.6.1 Optimisation of duration of IBA treatment for root induction

IBA has long been one of the preferred and active auxins to promote root formation in in vitro

cultured plant shoots (Epstein & Muller, 1993). Previous root initiation trials with the existing

root induction protocol for V. grandis by McComb, Newell & Arthur (1986) showed inconsistent and poor root growth of the V. grandis clone for each trial. In order to determine the optimum

IBA pulsing period for root induction, shoot replicates were grown on rooting media supplemented with 10µM of IBA (as per root induction protocol by Bennett, et al., 2004).

For each day of treatment 25 shoots of GRD1 were randomly chosen and the media of the control was not supplemented with IBA. The experiment was designed for 7 days; on each day

5 shoots, from 5 randomly selected vessels, were transferred to rooting media that contained no

IBA. These were then kept in this media for 28 days, the experiment was repeated twice; the numbers of roots were counted and the means calculated for each day of pulsing. For this experiment only the root numbers/shoot were analysed in order to determine the most effect IBA pulsing time. This was repeated with GRD3 clones.

2.6.2 Optimisation of IBA concentration for root induction

The root formation performance is affected by the concentration of auxin, the type of auxin and the plant species. IBA is often the preferred auxin for root induction, as plant tissues can rapidly oxidise other auxins such as IAA (DeKlerk, et al., 1997). IBA can either promote the formation

of adventitious and lateral roots, or it can inhibit this process – this variation depends on the IBA

concentration and the plant species (Muller, 2000).

For this experiment different IBA concentrations were tested to determine the optimal

concentration to achieve the greatest number of shoots with root induction and growth. The

randomly selected shoots (GRD1), 25 for each treatment, were pulsed for 6 days on media

supplemented with different IBA concentrations (0, 2.5, 5, 10, 20, 40, 80 and 160µM). After this, all shoots were randomly transferred to culture vessels rooting media without IBA, visible root formation was recorded for each shoot on a daily basis and then totals scored after 28 days for statistical analysis and evaluation.

2.6.3 Trials with auxin combinations for root induction

In contemporary plant science research it has become evident that endogenous plant auxins interact with exogenous auxins resulting in diverse fast and slow responses (Woodward & Bartel,

2005). Different auxin combinations have been successful for root induction in numerous plant

species due to their various properties (Martin, 2003; Martin, 2003/4; Pati, Rath, Sharma, Sood

& Ahuja, 2006; Nandagopal & Kumari, 2007), so several combinations of auxins were

experimented with in order to achieve more overall consistency in root response and to

determine the optimal exogenous auxin treatment for root induction.

For this experiment it was assumed that 80 µM of an auxin combination would give a comparable result, as was achieved with 80 µM IBA, in order to determine greatest effectiveness for root induction and formation. The randomly selected shoots (GRD1), 25 for each treatment, were pulsed for 6 days on media supplemented with different auxin treatments: Treatments: no auxin, IAA (80µM), NAA (80µM), IAA (40µM) + NAA (40µM), IAA (40µM) + IBA (40µM),

NAA (54µM) + IAA (26µM), NAA (40µM) + IBA (40µM), IAA (54µM) + NAA (26µM), IAA

(26µM) + NAA (26µM) + IBA (26µM) or IBA (80µM - control). After this, all shoots were randomly transferred to culture vessels rooting media without auxins, visible root formation was recorded for each shoot on a daily basis and then totals scored after 28 days for statistical analysis and evaluation.

2.7 Trials with different in vitro substrates for root induction

In 2003, Newell and co-workers reported results of experiments on shoot microcuttings of

Verticordia species using a porous agar medium that contained air filled porosity of 10% to 29% to enhance root formation. The researchers concluded that root induction and length was significantly increased compared to standard agar solidified media (Newell, 2003). Another alternative in vitro rooting substrate being researched by Newell et al., (2005) involves in vitro – soil medium (IVS), which consists of an aerobic rooting medium and the results were compared with agar-solidified media. It was concluded that for numerous woody perennials (including species from Myrtaceae) the rooting performance in IVS was successful and this method could reduce the stress and plant loss during the acclimatisation stage of micro cuttings from sterile in vitro conditions to in vivo field conditions (Newell, 2006). Newell, et al., (2005) and Yan &

Sedgley (2006) experimented with different substrates including IVS (in vitro soil media) instead of agar based media on difficult to root recalcitrant woody plants to increase root formation and vigour.

For this experiment different substrates were trialled; some were commercially available whilst others were prepared prior to the experiment. The shoots (25 for each treatment, randomly chosen) were pulsed on agar/gelrite media supplemented with 80 µM IBA and after 6 days these were randomly transferred to culture vessels that contained a uniform amount of sterilised substrate (autoclaved at 121oC for 20 minutes for each different substrate). This was supplemented with 50 ml sterilised liquid nutrient solution containing ¼ strength MS macronutrients and full strength micronutrients supplemented with 2% sucrose, pH adjusted to

5.5. The substrates included: Agar/gelrite, White sand (Ø ≤ 1 mm), white sand (Ø ≥ 1 mm), sterilised soil (2 peat: 1perlite), Jiffy pots®, black sand (Ø ≤ 1 mm) and rockwool®. This was repeated with fewer treatments that included: Agar/gelrite, blacks and sterile soil (2 peat: 1 perlite).

2.7.1 Comparison of sterilized soil with different nutrients

In order to test if GRD1 clones will initiate roots (after a 6 day IBA pulsing) in sterilized soil without additional nutrients, the following experiment was conducted: 25 shoots for each treatment (randomly chosen) were pulsed on agar/gelrite media supplemented with IBA and after

6 days these were randomly transferred to culture vessels that contained a uniform amount of sterilised sterilised soil (2 peat: 1 perlite). According to treatment some were supplemented with

50 ml sterilised liquid nutrient solution containing ¼ strength MS macronutrients and full

strength micronutrients supplemented with 2% sucrose, pH adjusted to 5.5 (control), some with

just 50 ml sterilised milliQ water supplemented with 2% sucrose, pH adjusted to 5.5 and the

others just with 50 ml sterilised milliQ water. These culture vessels were kept in the culture room

for 28 days, after which the roots were scored. The entire root system of each shoot was carefully

separated and scanned for root surface area and length for comparison and statistical analysis.

2.7.2 Comparison of in vitro sterilized soil to ex vitro soil

In order to test if GRD1 clones will form roots (after a 6 day IBA pulsing) in soil ex vitro without additional nutrients, the following experiment was conducted: 25 shoots of GRD1 for each treatment (randomly chosen) were pulsed on agar/gelrite media supplemented with IBA and after

6 days these were randomly transferred to crackpots (direct soil) containing the same soil composition as the control (sterilised soil in culture vessels that contained a uniform amount of soil media supplemented with 50ml sterilised milliQ water, no nutrients). The culture vessels were kept in the culture room and the other shoots were transferred to the greenhouse in seed trays with vented lids at a temperature regime of 20-30 oC in high humidity (~90% - 98%).

After 28 days the roots were scored; the entire root system of each shoot was carefully separated,

scanned for root surface area and length and root numbers were counted for comparison and

statistical analysis. All fresh shoots were separated from all roots and weighed then all shoots

and roots were dried at 70º C for 24 hours and weighed again to determine the shoot and root

biomass for statistical analysis.

2.8 Survival in greenhouse during acclimatisation phase

For comparison 180 shoots of GRD1 clones (after in vitro IBA pulsing) were transferred to

sterilized soil (with just 50ml millQ water) in vitro and then after 28 days were re-potted in to crackpots containing the same soil mixture, which were placed randomly in humidity tents that allowed for retention of high humidity (90-98%) with periodical misting. In order to facilitate

rooting simultaneously with acclimatisation 180 shoots of GRD1 clones (after in vitro IBA

pulsing) were, at the same time, placed randomly (arranged in a randomised complete block

design) in crackpots containing a uniform soil mixture in humidity tents that allowed for

retention of high humidity (90-98%) with periodical misting. The survival rates for both

treatments were scored in intervals of 28 days for the initial acclimatisation in humidity tents,

and then the shoots were moved out of the humidity tents to reduce humidity (~60%). All shoots remained in the greenhouse until roots were visible at the bottom of the crackpots or until they died. All were scored after 56 days. The surviving plantlets were scored every 28 days, and then remaining shoots were moved to a shady area in the glasshouse (at 84 days) where the humidity was less, but a periodical watering regime was in place. After 112 days the surviving shoots were scored again and then moved to the shade house for further in vivo acclimatisation. The shade house had a restricted water regime of once in the morning (10 mins) and once in the afternoon

(10 mins). These surviving plantlets were then re-potted into larger pots containing the peat and perlite mixed with sand (1:1:1) for additional drainage. Long-term survival after re-potting continued to be assessed every 28 days.

The pulsed shoots were transferred to the greenhouse on March 8th, 2008, and this process was

repeated 6 times. Again after 112 days all surviving plants were moved to the shade house with

ambient temperatures and regular watering. Long-term survival was monitored for over 252 days in total.

2.9 In vitro shoot multiplication of new GRD clones

Following the success of the shoot multiplication protocol for GRD1 & 3 the effectiveness was tested for other clones. Shoot multiplication for new clones introduced into culture (Table 2.1) was not achieved for all clones; VOV, VDF and VFR did not produce shoots that could be sub- cultured over time. Shoot multiplication proved to be difficult for GRD 4, 5, 7, 8 and 13. The most successful shoots for multiplication were GRD 6, 9, 10, 11, 12 and GF1, 2, 3 and 4 (no

results shown here), as these clones continuously produced multiple and uniform shoots. The

improved protocol for shoot multiplication was used on all new clones to produce comparable

shoots with a uniform length of ~4.5-5cm for root induction.

2.9.1 Root induction of successful new clones

Root induction trials with the most successful new clones commenced approximately after 1 year in culture. The optimised root induction protocol was applied to shoots that were pulsed on

IBA/agar media and then transferred to crackpots with a uniform soil mixture, placed in the humidity tents in the greenhouse and monitored. In order to facilitate rooting simultaneously with acclimatisation, 40 shoots of various clones (after in vitro IBA pulsing) were placed randomly (arranged in a randomised complete block design) in crackpots containing the uniform

soil mixture in humidity tents that allowed for retention of high humidity (90-98%) with periodic

misting. The results were scored every 28 days for analysis. These trials were repeated 3 times

during 2008/09. The means for survival were calculated (rounded to a complete number) and

results were combined for final analysis for all plantlets that performed well in culture. The

following clones were used because of their similar physiology; e.g. fairly uniform length, large

leaves and vigorous appearance – GRD 6, 9, 10, 11, 12, GF1, 2, 3, and 4 (although 1 and 2 had

longer internodes than 3 and 4). All surviving plantlets were re-potted at 140 days (28 days later than the experiments with GRD1) in peat and perlite mixed with sand (1:1:1) for extra drainage.

Re-potting was done later (compared to trails with GRD1) because not all plantlets had roots visible simultaneously by 56 days.

2.10 Statistical analysis

All experiments were randomised (or complete randomised block design was used) and analysis of variance (ANOVA), as a linear model, was applied to determine differences of means between clones and treatments. Levene’s test of homogeneity of variance was applied to all data sets and where variance were found to be unequal a log + 1 transformation was performed. Different post hoc tests were used to determine differences between treatments; Tukey’s B multiple-range test

(P ≤ 0.05), or Dunnet’s t-test, where all groups were compared against the control group. All root

lengths and surface areas were measured using an automatic root analyser – Delta-T-Scan and

Delta-T-Scan software. For the analysis of long-term survival of rooted explants in the shadehouse a χ2 test was used and resulting values were compared to a χ2 critical table to

determine significant probability values (P ≤ 0.05). SPSS version 17 was used for all statistical

analysis.

3. RESULTS

3.1 Survival of new clones initiated into culture

A total of 380 shoots did not survive the first 2 weeks because of shoot death or contamination.

Many shoots had varying axillary bud formation and shoot elongation, some to the extent that

sub-culturing was not possible; these shoots were just transferred to fresh media every 28 days.

Following subsequent sub-culturing of the clones every 28 days many more shoots died; after 6

months there were no V.ovalifolia (VOV), V. fragrans (VFR) or V. densiflora (VDF) shoots

remaining (Table 3.1).

Table 3.1: Percentage Survival of new microcuttings over a 28 day period with cuttings being transferred to new media in 50ml flasks daily. ______Survival (%) Clone Start Week 1 Week 2 Week 3 Week 4 ______(n)______

GRD4 18 39 33 22 22 GRD5 25 64 40 28 24 GRD6 35 69 69 69 69 GRD7 42 50 45 38 31 GRD8 34 68 65 56 56 GRD9 17 71 59 53 47 GRD10 28 89 86 79 79 GRD11 18 56 28 22 17 GRD12 23 65 30 13 13 GRD13 27 70 41 37 37 GRD14 32 69 41 34 31 GRD15 24 38 21 17 17 GRD16 17 47 35 35 35 GRD17 48 44 33 25 23 VOV1 27 37 30 22 0 VOV2 14 29 14 14 0 VOV3 28 57 28 28 7 VOV4 17 94 29 24 0 VOV5 21 57 43 43 5 VFR1 29 52 52 45 45 VDF1 24 54 46 42 37 ______

3.2 Changes to shoot multiplication protocol

It was observed that using the original media formulation resulted in shoot multiplication of

GRD1 clones having variable shoot length and proliferation, and in GRD 3 shoot multiplication was prolific, with but with no consistent elongation; most of the numerous shoots were less than

1 cm in length. Sub-cultured shoots of uniform length (~ 2 - 5 cm) from both clones were needed for further experimentation and comparisons. In order to improve shoot elongation necessary for shoot multiplication and proliferation of unvarying shoot lengths it was decided to modify the

BAP concentration supplemented to the growth media.

The highest number of shoots with a more uniform shoot height was achieved at 0.25µM BAP, as the shoots of both clones (GRD1 – 2.3 cm & GRD3 – 2.1cm) were comparable for shoot elongation (Fig 3.2). All other BAP concentrations produced shoots of varying lengths (the shortest growth 0.7cm for GRD3 at 1µM and GRD1 was 1.1cm at 0.45 µM). The new shoots appeared much more vigorous and healthier in growth (a more visibly uniform leaf size and uniform shoot lengths) compared to most explant shoots before the experiment. The most unvarying height for both GRD1 & GRD3 shoots were achieved at 0.25 µM of BAP added to the shoot multiplication media. These results substantiated research that suggests that axillary bud formation and shoot height could be promoted by using low concentrations of BAP.

Consequently the BAP concentration was changed to 0.25 µM for all following shoot multiplication media.

3.0

GRD1 2.5 b b b GRD3 a a 2.0 a a a a a a a 1.5 a a a a a a a a a a 1.0 Meanshoot height (cm)

0.5

0.0 0 0.1 0.15 0.2 0.25 0.3 0.35 0.4 0.45 0.5 1 BAP concentration (mM)

Figure 3.2: Comparisons of different BAP concentrations to improve shoot heights in vitro. The concentrations of 0.25 and 0.3µM BAP were significantly different (P ≤ 0.042) for GRD 1. For GRD 3 the concentration of 0.25µM BAP was significantly different (P ≤ 0.029). Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

3.3 Changes to root induction media composition

The most commonly used auxins for root induction includes indole-3-butyric-acid (IBA), indole-

3-acetic-acid (IAA) and α-naphthalene-acetic-acid (NAA), with IBA often being the most

common one used for root induction. In previous root initiation trials, using the existing root

induction protocol (root induction media supplemented with 10µM of IBA) for V. grandis by

McComb, Newell & Arthur (1986), the results showed inconsistent and poor root growth of the

V. grandis clone (GRD1) for each trial. It was decided to determine optimal IBA pulsing duration and concentration.

3.3.1 Optimisation of duration of IBA pulsing

In order to determine the optimum IBA pulsing period for root induction, shoot replicates were grown on rooting media supplemented with 10µM of IBA. Root formation differed for the shoots that were pulsed with IBA for 5 and 6 days; root initials became visible just 7 days after the transfer from IBA pulsing media to non-IBA root growth media, and most shoots in all of these culture vessels had adventitious root growth visible after 17 days (Figure 3.3a). Shoots that were pulsed with IBA for 4 days showed root growth on most shoots after 23 days, whereas the shoots that were pulsed with IBA for 1, 2, 3 and 7 days showed poor root induction, with many of these shoots not forming roots at all. The highest percentage of root formation after 28 days was only 45% after 6 days pulsing on 10 µM IBA, which was very low when compared to the original root induction protocol results (McComb, et al., 1986).

4 a) b

3 b

b 2

1 a Mean number of roots per shoot per roots of number Mean a a a

0 100 b)

40 b Percentage rooting Percentage

b 20 a a a a 0 1 2 3 4 5 6 7

IBA pulse duration (days)

Figure 3.3: Comparison of duration of IBA pulsing for root formation and growth in vitro. a) The means of roots for days 1, 2, 3 & 7 were not statistically different (P ≤ 0.125), but different to the means of roots (P ≤ 0.031) for days 4, 5 & 6 which had no statistical difference. b) The greatest percentage of roots after 28 days resulted with shoots pulsed for 6 days. Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

Some shoots of GRD3 clones readily displayed root formation on shoot multiplication media so this experiment was repeated twice for GRD3 clones. For GRD3 only the control treatment had one shoot with visible roots compared to the other treatments, which had no results (after 28 days all shoots had no visible root formation); consequently no analysis was possible.

All further experiments for root induction only used GRD1 clones and 6 days auxin pulsing for root induction.

3.3.2 Optimisation of IBA concentration

Root formation was greatest for the concentration of 80 µM IBA with a mean of 7 ± 1.2 roots/shoot, followed by 6.3 ± 1.0 roots/shoot for 40 µM IBA and 5.7 ± 1.1 roots/shoot at 20 µM

IBA (Fig 3.4). Root initials for these treatments became visible by day 5 and 6, after transfer to media without IBA. Root formation on shoots pulsed on media with the other IBA concentrations became visible much later (day 18-25), if at all, with many shoots not producing any roots. The lower concentrations (0=control, 2.5µM, 5 µM & 10 µM) of IBA had little effect on root response and formation. All further experiments used 80 µM of auxin for root induction pulsing.

10 a)

b 8 b b

Mean number of roots per shoot per roots of number Mean a 2 a a a 0

120 b)

100 b b b 80

Percentage rooting Percentage 40

20 a a a a a 0 0 2.5 5 10 20 40 80 160

IBA Concentration (uM)

Figure 3.4: Comparison of in vitro root induction and growth using different IBA concentrations. a) Mean roots per shoot. Treatments with 0, 2.5, 5, 10 and 160 µM IBA were not statistically different (P ≤ 0.237), but different to the means of roots (P ≤ 0.029) for treatments with 20, 40 and 80 µM IBA were not statistically different. b) Mean percentage rooting. Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

3.3.3 Auxin combinations for in vitro root induction

Root formation in the control (80 µM IBA) was again visible 6 days after transfer to an auxin free media and after 17 days 24/25 shoots had roots from this treatment (Fig 3.5). Root formation for all other treatments were significantly lower and most roots appeared visible after 19 days.

The treatment with no auxin had 1/25 shoots with 2 small (>0.3cm) roots. Treatments containing

NAA (3, 4, 6, 8 and 9) produced large calluses at the shoot base and means of 0.8 ± 0.3 and

1.7±0.4 roots/shoot. IAA did was not very effective for root induction; Treatments 2 and 4 only produced a mean of less than 1 ± 0.2 root/shoot. Treatment 5 and 7 contained an auxin combination with IBA and produced means of 3.4 ± 0.5 and 3.8 ± 0.7 roots/shoot, which were more roots than treatments without IBA, but these results were not statistically different. For all further experiments 80 µM IBA was used for root induction.

Commercial auxin applications (Clonex powder & gel for soft and semi-hard wood) and prepared auxin solutions for dipping the shoots prior to transfer to soil for root induction and simultaneous acclimatisation was also trialled. However, within a few days all shoots were dead, so this was not further investigated and as no results were obtained, they were not recorded.

Treatments: 1. no auxin 7 2. IAA (80uM) 3. NAA (80 uM) 4. IAA (40uM) + NAA (40 uM) b 6 5. IAA (40 uM) + IBA (40 uM) 6. NAA (54 uM) + IAA (26 uM) 7. NAA (40 uM) + IBA (40 uM) 5 8. IAA (54 uM) + NAA (26 uM) 9. IAA (26 uM) + NAA (26 uM) + IBA (26 uM) 10. IBA (80uM) control a 4 a

a 2 a

Mean number of roots/shoot of number Mean a a 1 a a a 0 0 2 4 6 8 10 Auxin treatment

Figure 3.5: Comparison of in vitro root induction and growth using different auxin combinations and concentrations. There were no differences (P ≤ 0.351) between treatments 1 – 9, compared to the control treatment 10 that was significantly different ( P ≤ 0.017). Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

3.4 Different substrates

Many alternatives to gel media have been investigated; e.g. foam, vermiculite, vermiculite and gelrite, rock wool, filter paper bridges, glass beads, coir, luffa –sponge, jute fibre and in vitro – soil / IVS, as these substrates can facilitate improved gas exchange conditions.

Trials with different substrates (Figure 3.6) resulted in good rooting response with black sand

(8.1 ±2.7 roots/shoot, many branched with lateral roots), rock wool (6.6 ±2.9 roots/shoot) and agar (4.4 ±2.0 roots/shoot, few branched with lateral roots). Also shoots grown in black sand and rock wool appeared different to those grown in agar medium by having more, longer and some laterally branched roots. The first roots became visible in agar after 7 days, in rock wool several roots protruded at the sides after 13 days and in black sand some roots were visible at the side and bottom of the culture vessels by day 10. In the other treatments, the first roots became visible later; white sand (0.9 ±2.7 roots/shoot) by day 17, coarse white sand (1.7 ±2.9 roots/shoot) by day 16, sterile soil (2.7 ±2.8 roots/shoot) by day 14 and roots protruded from soil jiffies (3.3 ±2.8 roots/shoot) by day 14. Overall, the early visibility of roots in the different substrates suggests a faster rooting response (compared to previous trials with agar). However, the individual roots growing from shoots in agar were the only ones that could be clearly observed on a daily basis.

Treatments: Agar = control 10 Sand 1 = white sand ( 1mm)  Sand 2 = white sand ( 1 mm) b St soil = sterilised soil (peat:perlite 2:1) 8 Jiffies = commercial brand Sand 3 = black sand ( 1 mm) b Rockwool = commercial brand 6 b

4 a a

Mean number of roots/shoot of number Mean a 2 a

0 Agar Sand 1 Sand 2 St Soil Jiffies Sand 3 Rockwool Growth substrates

Figure 3.6: Comparison of in vitro root induction and growth in different substrates. The means of roots between treatments white sand (≤1mmØ), coarse white sand ( ≥2mmØ), sterile soil (peat : perlite = 1:3) and commercial soil jiffies were not statistically different (P ≤ 0.276), but different compared to the means (P ≤ 0.022) of the roots between treatments black sand (≤1mmØ), commercial rock wool cubes (5cm3) and the control (agar/gelrite medium) which were not significantly different. This experiment was repeated twice to compare treatments Control, Sand 3 and Rockwool only, as they had higher means of root numbers per shoot, confirming no significant difference (P ≤ 0.153). Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

The roots from shoots grown in soil jiffies and rock wool were problematic to score, as each contained 5 shoots and the roots were difficult to completely separate and remove intact after 4 weeks of growth. However, each shoot had formed at least 1 root (often branched with lateral roots), and some shoots had small white outgrowths from the swelling at the shoot base; it is possible that these could have developed into roots if given more time.

3.4.1 Comparison of agar, black sand and sterile soil

The root formation of GRD1 clones in black sand was statistically comparable with agar (Figure

3.6), so the affect of root induction in these substrates was compared to sterile soil rooted shoots.

Root numbers alone, where short and long roots are counted as equal, are not always an ideal

parameter when comparing plant root systems and therefore root surface area and root length

were also measured.

There were no statistical differences for means of root numbers (P=0.012) between treatments

(Figure 3.7a). The root surface area means in Figure 3.7b (13.9cm2 ± 1.1) for shoots maintained in sterile soil were statistically different compared to root surface area means of roots grown in agar (8.4cm2 ± 1.6) and in black sand (11.9cm2 ± 1.8). The sterile soil and black sand roots (less in black sand) were visibly different as well; these had more branched lateral root formation compared to roots from agar that consisted of single roots of varying lengths. It was also observed, whilst separating the roots from the substrate, that not all roots were submersed in the nutrient liquid. The mean lengths (Figure 3.7c) were for roots grown in sterile soil (6.2cm ± 2.0) were statistically different compared to roots grown in black sand (5.7cm ± 1.8) and agar (4.3 ±

1.7). Roots grown in agar were mostly single roots with no branching, compared to the branched roots of various thicknesses from the other treatments of in vitro black sand and in vitro sterile soil (Fig 3.8).

a) a a 6 a 4

2 Mean number of roots/shoot of number Mean

0 6 b)

) b 2 5 a a 4

1 Mean root surface area (cm area surface root Mean

0 16 c) b 14 a 12

10 a 8 6 4 Meanroot length (cm) 2 0 Agar Sterile Soil Black sand Black

Substrate

Figure 3.7: Comparison of in vitro root induction and growth in different substrates a) Mean roots per shoot. nsd (P=0.012). b) Mean root surface areas grown in agar/gelrite = control were statistically not different to black sand (P=0.113) compared to roots grown in sterile soil (P=0.014). c) Means of root length grown in agar/gelrite = control were statistically not different to black sand (P=0.245) compared59 to roots grown in sterile soil (P=0.037). Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

a) b) c)

Figure 3.8: Comparison of harvested and washed roots grown in different in vitro substrates. a) Shoot with root formation grown in agar. b) Shoot with root formation grown in black sand. c) Shoot with root formation grown in sterile soil. For further in vitro experiments only sterile soil was used.

3.5 Comparison of sterile soil with different nutrients The previous experiment achieved good root induction on GRD1 clones using the aerated, porous substrate of sterile soil instead of an agar medium. This led to experimenting root induction with sterile soil supplemented with just sucrose, with complete nutrients and with just DDI water as a control.

The mean root numbers for sterile soil +DDI (4.1 ± 1.2), sterile soil + sucrose (4.6 ±

1.8) and for the control, sterile soil + nutrients (5.6 ±1.7) were not statistically different

(Fig 3.9a). The means for root surface area (Fig 3.9b) for shoots grown in sterile soil +

DDI (3.9 cm2 ± 1.6) and for shoots grown in sterile soil+ sucrose (5 cm2 ± 2.0) were statistically different from shoots grown in the control of sterile soil + nutrients (11.8 cm2 ± 1.9). The root surface area for the control treatment (sterile soil + nutrients) was also less than in the previous experiment (Fig 3.7b, 13.9 cm2 ± 2.0). The mean root lengths for sterile soil +DDI (4.4 cm ±0.4), sterile soil + sucrose (4.6 cm ±0.1) and for

60 the control, sterile soil + nutrients (6 cm ± 0.6) were not statistically different (Fig 3.9c).

This experiment showed that root induction and growth was possible without added nutrients in the substrate, so just soil was used for further experiments.

7 a) a 6 a 5 a 4

3 2

Mean number of roots/shoot of number Mean 1

0 7 b) b ) 2 6 a 5 a

3 2

Mean root surface area (cm area surface root Mean 1

0 140 c) a 120

100

60 a a 40 Meanroot length (cm) 20

Sterilesoil & DDi Sterilesoil & sucrose Sterilesoil & nutrients

Sterile soil with different treatments

Figure 3.9: Comparison of in vitro sterile soil with different nutrients (no nutrients). a) Mean root numbers b) Mean root surface c) Mean root Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

3.6 Ex vitro root formation and acclimatization

Combined in vitro shoot multiplication with ex vitro root formation and acclimatisation has been

trialled to enhance survival of plantlets, as well as a cost and time reducing strategy. For this

experiment in vitro (sterile soil) and ex vitro (direct soil in greenhouse) root formations were

compared.

It was not possible to observe when root initials began to appear and compare for either

treatments. Root numbers (Fig 3.10a) were not statistically different (sterile soil 2.1 ± 0.2

roots/shoot, direct soil 1.9 ± 0.1 root / shoot). The mean root surface area for sterile soil was 7.9

cm2 ±0.9 compared to 6.3 cm2 ± SE 1.0 for direct soil. Even though the results were statistically different (Figure 3.10b), each shoot had roots after 28 days. After 28 days of growth the mean root length in sterile soil was 4.8 cm ±0.4 and the mean length in direct soil was 4.7 cm ± 0.4,

which were statistically not different (Figure 3.10c). Root morphology appeared similar, as root

branching were visible on roots from both treatments. Means of dry shoot biomass (Figure

3.10d) were not statistically different between treatments (sterile soil 0.88g ± 0.3, direct soil

0.64g ± 0.3) and the means for dry root biomass (Fig 3.10e) were also not statistically different

(sterile soil 0.53g ± 0.2, direct soil 0.48g ± 0.2).

1.2 a) d) 0.6 a a 1.0 a 0.5 0.8 a 0.4 0.6 0.3 roots/shoot Meanshoot 0.4 dryweight (g) Meannumber of 0.2

0.2 0.1

0.0 0.0

2.5 b) 6 e) a a a 2.0 a 5 ) 2 4 1.5 3

Mean root Mean 1.0 root Mean

dryweight (g) 2 surface area (cm area surface 0.5 1

0.0 0 root root 10 c) induction a induction Invitro 8 vitro Ex a 6 Mean root Mean

length(cm) 4

0 root root induction induction Invitro Ex vitro Ex

Figure 3.10: Comparison of in vitro and ex vitro root induction and growth after being in soil for 28 days. a) The mean number of roots per shoot b) Mean root surface areas c) Mean root d) The mean shoot dry weights e) Mean root dry weights. Means were analysed using Anova; n=25; different letters above bars indicate significant differences (ANOVA, Tukey’s test P ≤ 0.05); Error bars = ± SE.

3.7 Survival

The greenhouse environment consists of a septic environment with higher light levels and lower relative humidity which are the opposite to in vitro conditions. Due to these different ambient

conditions transfer from in vitro to ex vitro is stressful for the plantlets, but the main success of

micropropagation largely depends on the plantlets capability to resume growth during and after

acclimatisation. This experiment was started simultaneously for all shoots at 28 days following

the completion of IBA pulsing and long-term survivals were recorded (Figure 3.11)

Shoots kept in in vitro conditions for root growth, then transferred to soil in crackpots after 28

days, did not acclimatize easily to the greenhouse conditions; many wilted early which often

resulted in death. After 140 days there was 100% mortality recorded for the in vitro rooted

shoots. There was a high survival rate for shoots transferred to direct soil immediately after the

IBA pulsing period in ex vitro conditions for acclimatisation in the greenhouse. The greatest plant loss occurred at 28 days when plants were moved out of the enclosed crackpot trays (high humidity) into lower humidity levels of the greenhouse. The greatest shoot loss was observed in the first 28 days for shoots transferred to the humidity tent in March. Some plantlets did not acclimatise and survive every time environmental conditions were changed (transfer to lower humidity; e.g. from closed trays, to open trays, from greenhouse to shade house); the greatest mortality rate was scored for the first trial. After a total of 6 repeats (Table 3.2) it was evident

that with each change of environmental conditions total plantlet mortality increased, as some

failed to acclimatise and thrive. After 112 days all surviving plantlets were re-potted and the following mortality rates (140 days) were further recorded. Following re-potting, most of the 64

surviving plantlets displayed new shoot growth: either branching or shoot elongation, which

occurred at the end of winter and in early spring. After 252 days more than 70% of the plants (ex

vitro root growth) had survived (Figure 3.11; Table 3.2).

100 in vitro root growth ex vitro root growth

40 Percentage survival Percentage 20

0 28 56 84 112 140 168 196 224 252

Time since transfer to soil (days)

Figure 3.11 Periodic assessment of ex vitro grown roots and in vitro grown roots of GRD1 (180), acclimatisation and survival (initial experiment) was assessed every 28 days after the initial transfer to soil. All plants had visible roots after 48 days. Plants were re-potted into bigger pots with fresh soil (peat, perlite, sand 1:1:1).

The experiment with in vitro pulsed shoots and ex vitro root growth combined with simultaneous

acclimatisation was repeated with larger sample sizes and the results were recorded (as follows

in Table 3.2). After 252 days more than 72% of the GRD1 plants survived simultaneous ex vitro

root growth and acclimatisation.

Table 3.2: Results from repeat experiments of survival rates of 1080 GRD1 shoots (ex vitro root induced) in a total of 27 trays @40 shoots each) during 2008 & 2009.

Period Plants survived Percentage

28 days 1068 98.9%

56 days 1029 95.3%

84 days 987 91.4%

112 days 956 88.5%

140 days 845 78.2%

168 days 839 77.6%

196 days 827 76.6%

224 days 813 75.3%

252 days 786 72.8%

______

During the following 2 years after these experiments started most plants had died, even though they were initially thriving for the duration (252 days) of the experiment.

3.8 Effectiveness of improved protocols on new clones

3.8.1 Shoot growth and maintenance

The improved shoot multiplication was not successful for all new clones introduced into culture.

All clones of VOV, VDF and VFR did not produce shoots that could be sub-cultured over time, or they eventually died. The most successful new clones for shoot multiplication were GF 1, 2, 3 and 4, GRD 5, 8, 9, 10, and 11; these all produced shoot material that was appropriate for subcultures and for root induction trials.

3.8.2 Root induction, growth and acclimatisation

Root induction trials with the most successful new clones commenced approximately after being in culture for over 1 year. The optimised root induction protocol was applied to shoots where root induction and acclimatisation would occur simultaneously. Shoots were pulsed on IBA/agar media and then transferred to crackpots with IVS, placed in the humidity tents in the greenhouse and monitored. Long-term survival rates were monitored.

The results from Figure 3.12 show that only an average of 51% of plantlets survived acclimatisation in the long term (GRD3 & GRD4 did not survive). The majority of plantlets showed roots at the bottom of the crackpots by 84 days and initial survival rates were relatively high during this period. At 112 days ~79% of the total plantlets were re-potted, but in the

1 remaining weeks of the experiment many plantlets did not survive. After 1 /2 years post ex vitro transfer most of these plantlets had not survived and during this time very few displayed visible shoot growth or vigour.

GF1 GF2 GF3 GF4 GRD3 GRD4 GRD5 100 GRD8 GRD9 GRD10 80 GRD11

40 Percentage survival Percentage 20

0 50 100 150 200 250

Time since transfer to soil (days)

Figure 3.12: Periodical assessment of acclimatization and ex vitro survival over 252 days of 11 clones of V. grandis (40 shoots of each clone). Plants were assessed every 28 days and all surviving plants had visible roots 84 days after transfer to soil.

4. DISCUSSION

4.1 Collection and survival of new clones initiated in to culture

Micropropagation has become an invaluable tool in the plant propagation industry; e.g.

ornamental horticulture and agriculture (Johnson, 1996; De Jeu & Cadic, 2000; DeKlerk, 2002;

Sarasan, et al., 2006). This propagation method presents a viable option to achieve commercial

success for the production of disease free plants in large quantities (Bandyopadhyay, et al., 1999)

and in general is based on the principle of establishing and maintaining healthy micro-shoots in

culture for multiplication so that these produce rooted micro-cuttings and plantlets (DeKlerk,

2002; Liu & Bao, 2003). There are no effective vegetative propagation techniques for

reproducing numerous mature plants that can equal this.

New clones of V. grandis and other verticordia species were introduced into culture to test if optmised shoot multiplication and root induction media could be successfully applied to these.

The introduction of new clones was only partially successful (Table 3.1), even though the source plants for cutting materials were carefully selected from wild verticordia populations in natural habitats around Eneabba. Cuttings from different individual explants are likely to behave differently under culture conditions (Geneve, 1989; Anthony, McLean & Lawrie, 2000). Most of the clones that did not survive were contaminated either by exogenous and/or endogenus microorganisms which proved to be detrimental (Idczak, & Brielmaier-Liebetanz, 2003). The surviving clones were sub-cultured for over 1 year and numerous viable cultures were produced for further experiments.

4.2 Changes to shoot multiplication protocol

Some research suggests that high BAP concentrations could inhibit shoot elongation (Lall,

Mandegaran & Roberts, 2006), as well as stunt the growth of shoots and reduce the number of shoots produced (Martin, 2003). This was evident when using the original shoot multiplication protocol (McComb, et al., 1986); in the clones GRD1 and in GRD3 shoots were not consistently elongated, but shoots of 2-5cm in length were needed for further experiments. The results

(Figure 3.2) substantiated the research that suggests that axillary bud formation and shoot elongation could be promoted by using very low concentrations of BAP; e.g. 0.2µM to 0.5µM

(Chalupa, 1987, Lall, et al., 2006).

BAP and kinetin are generally considered as anti-senescence hormones and are known to reduce the apical meristem dominance and stimulate both axillary and adventitious shoots formation from meristematic tissues, resulting in shoot multiplication. Like auxins, cytokinins can produce a paradoxical effect whereby they produce an effect or its opposite depending on concentration

(Madhulatha, Anbalagan, Jayachandran, & Sakthivel, 2004). The results confirmed that a reduced BAP concentration of 0.25µM improved shoot elongation for both clones (GRD1 &

GRD3) and the new shoots of GRD3 appeared much more vigorous and healthier (more uniform shoot heights, leaf size and number of axillary shoots) in growth compared to most explant shoots before the experiment (variable shoot heights, leaf sizes and number of axillary shoots).

Shoot length can determine the efficiency of root formation, as longer shoots are in a better physiological condition for root induction (Lin, Bergann & Stomp, 1995; Thengane, Bhosle,

Deodhar, Pawar & Kulkarni, 2006; Xu, et al., 2008). The longer shoots were easier to transplant

and the uniform lengths facilitated better comparisons for all subsequent experiments.

Consequently, the protocol was altered for the BAP concentration and all shoot multiplication

media used the reduced BAP concentration of 0.25µM.

4.3 Changes to root induction media composition

The success of micropropagation largely depends on achieving root induction and then ex vitro

acclimatisation for all in vitro grown plantlets, but acclimatisation of rooted in vitro cultured

plantlets have many morphological limitations; e.g. unusual tissue organisation, incomplete

vascular connections between shoot and root resulting in low hydraulic conductivity and limited

transport of nutrients (Romano & Loucao, 2003). In vitro cultured plantlets have nutrients and water readily available and this may explain the anatomical and morphological differences in development of tissues; e.g. leaves have an undeveloped cuticle, lack of secondary xylem and phloem tissues that enhance conduction and minimal vascular differentiation in adventitious roots (Metivier, et al., 2007).

One of the most commonly used auxins for root induction includes IBA and the effect depends on the plant species, the concentration and the purpose of application (De Klerk, et al., 1999;

Ibrahim & Debergh, 2001). Different auxins, concentrations or combinations, may have to be applied depending on rooting treatments (in vitro or ex vitro) and on plant species (DeKlerk, et al., 1997). The application methods vary greatly, depending on the culture methods; for in vitro root induction the auxin is added to the culture medium, for ex vitro root induction the stems can be dipped in a highly concentrated auxin formula (Ibrahim & Debergh, 2001). The original root

induction protocol (McComb, et al., 1986), which required etiolation of shoots prior to root induction, produced inconsistent root induction and poor root formation with GRD1 clones.

4.3.1 Optimisation of duration of IBA pulsing

IBA is often the preferred auxin for root induction (Epstein & Muller, 1993), as plant tissues can rapidly oxidise other auxins such as IAA (DeKlerk, et al., 1997; Hausmann, 2003). Plants utilise

different auxins at different uptake rates (Nissen & Sutter, 1990; DeKlerk, et al., 1997; Simon &

Petrasek, 2011), which could explain the variation of root numbers formed (Figure 3.3). Plants

are generally not very sensitive to auxins during the first 24 hours, but after this time has lapsed

cells become more competent and begin to respond. Root initiation usually occurs between 72

and 96 hours as cells become activated, respond and result in root formation (De Klerk, et al.,

1999). A variation of sensitivity to auxins has been observed in clones of the same plant and this

can affect the transport of auxins to the plant cells and affect rooting response (Kim, et al.,

1998). Rooting response (number of primary roots) can depend on the duration of IBA pulsing

(Glocke, Delaporte, Collins & Sedgley, 2006) and the highest mean of root numbers was

recorded for 6 days of pulsing on IBA, and the mean number of roots (Figure 3.3) declined

rapidly thereafter. The results clearly showed that auxin pulsing is only necessary for a short

period of time; so all the subsequent root induction experiments used a 6-day IBA pulsing

duration.

The spontaneous root formation in GRD3 on shoot multiplication media occurred repeatedly

with some shoots, but always in isolated shoots of a culture vessel, and did not occur again when

the same shoot was sub-cultured. A possible explanation for this could be that plants contain the

72 natural, most abundant and endogenous auxin IAA. IAA is synthesised by the plant itself and in these particular shoots it may concentrate locally at the base of the shoot cutting causing cell enlargement, cell division (callus formation) and eventually root formation (Nordstrom, Jacobs

& Eliasson, 1991; Epstein & Muller, 1993; Hobbie, 1998). However, the results with GRD3 were very inconsistent, so all further experiments used only GRD1 clones.

4.3.2 IBA concentration

IBA can either promote the formation of adventitious roots, or it can inhibit this process; this variation depends on the IBA concentration and plant species (Muller, 2000). Variation in IBA concentrations can result in great variation of root response (Glocke, et al., 2006) and high IBA concentrations can increase the rooting rate (Xu, et al., 2008). These different rooting responses

(Figure 3.4) can be explained with the increased concentration of IBA allowing for faster action on the differentiating cells to become competent and enhance the cell division of the first root initials (Smith & Thorpe, 1975). It was observed that very high concentrations of IBA (160 µM

IBA in Figure 3.4) had an inhibitory effect on root formation (DeKlerk, et al., 1997; Wynne &

McDonald, 2002).

4.3.3 Auxin combinations for in vitro root induction

In contemporary plant propagation research it has become evident that endogenous plant auxins interact with exogenous auxins resulting in fast and slow responses (Woodward & Bartel, 2005).

Different auxin combinations have been successful for root induction in various woody plant

species, including medicinal woody plants, rose, apple and others (Marks & Simpson, 2000;

Martin, 2003; Martin, 2003/4; Pati, et al., 2006; Nandagopal & Kumari, 2007). Treatments

containing NAA (80µM, 54µM, 40µM and 26µM, Figure 3.5) produced large calluses (not

measured) at the shoot base and means of 0.76 – 1.73 roots/shoot. This could be because NAA

remains longer in the tissue as it is more persistent than IBA, hence the callus formation, which

may have blocked the root meristemoids from developing any further (De Klerk, et al., 1997).

IAA did not prove to be that effective for root induction; treatments with 80µM and 40µM IAA

(Figure 3.5) only produced a mean of less than 1 root/shoot. A possible explanation for this occurring could be the fact IAA is an endogeneous auxin synthesised by plants and degrades rapidly, so that only very small amounts of free auxin can be taken up by the target cells. IBA,

IAA (endogeneous auxins) and NAA (synthetic auxin) are conjugates where free auxin can be

released for preferential uptake (Van der Krieken, Breteler, Visser, Mavrdour, 1993). The

endogeneous auxins are less stable and IAA photo-oxidises more rapidly than IBA; IAA oxidises

by 50% and IBA by 10% in 24 hours, whereas NAA is a very stable, synthetic auxin that does

not oxidise (Nissen & Sutter, 1990; Epstein & Muller, 1993; Hausman, 2003; Simon & Petrasek,

2011). Treatments containing 40µM IBA combined with 40µM of IAA or NAA (Figure 3.5)

produced means of 3.43 and 3.85 roots/shoot. These results indicated that IBA was the more

active auxin for root induction and formation. IBA seemed to have a superior effect on root

induction and formation, which could be due to the preferential uptake and metabolisation of

IBA by the shoot tissues (Muller, 2000), but this was not further tested. The high concentration

of exogenous IBA applied to the wounded shoots for 6 days is synthesised by the receptor cells

targeted; initially wounding related compounds and auxin activate the cells at the cut site. This

is followed by cell division (after 24 hours) and primordial growth can begin to develop from

meristemoids after 96 hours with outgrowth from the stem occurring after 120 hours (De Klerk,

et al., 1999). This could explain the observed results as only the control treatment was

significantly different from the other treatments, and consequently all further experiments for

root induction only used 80 µM of IBA.

4.4 Different substrates

Roots are vital to any plant as they serve a multitude of functions; anchorage and support, as

well as being the supply channel for water and nutrients to the shoot. Tissue culture techniques

have utilised the universal application of gelling agents as the most favourable anchorage and

support system for the plant cultures. Gelling agents have long been the preferred substrate for

root induction and formation, but the limitations of agar and gelrite gelling agents for root

formation have also been investigated (DeBergh, 1983; Williams & Taji, 1987; Kumar, et al.,

2003; Peira-Netto, Petkowicz, Cruz-Silva, Gazzoni, Mello & Silveira, 2007). Root induction in

porous substrates has also been trialed as porous substrates are thought to alleviate some of the

physiological deficiencies that limit acclimatisation; one of the limitations in the physiology of in

vitro cultured plants is that the roots do not function properly due to reduced numbers or lack of

root hairs (Jay-Allemand, Capelli & Cornu, 1992; Lin, et al., 1995; Gangopadhyay, Das, Mitra,

Poddar, Modak & Mukherjee, 2002; Mohan, Soccol, Quoirin & Panday, 2004; Gangpadhyay,

Bandypadhyay, Gangopadhyay & Mukherjee, 2004 & Gangopadhyay, Roy & Mukherjee, 2009).

Porous substrates could overcome these short- comings and increase the survival rate during

acclimatization (Barrett-Lenard & Dracup, 1988; Rossetto, Dixon & Bunn, 1992; Hoarau &

Cornic, 1998; Zobayed, Zobayed, Kubota, Kozai & Kasgawa, 2000; Gangopadhyay, et al., 2004;

Gangopadhyay, et al., 2009).

Most roots grown in black sand, sterilised soil, rock wool, soil jiffies and coarse white sand (Fig

3.6) were positively gravitropic; the downward growth of the roots resulted from the additional rooting space that these substrates provided (4-7cms depth depending on substrate) compared to

~2cm depth that agar provided. Surprisingly, root formation was not statistically different in the

2 white sands, sterile soil and soil jiffies – it was expected that root formation/numbers in sterile soil and soil jiffies would be greater in these more aerated substrates.

Generally, roots can sense their environment (air, humidity gradients, nutrients, substrate and

water) that they touch and respond physiologically with growth accordingly; this response is

known as thigmotropism (touch), but the exact mechanism is not yet fully identified (Perrin,

Young, Murthy, Harrison, Wang, Will, & Masson, 2005; Gilroy, 2008; Gangopadhya, et al.,

2009). Experimentation with different substrates clearly showed different growth responses

and results from this experiment were not conclusive; e.g. destruction of roots when separating

(rock wool and soil jiffies), thus results may be incorrect as soil jiffies are designed for direct

transfer into soil where it biodegrades and rock wool is commonly used as a substrate for

hydroponic cultures. For in vitro root induction experiments these were too cumbersome and

probably did not provide adequate rooting space for 5 shoots. The different root growth

responses were evident in the relatively poor results for both white sands compared to black sand

and lower means for root numbers in sterile soil compared to agar and black sand (Fig 3.6).

4.4.1 Comparison of Agar, black sand and IVS

Root physiology, length and surface area/distribution (root architecture) are vital parameters for plant growth and survival, but this is difficult to quantify with living plants (Lynch, 1995; Davies

& Jacobs, 2005; Jacobs, Salifu & Seifert, 2005). Root numbers alone, where short and long roots are counted as equal, are not an ideal parameter when comparing plant root systems.

Therefore it is better to measure more than just one parameter in order to get more comprehensive data on root systems by analysing the means of root length and surface area, as well as shoot and root biomass for differences in substrate affects (Melhuish, 1968). The alternative in vitro rooting substrate being researched by Newell (et al., 2005) in vitro – soil medium (IVS) and other porous substrates (Barrett-Lenard & Dracup, 1988; Rossetto, et al.,

1992; Zobayed, et al., 2000; Gangopadhyay, et al., 2002; Mohan, et al., 2004; Gangpadhyay, et al., 2004) are aerobic and less dense than solidified agar media.

Root formation in the alternative substrates of black sand and sterile soil (Figs 3.7 & 3.8) was compared with agar-solidified media. The results showed the improved root performance (root surface area and length) observed for sterile soil, which was statistically different to the other treatments (Figure 3.7). Sterile soil is not a substrate that is uniform in density or particle size

(it consisted of peat and perlite 1:3), so different root sizes can occupy the substrate pores, which could explain the increased root numbers, root length and surface area (Pierret, Moran &

Doussan, 2005). This could be due to the different mechanical resistance of the substrates and the stimulation that this provides to the root and could explain the variation in root morphology of shoots grown in sterile soil (Gangopadhya, et al., 2009). It is hypothesised that medium

hypoxia and the build-up of ethylene and CO2 observed in agar substrates can limit root initiation

and elongation. Using a porous substrate could reduce these effects, as the porous media could

allow for greater diffusion of inhibiting effects of ethylene and CO2 (Newell, et al., 2003).

However, most of these root growth patterns observed are the short-term effects in response to physiological stimuli and cannot be regarded as root growth patterns, but rather as spontaneous root growth in individual shoots (Grabov, Ashley, Rigas, Hatzopoulos, Dolan & Vicente-Agullo,

2005). This experiment showed that roots could display a high level of morphological plasticity in response to the different substrates that they were grown in (Figure 3.8).

4.5 Comparison of sterile soil with different nutrients

The previous experiment achieved good root induction on GRD1 clones using an aerated, porous substrate of sterile soil instead of an agar medium. The limitations of agar-based media have been discussed (Newell, 2005; Gangopadhya, et al., 2009) and as a result, some research has been done on ex vitro rooting of in vitro cultured plantlets (Kim, et al., 1998; Martin, 2003; Xu, et al., 2008; Gangopadhya, et al., 2009). The process of ex vitro root growth usually entails that the in vitro grown shoots receive exogenous auxin treatment before being transferred for acclimatisation and root growth to ex vitro conditions. This method is often applied as a cost and time reducing strategy (Martin, 2003; Thomas & Schiefelbein, 2005; Feyissa, et al., 2007). The experiment showed that root induction and growth was possible (Figure 3.9) without added nutrients in the substrate, as the pulsing on IBA media induces root formation. The immediate application of IBA after wounding did not seem to adversely affect root formation and the results from the sterile soil treatments with no nutrient supplements were comparable (no statistical

78 difference) to the other treatments for mean root length and numbers (Fig 3.7). This suggested that roots could be induced with a high concentration of IBA pulsing (in vitro) and these results were comparable to results from experiments by De Klerk, et al., (1999) where cuttings were exposed to very high auxin concentrations ex vitro before being planted out in to soil directly.

For the verticordia shoots in this experiment the adventitious roots formed spontaneously as a direct result of exogenous auxin application. There was no statistical difference in root length or surface area between treatments with just DDI and DDI with sucrose. It has been suggested that starch accumulation is a prerequisite for adventitious tissue regeneration and root formation

(Jasik & De Klerk, 1997). Sucrose from the media can be synthesised to starch and has a regulatory effect on adventitious organ formation; starch grains accumulate in the target cells that then produce adventitious roots, but then degrade after cell division has occurred

(Capellades, Lemeus & de Bergh, 1997; De Klerk, et al., 1999, Hazarika, 2003). However, in this study the sucrose concentration was not apparent to have an effect on adventitious root formation by increasing sensitivity to IBA (Calamar & De Klerk, 2002), as the shoots in the treatment without sucrose had comparable root formation. The IBA concentration of the pulsing media was high enough for free auxin to reach the target cells and to stimulate rooting response

(Figure 3.9). Root induction and growth occurred in sterile soil without nutrients and a carbon source, so this suggested that the in vitro root induction process could be eliminated to save time and costs.

4.6 Ex vitro root formation and acclimatization

For acclimatisation from in vitro to ex vitro to be successful physiological changes to correct the abnormalities in leaf morphology, photosynthetic capacity and water relations must occur.

Improved physiological functions of in vitro plantlets should facilitate rapid acclimatisation to ex vitro conditions and thereby increase plantlet survival rates (Rossetto, et al., 1992, Bonal &

Monteuuis, 1997; Hazarika, 2003; Martin, 2003; Feyissa, et al., 2007). Thomas & Schiefelbein

(2005) compared in vitro with ex vitro physiological rooting responses and examined if there were any significant differences in gene expression pattern. They discovered that gene expression was identical between in vitro and ex vitro root induction.

For both treatments soil (in vitro = sterile soil) was used (Figures 3.10), which is more aerated than an agar medium, and this had positive effects on root induction and development, as all stages of rooting in plants require oxygen (Newell, 2006). Medium hypoxia is assumed by

Newell (2006) to occur in agar rooting medium and this could limit not only root induction but also root elongation, as compared to the root growth patterns observed in IVS (Newell, et al.,

2005). Another advantage of using a sterile soil media is that it has better water holding capacity over agar media and therefore can prevent medium hypoxia. In this experiment the biomass of shoots and roots for in vitro and ex vitro soil had no statistical difference (Fig 3.10), and neither visible shoot growth nor wilting was observed for the duration of the experiment.

These results indicated that possibly the increased aeration of a soil media could allow for greater root growth, because aerated media allows for greater diffusion of ethylene caused by in vitro pulsing with auxins, which enhances ethylene synthesis (Newell, et al., 2005). Agar medium does not readily allow for rapid gas diffusion, hence the accumulation of ethylene at the basal part of the stem. This accumulation can inhibit root induction, growth and elongation (De

Klerk, 2002). Ex vitro rooting would eliminate the process of transfer and reduce the risk of root destruction (Gangopadhya, et al., 2009).

Combined in vitro shoot multiplication with ex vitro root induction and acclimatisation has been trialled to enhance survival of plantlets, as well as a cost and time reducing strategy (McClelland, et al., 1990; Kim, et al., 1998; Martin, 2003; Thomas & Schiefelbein, 2005; Feyissa, et al., 2007;

Xu, et al., 2008). Newell (2006) alleged that agar medium can suppress rooting performance in many Australian plant species, so the rooting potential of these are difficult to be accurately determined. He concluded that survival rates dramatically increased when the root induction phase for many Australian woody plant species occurred ex vitro and simultaneously with acclimatisation on IVS without de-flasking and transferring, when compared to the conventional root induction on in vitro agar medium followed by transfer to ex vitro substrates and conditions.

For this experiment the in vitro shoots grown in sterile soil were re-potted into crackpots at the beginning of this experiment. The positive results (Figure 3.10) demonstrated that it was possible to multiply shoots in vitro and then to root and acclimatise these shoots ex vitro simultaneously.

This process of combining Stage 3 and 4 of micropropagation excludes disturbing and breaking roots during transfer, using soil in aseptic laboratory conditions can be avoided and it is a time and money saving strategy. Ultimately the effectiveness and success of this relies on the survival of the plantlets (Martin, 2003).

4.7 Survival

Trials with ex vitro rooting and simultaneous acclimatisation have been considered promising for the micropropagation of many different plant species. This is due to the reduction of

establishment and production time, labour and associated costs, as ex vitro rooting can reduce the overall costs by 35% (McClelland, et al.; Kim, et al., 1998; Martin, 2003; Thomas &

Schiefelbein, 2005; Feyissa, et al., 2007; Xu, et al., 2008). The appropriate substrate, with high water holding capacity, can be beneficial to acclimatisation as well as the reduction of light intensity in the greenhouse by using shade cloth (Viegas, Pereira, Ambrosano & Batista, 2005).

The results (Fig 3.11) suggest that ex vitro root growth can occur simultaneously with acclimatisation as a time and cost saving strategy. The high long-term survival rate (Figure

3.11) could possibly be due to several factors; e.g. possible in vitro rejuvenation (Haapala, et al.,

1 2004; Andreu & Marín, 2005) of plantlets due to the long time in culture (more than 2 /2 years)

with constant sub-culturing, endo- and exogenous auxin levels (Ibrahim & Debergh, 2001) and favourable environmental conditions (DeBergh, 1991; Bonal & Monteuuis, 1997; Hazarika,

2003). The beneficial environmental ex vitro conditions included the medium substrate that

possibly allowed for flushing of excess auxin, better gas exchange and water retention/drainage,

additional to relative humidity and the quality/quantity of light (Gangopadhya, et al., 2009).

The wilting and death (Figures 3.11 & 3.12, Table 3.2) of some plants was probably due to the

combination of different factors during ex vitro acclimatisation; initially the lack of ventilation in the humidity tent (even though the crack pot trays had vented lids) and increase in ambient temperature due to the lack of ventilation. It could have been possible that these shoots could not adequately control plant water loss due to the limited stomata function (Gilly, Rohr & Chamel,

1997) combined with ambient humidity and temperature fluctuations. This may explain why shoots did not fully recover, acclimatise or survive in the first 56 days for the first trial.

Over time stomatal performance must have improved in the remaining plantlets, as these acclimatised and could control transpiration (this is only evident by survival and was not tested

further), even though 100% survival was not achieved. Transpiration control and stomata

function can usually improve between 8-12 weeks (Figs 3.11 & 3.12, Table 3.2). The

autotrophic growth of new shoots and/or shoot elongation further confirmed this. The

functioning roots also would have contributed to the water retention capacity as well as nutrient

uptake required for further growth (Diaz-Perez, Sutter & Shackel, 1995).

The long-term survival of over 70% of in vitro cultured plantlets with ex vitro induced roots

(Figs 3.11, Table 3.2) indicated that the abnormalities and limitations in shoot and root

morphology and physiology were gradually overcome and these became functional during ex

vitro acclimatisation (Pospislova, et al., 1999). In comparison the in vitro root induced shoots

(Figure 3.11) did not perform as well, which could have been due to the additional stress of

transplanting, due to root damage that occurred during this process, and sudden changes of

ambient conditions (Hazarika, 2003). However, high mortality rates after the conclusion of this experiment suggest other factors negatively impacting on long-term survival, but due to time constraints this was not further investigated.

4.8 Effectiveness of improved protocols on new clones

One of the aims of this research was to test the effectiveness of the improved shoot and root protocols on other verticordia clones that were introduced into tissue culture. Not all new clones were successful for micropropagation (Chapter 3.8.1), but those that could be continuously subculture were trialled for ex vitro root formation combined with simultaneous acclimatisation

(Chapter 3.8.2) and monitored for long- term survival (Figure 3.12) for comparison to GRD1.

4.8.1 Survival of new clones

The success of ex vitro root induction for most new clones was confirmed by the fact that all

plantlets had roots. Initial acclimatisation was also a success, but long-term survival was less successful (Figure 3.12). The stomatal function may have developed over time (8-12 weeks) in

many plantlets, so the plantlets could control transpiration and water loss with the reduced humidity (Fila, et. al., 1998; Pospisilova, et al., 1999), but functional roots were needed for

nutrient uptake to facilitate growth and survival (Mulataya, et al., 2002; Hazarika, 2003), however, evidence of autotrophic growth was insufficient. The poor long-term survival rates suggest that not all plantlets repaired their in vitro functional abnormalities, in particular, the lack of junction between the stem vascular tissue and the adventitious roots, as they failed to thrive and grow. The lack of functional roots (this was not tested, as plantlets had roots) could explain why these results do not compare to the results of acclimatisation and survival of GRD1 clones

(Chapter 3.7). This may also be due to the genotypic and phenotypic variation that is a common occurrence observed in micropropagated plants (e.g. different internodes in GF clones, different leaf sizes in all clones), and these variations can pre-exist in the ex-plant material or result from in vitro conditions (Perez, et al., 2011). There was inconclusive evidence with regards to in vitro rejuvenation (Haapala, et al., 2004; Andreu & Marín, 2005) having occurred, even though most plantlets had root formation. The clones GF 1-4 originated from new shoot (possibly juvenile) growth (after fire destroyed the explant) and performed similarly to the clones taken from seasonal, fresh growth (Table 3.1), which is usually mature vegetative growth (Hackett &

Murray, 1992).

5. CONCLUSION This study confirmed that the improved shoot multiplication and root induction protocols produced overall good results with most V. grandis clones, but the best results were achieved with a single clone (GRD1) that had been in culture for over 5 years. It was possible to introduce new V. grandis clones into tissue culture, even though the success was limited for the other verticordia species; V. ovalifolia, V. fragrans and V. densiflora. The new shoot multiplication

protocol attested to the effectiveness with good results for the new V. grandis clones.

The improved shoot multiplication protocol produced more uniform elongation for most clones,

which was necessary for further experimentation; most new clones performed well on the lower

concentration of 0.25 µM BAP, but elongation was greatly improved for the GRD3 clone. This

resulted in having an endless supply of shoots from most clones that had more uniform lengths

needed for further experimentation, as this allowed for better comparisons (Lin, et al., 1995).

The optimal auxin for root induction was IBA and the best concentration was determined to be

high at 80 µM, and the optimal in vitro auxin pulsing duration was found to be 6 days. This

optimised root induction protocol worked well for most GRD clones. Recent studies by Newell

(2006) hypothesized that Australian native plants perform better in an aerated medium compared

to agar media. Experiments with different substrates, other than a gelled medium, were trialled

on GRD1 and a soil mixture of peat and perlite (1:3) was found to support this with good results.

Other studies (Kim, et al., 1998; Hazarika, 2003) have suggested that species, which are

problematic with in vitro root induction, often improve root formation when combined with ex

vitro acclimatisation. The effectiveness of combined root formation and growth, after in vitro

IBA pulsing, and acclimatisation ex vitro was verified by high percentages of root formation and

subsequent growth and survival. This process saved not only time, but eliminated the excessive

handling of delicate shoots with roots, prevented breaking roots whilst de-flasking from in vitro

media and transfer to ex vitro growth media (soil), and preparing soil in an aseptic laboratory was avoided as well. Simultaneous ex vitro root formation, after IBA pulsing, and

acclimatisation was achieved and confirmed by some of the best survival rates in most

experiments for Verticordia grandis, especially the GRD1 clone. The alternative trials with ex

vitro auxin dipping (instead of in vitro auxin pulsing) prior to out-planting in the greenhouse for

acclimatisation were not successful and resulted in 100% mortality of shoots. An illustration of

how this approach is incorporated into a more standard micropropagation approach is illustrated

in Figure 4.13 where the elimination of the need for roots prior to transfer to soil is illustrated.

Long-term survival was monitored for 252 days, which included re-potting plants after 112 days

into larger containers and a peat – perlite – sand (1:1:1) mixture for added drainage. Plants that

had roots induced in vitro and that were then transferred for acclimatisation to the greenhouse

after 28 days had a high mortality rate (possible due to roots being damaged during transfer) and

after 140 days there was 0% survival. In contrast, all ex vitro root grown and acclimatised

plants had a high survival rate at over 72% after 252 days, when the experiments concluded.

Figure 4.13: The incorporation of ex vitro rooting process into the micropropagation process - adapted from George et al., 2007.

However, survival continued to decline after experiments had concluded at 252 days, and after more than 2 years there were no plants left. This indicates that more research into enhancing long-term plant survival and field performance would be required if commercial production for

Verticordia grandis was to be pursued.

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