MICROPROPAGATION OF WASABI (WASABZA JAPONZCA) AND

IDENTIFICATION OF PATHOGENS AFFECTING PLANT

GROWTH AND QUALITY

Georgina Rodriguez B.Sc., Simon Fraser University, 2000

THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS

FOR THE DEGREE OF

Master of Science

in the Department

of

Biological Sciences

O Georgina Rodriguez

SIMON FRASER UNIVERSITY

Fall 2007

All rights reserved. This work may not be reproduced

in whole or in part, by photocopy or other means,

without permission of the author.

Name:

Degree:

Title of Thesis:

APPROVAL

Georgina Rodriguez

Master of Science

Micropropagation of wasabi (Wasabia japonica) and identification of pathogens affecting plant growth and quality

Examining Committee:

Chair: Dr. I. Novales Flamarique, Assistant Professor

Dr. Z.K. Punja, Professor, Senior Supervisor Department of Biological Sciences, S.F.U.

Dr. A.R. Kermode, Professor Department of Biological Sciences, S.F.U.

Dr. A.L. Plant, Associate Professor Department of Biological Sciences, S.F.U.

Dr. A. Uvesque, Section Head Biodiversity (Mycology and Botany), Agriculture and Agri-Food Canada Public Examiner

24 October 2007 Date Approved

ii

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Revised: Fall 2007

ABSTRACT

Wasabi (Wasabia japonica Matsum.) is grown for its rhizome which is a culinary

ingredient. Wasabi plantlets were regenerated from meristem-tips taken from axillary

buds on greenhouse-grown plants. Axillary shoots developed at 3.8*0.13 per meristem-

tip every 8 weeks on 0.8 mg/L N6-benqladenine (BA) and rooted on hormone-free

Gamborg's B5. These plants were used in subsequent pathogenicity studies. Pythium

dissotocum and Pythium intermedium were recovered from plants with root loss.

Hydroponically-grown plants were inoculated with mycelial plugs of each Pythium spp.

and after 2 months, root dry weight was 16% of the controls. Isolation studies from

blackened rhizome vascular tissue were also conducted. Microscopic observations and

isolation data showed that bacteria were associated with symptomatic plants.

Pectobacterium carotovorum subspecies carotovorum (Pcc) was identified. Symptoms

developed when roots were wounded and inoculated with Pcc. Pcc was recovered from

the rooting medium used by growers and found to be identical to the wasabi isolate.

Keywords: histopathology, hydroponic culture, Pectobacterium carotovorum, Pythium

dissotocum, Pythium intermedium, root loss, sterilization, subculture, tissue culture,

vascular blackening, Wasabia japonica, wounding

ACKNOWLEDGEMENTS

I would like to thank Brian Oates of Pacific Coast Wasabi Ltd., Vancouver, B.C.,

for generously providing wasabi plants and funding for this research. Without your

contribution this would not have been possible. I am grateful to Dr. T. Barasubiye and

Dr. C. A. Levesque of the National Fungal Identification Service, Agriculture and Agri-

Food Canada, Ottawa, Ontario, for assisting in the identification of the two Pythium

species. Also thanks to Len Ward and Dr. Solke de Boer of the Canadian Food

Inspection Agency, Charlottetown, P.E.I., for supplying me with Pectobacterium

carotovorum strains. Many thanks goes to Terry Holmes at Pacific Forestry Research

Centre, Victoria, B.C., who prepared and photographed the wonderful images of wasabi

and bacteria.

I am very fortunate to have worked under the guidance of Dr. Punja, and am proud

to have studied in his laboratory. Thank-you to my committee, who provided support

throughout this experience. And many thanks to all of my lab' colleagues, you were a

source of support, inspiration and a lot of fun.

Finally, I am indebted to my family, whose commitment to my studies was

invaluable. Thank-you Guillermo for all that you sacrificed, Mom and Dad for giving me

the opportunity to pursue a great education, and my amazing children for your contagious

TABLE OF CONTENTS

. . ............................................................................................................................ Approval 11

... Abstract .......................................................................................................................... 111

.......................................................................................................... Acknowledgements iv

Table of Contents ............................................................................................................... v ... List of Figures ............................................................................................................. VIII

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

Chapter 1: General Introduction ..................................................................................... 1 1.1. Wasabi biology .......................................................................................................... 1

..................................................................................................... 1.2. Wasabi production 2 ............................................................................................................ 1.3. Tissue culture 6

1.3.1. Meristem tip micropropagation and axillary shoot propagation ........................ 6 ............................................................................... 1.3.2. Micropropagation of wasabi 7

................................................................................................. 1.4. Pathogens of wasabi -8 ................................................................................................ 1.5. Tissue discolouration -9

1.5.1. Stem blackening in wasabi and Phoma wasabiae .............................................. 9 ..................................................... 1 S.2. Tissue discolouration in other plant species 11

......................................................................................... 1 S.3. Phenolic compounds 12 ................................................................................................................... 1.6. Pythium -14

............................................................................ 1.6.1. Taxonomy and identification 14 1.6.2. Pythium spp . as pathogens ................................................................................ 15

.......................................................................................... 1.6.3. Pythium dissotocum 17 ........................................................................................ 1.6.4. Pythium intermedium 1 8

............................................................. 1.6.5. Control methods for Pythium diseases 19 ....................................... 1.7. Pectobacterium carotovorum (syn . Erwinia carotovora) 23

....................................................................................................... 1.7.1 . Taxonomy 23 ........ . 1.7.2. Pectobacterium carotovorum subsp . wasabiae and subsp carotovorum 24

1.7.3. Soft-rot disease caused by Pcc ......................................................................... 25 1.7.4. Control methods for soft-rot diseases ............................................................... 26

................................................................................................. 1.8. Research objectives 27

Chapter 2: Micropropagation of Wasabi (Wasabia japonica) from Meristem Tips and Axillary Shoots ................................................................................................. 29

............................................................................................................. 2.1. Introduction 29 .......................................................................................... 2.2. Materials and methods 31

......................................................... 2.2.1 . Plant material and meristem sterilization 31

................................................................................................. 2.2.2. Shoot initiation 32 ............................................................................................ 2.2.3. Shoot proliferation 35 ............................................................................................. 2.2.4. Root development 35 ............................................................................................. 2.3. Results and discussion 36

....................................................................................... . 2.3.1 Meristem sterilization 36 ............................................................................................ 2.3.2. Shoot proliferation 36 ............................................................................................ 2.3.3. Root development -38

............. Chapter 3: Root infection of wasabi (Wasabia japonica) by Pythium species 40

3.1. Introduction ............................................................................................................. 40 3.2. Materials and methods ............................................................................................ 44

3.2.1. Isolation and identification of fungi ................................................................. 44 .................................................................................................... 3.2.2. Plant material 44

................................................ 3.2.3. Inoculation and root colonization experiments 45 3.3. Results and Discussion ............................................................................................ 46

3.3.1. Isolation and identification of fungi ................................................................. 46 3.3.2. Inoculation of wasabi plants and assessing root infection .............................. 46

Chapter 4: Rhizome blackening of wasabi caused by Pectobacterium .................................................................................... carotovorum subsp . carotovorum 51

............................................................................................................. 4.1. Introduction 51 4.2. Materials and methods ............................................................................................ 53

4.2.1. Microbial isolation and microscropy ................................................................ 53 4.2.2. Plant material for pathogenicity tests ............................................................... 55 4.1.4. Bacterial identification ..................................................................................... 58

....................................................................................... 4.2.5. Bacterial inoculations 60 .................................................................... 4.2.6. Bacterial plus fungal inoculations 61 ................................................................... 4.2.7. Identification of inoculum sources 61

..................................................................................................................... 4.3. Results 62 ................................................................ . 4.3.1 Microbial isolation and microscopy -62

.......................................................................................... 4.3.2. Fungal inoculations 62 ........................................ 4.3.3. Bacterial identification

..................................... 4.3.4. Bacterial inoculations 9 .................................................................... 4.3.5. Bacterial plus fungal inoculations 64 ..................................................................... 4.3.6. Identification of inoculum source 65

............................................................................................................... 4.4. Discussion 65

Chapter 5: General discussion and conclusions ............................................................ 70 .......................................................................................... 5.1. Tissue culture of wasabi 70

...................................................................................... 5.1.1. Pathogen-free material 70 ................................................................................................. 5.1.2. Acclimatisation 70 . . ............................................................................................ 5.1.3. Commerclallsation 71

5.2. Low root mass in wasabi .......................... .. ........................................................ 73 ....................................................................................... 5.2.1 . Pythium spp . control 73

5.3. Rhizome vascular blackening .................................................................................. 74 5.3.1. Pcc isolation variation and the possible involvement of additional

microbes ........................................................................................................... 74 ............................................................. 5.3.2. Defining symptoms and causal agents 76

5.4. The interaction between Pcc and wasabi ................................................................ 78 ............................................................................................................. 5.5. Conclusions 80

......................................................................................................................... References 81

vii

LIST OF FIGURES

Figure 2.1. Tissue culture and micropropogation of wasabi ( Wasabia japonica cv. Daruma). ................................................................................................. -33

Figure 3.1. The effect of Pythium dissotocum and Pythium intermedium on wasabi roots. .................................................................................................. 42

Figure 3.2. Root dry weights from 8-month-old wasabi plants inoculated with Pythium spp. .................................................................................................. 48

Figure 4.1. Wasabi rhizomes with vascular blackening. ................................................ S 4

Figure 4.2. Reproduction of wasabi vascular blackening symptoms ............................... 57

Figure 4.3 Identification of Pectobacterium carotovorum subspecies carotovorum using primer set Llr and G l f. .................................................. 66

LIST OF TABLES

Table 1.1. Biocontrol agents available for controlling diseases caused by ................................................................................................. Pythium spp. .2 1

Table 1.2. Examples of fungicides available for controlling disease caused by .................................................................................................. Pythium spp .22

Table 2.1. The effects of different cytokinins (kinetin and BA) on shoot proliferation on Gamborg's B5 with 2.2 g/L phytagel .................................. 37

Table 4.1. Reference strains of Pectobacterium carotovorum ssp carotovorum (Pcc) and P. carotovorum subspecies wasabiae (Pcw) used in this study ............................................................................................................. S 9

Table 4.2. Incidence of rhizome vascular blackening (rvb) and soft-rot (rot) in wasabi plants that were inoculated with Pcc and grown in potting mix at different temperatures ......................................................................... 65

CHAPTER 1: GENERAL INTRODUCTION

1.1. Wasabi biology

Wasabi ( Wasabia japonica Matsumura, syn. Eutrema japonica) is a perennial

herb that prefers cool, moist growing conditions (Hodge, 1974). It is a member of

Cruciferaceae or Brassicaceae family, but has a unique morphology; its petioles originate

from a vertical rhizome and can extend up to 50 cm in height on a mature plant. The

heart-shaped, simple, palmate-veined, evergreen leaves can reach a diameter of 25 cm in

a mature plant, and as the rhizome slowly grows, the lower leaves senesce and fall off

(Hodge, 1974; Follet, 1986; Chadwick, 1993). Axillary buds develop on the rhizome,

which can give rise to a smaller rhizome attached to the main rhizome. The main

rhizome can grow to 5 - 40 cm in length and 2 - 5 cm in diameter over a 15-month to 3-

year period (Hodge, 1974; Chadwick, 1993). At harvest, the rhizome is cylindrical to

conical in shape, slightly curved, covered with leaf scars, and pointed at the terminal bud

(Chadwick, 1993). In spring, flowers are produced from the crown of the plant and

peduncles can reach 2 m in height. Racemic inflourescence can elongate and form

secondary inflorescences (Chadwick, 1993). The flowers are white and arranged on

racemes (Hodge, 1974). Seeds are borne in siliques along the length of the peduncle and

contain up to 8 seeds. Each seed is 2 to 3 mm long and 1 mm wide, covered by a thin

brown seed coat (Chadwick, 1993).

Wasabi is grown primarily for its valuable rhizome, which is freshly-ground using

a shark skin or a very fine-toothed ceramic grater and eaten as a condiment in Japanese

cuisine. More commonly used wasabi pastes and powders contain little or no true

wasabi. Instead, the primary ingredient is horseradish (Amoracia rusticana Gaertu.) or

western wasabi, chinese mustard (Brassica juncea L.) with food colouring. Other parts

of the wasabi plant can also be eaten, including the leaves and petioles, which can be

pickled and are often used in processing with lower grade rhizomes and added to wasabi-

flavoured products (Chadwick, 1993).

Freshly-ground wasabi has a pungent flavour, due to the large number of myrosin

cells in the rhizome (Kimura et al., 1990). Myrosin cells contain water-soluble

glucosinolates and myrosinase enzymes, which are compartmentalised into the vacuole

and various cytoplasmic membranes, respectively (Koide and Schreiner, 1992). When a

rhizome is ground, the myrosin cells break and myrosinase hydrolyses the glucosinolate,

known as sinigrin (CIOH16KN09S2), forming various isothiocyanates (Jordt et al., 2004;

Hodge, 1974). Research has shown that isothiocyanates have many beneficial medicinal

properties, including biocidal and anticarcinogenic properties (Chadwick, 1993).

1.2. Wasabi production

Wasabi is native to Japan where it can be found growing along river banks and in

shaded mountain streams (Follet, 1986; 1987; Chadwick, 1993). Cultivation in Japan

dates back to the loth century and wasabi was introduced to Taiwan in 1914, when it was

still a colony of Japan (Lo C. T., 2000). The earliest report of cultivation outside of the

Orient was in 1903, when wasabi was introduced for trial in the United States (Fairchild,

1903). Trials were reportedly unsuccessful (Chadwick, 1993); however wasabi has since

been reintroduced as a minor crop. Today, wasabi is cultivated not only in Japan, but in

North Korea, New Zealand, Australia, Israel, Brazil, Thailand, Taiwan, USA, and Canada

(Follet, 1987; Palmer, 1990; Chadwick, 1993).

The surge in wasabi cultivation outside of Japan has been stimulated by the

increasing price for wasabi, which currently can fetch US$240 per kilogram for fresh

rhizomes. The price has largely been driven by an increase in demand, which has grown

exponentially in the last 35 years, and is exacerbated by limited production areas

(Chadwick, 1993). In Japan, only about 880 hectares are suitable for production of

wasabi, and these areas are being destroyed by roads, dams, and fertilizer and sediment

pollution from rice fields (Chadwick, 1993).

The limited area available for wasabi production is due to the specific conditions

required for wasabi growth. Wasabi prefers an air temperature of between 8 and 18 "C

and growth is halted at temperatures above 28 "C. Wasabi has a high requirement for

moisture, requiring high and constant rainfall throughout the year. High humidity in the

warmer summer months is beneficial since it prevents wilting. Wasabi does not grow

well in full light and leaves can burn in direct sunlight. In Japan, plants are shaded from

April to August using either evenly planted deciduous black alders (Alnus japonica) or 40

- 50% shadecloth (Follet, 1986).

Many cultivars of wasabi exist, and are named for their region of cultivation or

after the person who developed them (Chadwick, 1993). The two main cultivars grown

in Japan are "Midori" and "Daruma" (Follet, 1986). "Daruma" is also grown in Canada

as it can produce good-quality rhizomes suitable for the fresh market and it is regarded as

an adaptable cultivar able to tolerate slightly higher temperatures, although it has a low

tolerance to high light levels (Follet, 1986). Another cultivar grown in Canada is

"Mazuma" which produces a spicier rhizome.

The various wasabi cultivars can be grown in soil in or an aquatic system

resulting in one of two types of wasabi. "Oka" wasabi is produced by growing wasabi in

soil and yields a rhizome of lesser quality and value. Approximately 86 % of this wasabi

is used for processing and not eaten fresh (Follet, 1986), as the rhizome is less pungent

and can be light green to white in colour, yielding an aesthetically unpalatable paste when

ground. "Sawa" wasabi is produced by growing wasabi in an aquatic system, either in

streams or in artificial hydroponic or semi-hydroponic systems. This wasabi generally

yields a more pungent and green rhizome which is highly preferred for the fresh market

(Follet, 1986).

In Japan, only 65 % of wasabi grown in an aquatic environment is sold on the

fresh market due to strict quality requirements (Follet, 1986). The water quality of

aquatically-grown wasabi is critical to crop quality. Wasabi prefers a constant water

temperature of approximately 10 to 13 OC throughout the year. Although plants will grow

in water of higher temperature, rhizome growth can be compromised and plants become

more susceptible to disease (Hodge, 1974). Water must be in constant supply, free of

organic pollution, and contain a high level of dissolved oxygen (Follet, 1986). In Japan

where wasabi is grown in streams, the water originates from clear, cool, clean mountain

springs as opposed to runoff. Macro-nutrient content of the water may also be critical to

wasabi growth rate (Follet, 1986).

Wasabi can be harvested throughout the year, and cropping times range from 15

months to 3 years, depending on the growing environment, the propagation method used,

and the desired rhizome size (Hodge, 1974). Plants are harvested by hand and offshoots

are removed which can be later used for propagation (Follet, 1986). The rhizome is

trimmed of roots and leaves and then washed in preparation for the fresh market or

processing. The less valuable leaves and petioles may also be processed (Follet, 1986) or

eaten fresh in salads.

Growers obtain new planting material using three methods: vegetative propagation

using the vegetative offshoots obtained during harvest; using seeds which they produce

themselves; or acquiring plantlets obtained through tissue culture (Chadwick, 1993).

Traditionally, vegetative propagation is generally preferred as crop turnover is faster, less

labour-intensive (Follet, 1986), cheaper and according to some consumers, results in

rhizomes with better taste and texture (Chadwick, 1993). Up to 20 offshoots can be

obtained from a 2 year-old plant and can be planted during spring, summer, or autumn

(Chadwick, 1993). These offshoots contain a shoot meristem, a tiny rhizome, and

developed leaves and therefore establish relatively quickly, resulting in quicker crop

turnaround. Additionally, the offshoots are essentially free as they are obtained when

rhizomes are trimmed for sale. However, successive vegetative propagation reportedly

results in decreased growth rate and accumulation of pathogens (Follet, 1986), and

therefore it is recommended that seed propagation be used every third to fourth planting

(Adachi, 1987).

In Japan, seeds are often produced by the grower and harvested in summer. Seeds

are stratified and sown by hand into nursery beds in January and February. Beds are

regularly hand-weeded until September, when seedlings are selected, washed, trimmed

and planted into the wasabi beds (Follet, 1987). The increased labour cost associated

with obtaining growing material from seed is compounded by low germination rates and

high seedling mortality due to damping-off pathogens.

1.3. Tissue culture

1.3.1. Meristem tip micropropagation and axillary shoot propagation

Plants obtained from micropropagation are used as a source of disease-free plants

and eliminates increased cost associated with seed germination, and decreased growth

rate associated with vegetative propagation. Micropropagation refers to a very small

propagule that is used in culture in order to asexually propagate a plant. When the

propagule used is a meristem tip, the method is referred to as meristem tip

micropropagation (Hu and Wang, 1983). The excised meristem tip used in

micropropagation is comprised of the apical meristematic dome and three to five leaf

primordia, and excludes any differentiated provascular or vascular tissues (Pollard and

Walker, 1990).

The in-vitro shoot or plantlet that is derived from the meristem tip culture may be

induced to produce secondary shoots or axillary shoots. These axillary shoots may be

separated and placed on fresh culture medium in order to produce additional shoots and

each shoot can be rooted to produce a plantlet. This technique, known as axillary shoot

propagation, allows for a high propagation rate of the original meristem, which is

essential to commercial plant propagation (Pollard and Walker, 1990).

In the early 1960s, Morel pioneered micropropagation in order to clone the

Cymbidium orchid (Morel, 1960b), and the method gained popularity in the 1970s (Hu

and Wang, 1983). In 1974, Murashige developed the concept of different developmental

stages involved in micropropagation and axillary shoot propagation: Stage I: explant

establishment in aseptic culture, Stage 11: multiplication of the propagule, which can be

achieved by axillary shoot propagation, and Stage 111: rooting and hardening for

transplanting (Murashige, 1974). Stage I11 often occurs as two separate stages, and thus

is sometimes divided into Stages I11 and IV. However, in order to decrease production

costs, unrooted shoots can be removed from in-vitro culture so that rooting and hardening

can be achieved concurrently. Another cost-saving strategy that is desirable in

commercial settings is to combine multiplication of shoots and rooting; however, this is

not possible for all species (Pollard and Walker, 1990).

Micropropagation is widely used to propagate disease-free plants from infected

stock. It has been used following chemo- and/or thermo-therapy, or by itself in order to

produce virus-free plants in numerous species. Similarly, planting stock that is

systemically infected with mycoplasma, fungi, or bacteria can be rendered disease-free

by means of micropropagation (Hu and Wang, 1983).

1.3.2. Micropropagation of wasabi

Previous tissue culture studies on wasabi have focused on development of somatic

embryos and micropropagation from zygotic embryos (Eun, 1995; 1996; 1999). Few

published studies have used apical meristems (Eun et al., 1997) and axillary buds (Hung

et al., 2006; Hosokawa et al., 1999) for micropropagation. These studies did not report

the cultivar(s) or the sterilization method used. Axillary bud culture reported by

Hosokawa (1999) did not describe sterilization protocols, which is critical in order to

successfully establish explants from plants that have been grown under nonsterile

conditions, such as the greenhouse. In 2006, a publication on axillary bud culture

outlined a protocol for explant establishment and micropropagation. This study also

relied on the use of antibiotics to establish explants (Hung et al., 2006). This would have

been insufficient for the use of tissue culture plants in pathology experiments, as plants

were required to be free of microbes.

1.4. Pathogens of wasabi

Like other crucifers, wasabi is a host to a wide range of pests and diseases, which

can result in up to 100 % crop loss. Many viral, bacterial, and fungal pathogens have

been reported on wasabi, with most reports originating from Japan and Taiwan.

Viruses are reported to cause crop loss in Japan, Taiwan, and New Zealand.

Tobacco mosaic virus (TMV), turnip mosaic virus (TuMV), cucumber mosaic virus

(CMV), and alfalfa mosaic virus (AMV) have been reported on wasabi. These viral

diseases cause stunting and discolouration of leaf veins and/or leaves. Symptoms are

most visible in early spring, when new leaves emerge (Chadwick, 1993; Fletcher, 1989).

The most common fungal pathogens of wasabi include: Phoma wasabiae Yokogi,

thought to cause rhizome blackening; Sclerotinia sclerotiorum (Lib.) de Bary which

causes cottony or watery soft-rot in wasabi seedlings (Chadwick, 1993); Peronospora

alliariae f. sp. wasabiae Gaiimann which causes downy mildew (Chadwick, 1993).

Plasmodiophora brassicae Woronin is an increasing problem to Japanese growers

(Chadwick, 1993), causing swelling of the vascular tissue in the roots and rhizomes

which leads to nutrient and water deficiencies (Adachi, 1987; Chadwick, 1993). Two

reported fungi cause damping-off of seedlings, namely Pellicularia Jilamentosa (Pat.)

Rogers (Adachi, 1987; Chadwick, 1993) and Rhizoctonia solani Kuhn (Adachi, 1987;

Chadwick, 1993). Leptosphaeria maculans (Sowerby) P. Karst causes leaf spots and

blackens wasabi petioles (Broadhurst and Wright, 1998). Albugo wasabiae Hara is a

minor pathogen on wasabi causing white rust (Adachi, 1987; Nozu et al., 1978;

Chadwick, 1993). Other fungal diseases that have been mentioned in the Japanese

literature include Ascochyta brassicae Thuman, Septoria wasabiae Hara and Alternaria

brassicae (Berkeley) Saccardo (Kishi, 1988; Chadwick, 1993).

Bacteria reported on wasabi include: Erwinia spp. (Goto and Matsumoto, 1986b;

Adachi, 1987; Goto and Matsumoto, 1986a; 1987); Pseudomonas spp. (Goto and

Matsumoto, l986a; l986b); and Corynebacterium spp. (Adachi, 1987; Matsumoto et al.,

1985). Corynebacterium spp are believed to cause ring rot in wasabi (Matsumoto et al.,

1985; Adachi, 1987). Leaf spots, blights, vascular wilts, blackened leaf veins, plus

damage to the rhizome and the root vascular tissue, are symptoms of this disease

(Chadwick, 1993). In 1986, the following bacteria were isolated from Japanese-grown

wasabi: Erwinia carotovora subspecies carotovora (Jones) Bergey et al., E. carotovora

subspecies wasabiae (Goto), Erwinia rhapontici (Millard) Burkholder, Pseudomonas

marginalis (Brown) Stevens, and Pseudomonas viridijlava (Burkholder) Dowson.

However, rhizome inoculation experiments were not reported and therefore the effect

these bacteria have on wasabi quality was not determined. In 1998, Erwinia spp. and

Pseudomonas spp. were isolated from New Zealand-grown wasabi. However, bacterial

inoculation experiments were not conducted.

1.5. Tissue discolouration

1.5.1. Stem blackening in wasabi and Phoma wasabiae

The biggest challenge facing wasabi growers worldwide is a blackening of the

rhizome tissue, which results in lower grade or poor quality rhizomes (Chadwick, 1993).

In Japan, reports describe growers harvesting the crop early in order to minimize loss, or

abandoning fields where symptoms are severe (Chadwick, 1993). Blackening of petioles,

leaves and roots have also been reported (Lo et al., 2002; Chadwick, 1993; Adachi, 1987;

Goto and Matsumoto, l986a); however, these tissues are not as valuable and therefore are

of less concern. To date, no control measures are regarded as highly effective in reducing

rhizome blackening (Lo and Wang, 2000b; Lo et al., 2002; Chadwick, 1993; Lo and

Wang, 2001).

Rhizome blackening has been observed in the cortex, vascular tissue, epidermis,

and/or pith (Adachi, 1987; Chadwick, 1993; Sparrow, 2006; Goto and Matsumoto,

1986a). Regardless of the tissue in which blackening occurs, the condition is referred to

as blackleg (Chadwick, 1993; Yokogi and Notsu, 1933), streak disease (Lo et al., 1990;

Wang et al., 1992), internal black rot syndrome (Goto and Matsumoto, 1986a), or black

rot disease (Lo and Wang, 2000b). In 1933, Yokogi and Notsu described "sumiiribyo",

or blackleg as it has been translated, as blackening of the petioles, leaf veins and some

roots (Chadwick, 1993). Lo (1990) and Wang (1992) used the term streak disease as a

synonym for blackleg. Adachi (1987) described the disease as necrosis of the vascular

tissue (Chadwick, 1993), although photographic images depict leaf holes with visible

pycnidia, petiole blackening, vascular blackening and blackening of the rhizome pith.

Internal black rot syndrome was described by Goto and Matsumoto (1986a) as the black

discolouration of vascular tissue, cortex and epidermis of the rhizome and fibrous roots.

Lo (2000a, 2000b, 2001, 2002) used the term black rot disease to describe leaf spot,

petiole blight, root rot, and black rot and/or vascular streaking of the rhizome.

Although the disease name and exact symptoms are unclear, it is widely thought

that the Ascomycete P. wasabiae is the causal agent of all of the symptoms, including

vascular blackening in leaves, petioles and rhizome, leaf holes and blackened roots, plus

blackening in the rhizome pith, cortex and epidermis. P. wasabiae is described as a

highly destructive fungus causing disease in both soil- and water-grown wasabi (Lo and

Wang, 2000b). The fungus is reportedly systemic and has been introduced to new wasabi

growing regions on vegetatively propagated plants by farmers who acquire plantlets from

areas where wasabi is infected with the disease or by shared farming equipment (Lo et

al., 2002). The disease is perpetuated by the production of pycnidia and pycnidiospores,

which are sources of primary and secondary inoculums (Lo and Wang, 2000a; Lo et al.,

2002; Lo and Wang, 200Clb). Fungicide control programs were implemented in 1973;

however, they have been largely ineffective in controlling the disease (Lo et al., 2002).

In 1987 Adachi, suggested that vegetative propagation be limited to only two successive

plantings in order to break the disease cycle (Chadwick, 1993). Despite attempts to

control rhizome blackening, this disease continues to cause major crop loss and crop

devaluation.

1.5.2. Tissue discolouration in other plant species

Many plant species exhibit discolouration in their stems, roots, or leaves due to

either abiotic stress or microbial infection. Boron deficiency, wounding, UV light or low

oxygen levels can cause tissue blackening (Nautiyal, 1986; Matsuki, 1996), while

pathogenic fungi or bacteria are known to cause tissue discolouration in a wide range of

plant species.

At a cellular level, discolouration is independent of cell type when caused by

abiotic factors (Matsuki, 1996). For example, wounding of cassava (Manihot esculenta

Crantz) root, which may occur during harvest, causes the vascular tissue to become blue-

black within 24 - 48 hours due to phenolic compound accumulation. However, this

rapidly spreads to include all parenchyma tissues so that at three days post-wounding, the

entire root is discoloured (Rickard and Gahan, 1983). Boron deficiency in crucifers also

causes non-specific tissue discolouration. For example, in rutabagas (Brassica

napobrassica Mill.) and turnip (Brassica rapa var. rapa L.), discolouration in the root

can occur as scattered grouped or circular patterns throughout, usually in the lower two-

thirds of the root (Howard et al., 1994).

In contrast, tissue discolouration that occurs due to biotic infection is usually cell-

specific (Matsuki, 1996). Xanthomonas campestris pv. campestris causes black rot of

susceptible cabbage varieties (Brassica oleracea). The bacterium multiplies in the xylem

vessels causing the characteristic discolouration of vascular tissue (Staub and Williams,

1972). Similarly, phenolic compounds accumulate in xylem parenchyma cells of tomato

(Lycopersicon esculentum Mill.) and banana (Musa spp.) as a result of infection by the

fungi Fusarium oxysporum Schlecht f. sp. lycopersici (Sacc.) Snyder & Hansen (Mace et

al., 1972) and F. oxysporum Schlecht f. sp. cubense (E.F. Smith) Snyder & Hansen

(Ploetz, 2005), respectively.

1.5.3. Phenolic compounds

The discolourations observed in plants experiencing abiotic or biotic stress are often

due to the oxidation of phenolic compounds. These compounds are comprised of one or

more benzene rings with one or more hydroxyl group and are synthesized by the plant

and play a role in plant defence against pathogen attack. Phenolic compounds are

numerous and extremely diverse, differing between tissues and between species

(Beckrnan, 2000).

Phenolic-storing cells synthesize, store, and facilitate oxidation of phenolic

compounds when various environmental stresses occur. Phenolics are thought to be

synthesized in the thylakoids of plastids and relocated to the vacuole, where the hydroxyl

group of the phenolics can be maintained in a non-ionized, reduced state due to the high

H+ concentration maintained by the H + - A T P ~ S ~ pumps in the tonoplast (Beckman, 2000;

Wink, 1997). Phenolic-storing cells are distributed within the plant to allow for

maximum protection. For example, they may occur in root tips, which is also the

interface with potential pathogens (Mueller and Beckman, 1976).

When plants are attacked by a systemic pathogen, the potential for extensive and

rapid spread of the pathogen through the xylem is extremely high. Therefore, it is not

surprising that phenolic-storing cells can be found in the xylem parenchyma. This has

been well documented for banana, cotton, tomato, and potato (Mace, 1963; Mace and

Howell, 1974; Mace et al., 1972; Mueller and Beckman, 1974; 1976; Waggoner and

Diamond, 1956). Following vascular infection of many plant species, the stored

phenolics diffuse out of the vacuole and into the cytoplasm, where they are oxidized,

which is visualized as tissue discolouration (Beckman et al., 1989).

The oxidized phenolics play a role in lignification and vascular defence signalling.

Oxidized phenolics can polymerize with each other or with cellular proteins or

carbohydrates to cause lignification, which is an attempt by the host to physically block

the vascular infection (Beckman, 2000). Phenolic compound oxidation can also signal a

long-distance defence mechanism, by mediating the conversion of tryptophan to 3-indo-

leacetic acid (IAA)(Gordon and Paleg, 1961). IAA levels rise sharply as oxidized

phenolics also prevent the destructive oxidation of IAA. IAA build-up has been shown to

promote ethylene production (McKeon et al., 1995; Beckman, 2000). Studies where

vascular infection of a plant occurs, shows that cytokinin levels decrease in vascular

tissues (Misaghi et al., 1972; Cahill et al., 1986). Therefore, the IAA/ethylene/cytokinin

hormone balance is shifted due to the oxidation of phenolic compounds. This mobilizes a

periderm-like defence beyond the point of immediate infection and can promote lateral

growth through the formation of tyloses, gels, and/or crushed elements (Beckman, 1966),

which are in turn often lignified by phenolic compounds or suberized by lipid infusion.

These short and long distance responses initiated by phenolic oxidation result in sealing

off the endangered xylem. The unaffected xylem reconnects above this zone (Beckman,

2000).

When an organism is not a host-specific pathogen, this response is strong and rapid

and localised discolouration may occur. When the organism is a host-specific pathogen,

phenolic oxidation and subsequent tissue discolouration may occur; however, it is

ineffective in preventing infection of the xylem and disease will occur (Beckman, 2000).

1.6. Pythium

1.6.1. Taxonomy and identification

In 1858, Pringsheim created the genus Pythium with P. monospermum Pringsh. as

the type species (Martin, 1992). Pythium is in the class Oomycota (or

Peronosporomycota) and under the 6-kingdom system; Oomycota was moved from the

kingdom Fungi or Eumycota to the kingdom Chromista, phylum Heterokonta. This move

reflected taxonomist's view that Oomycota were "colourless" algae rather than true fungi.

The differences between true fungi and those in the class Oomycota include: 1)

Oomycota undergo oogamous reproduction and true fungi do not produce oospores; 2)

cells walls are composed of beta-glucans and cellulose rather than chitin; 3) Oomycota

produce heterokont (two unequal flagella) motile zoospores, while although some fungi

produce zoospores they are of one kind; 4) Oomycota mycelia are generally composed of

coenocytic hyphae that contain diploid nuclei, unlike the haploid nuclei contained within

septate mycelia; and lastly, 5) the cristae in the mitochondria of Oomycota are tubular,

whereas true fungi have flattened cristae (Rossman and Palm, 2006).

Although the Oomycota share similarities with some algae, the algae are generally

regarded as photosynthetic and not heterotrophic like Oomycota. For this reason, some

researchers now believe that the Oomycota should be placed into a new kingdom:

Straminopila, along with the brown algae and diatoms (Rossman and Palm, 2006).

Pythium spp. can be difficult to identify morphologically, due to the lack of

formation of defining structures in culture. One of the most reliable ways of identifying

Pythium spp. is by amplifying the internal transcribed spacer (ITS) regions and the 5.8s

gene of the nuclear ribosomal DNA, using universal eukaryotic primers. Subsequently,

the amplified DNA can be sequenced using specific primers for the resultant 2 amplified

regions. Finally, using a BLAST (Basic Local Allignment Search Tool) search,

sequences are compared to other sequences of the same regions in a database, such as

GenBank (Lkvesque and De Cock, 2004).

1.6.2. Pythium spp. as pathogens

Pythium spp. are found throughout the world and can be either saprophytic or

parasitic. They are often parasitic when conditions are favourable for the fungus and less

favourable for the host and may become highly pathogenic, causing rot of root, stems and

fruit, damping-off on seedlings (van der Plaats-Niterink, 1981), root discolouration and

plant loss (Owen-Going et al., 2003; Stanghellini et al., 1988). Their deleterious effects

may be heightened by creating entry wounds in a plant for other pathogenic microbes or

playing a role in the infection and translocation of viruses (van der Plaats-Niterink,

1981). Finally, it has been demonstrated that Pythium spp. can synergistically cause

disease with other fungi (Garcia and Mitchell, 1975a; Garcia and Mitchell, 1975b; Frank,

1972).

Infection occurs when zoospores produce germ tubes which subsequently form

appressoria, penetrating the plant root through an infection peg. Zoospores arise from

vesicles outside of sporangia. Sporangia are born on either resting thick-walled oospores

or directly from mycelia (van der Plaats-Niterink, 1981; Martin, 1992; Howard et al.,

1994). The ability to produce asexual motile zoospores means that Pythium diseases can

spread very rapidly under favourable conditions.

The two main factors determining zoospore germination and subsequent disease

level are temperature and water availability. The optimum temperature condition is

dependent on the Pythium spp. involved, and can differ for zoospore germination and

mycelial growth. A film of water in the growing medium is required for zoospore

motility and germination, plus the disease severity can be dependent on the amount of

excess water in the medium. Additional factors influencing infection and extent of

disease include pH, light intensity, cation content in the water, the presence of other

micro-organisms, and inoculum concentration (van der Plaats-Niterink, 198 1).

The virulence of Pythium spp. is determined by the capacity to produce enzymes

and phytotoxins which have been demonstrated in many species (van der Plaats-Niterink,

1981). Of the two species discussed below, only cellulolytic enzymes have been found to

be produced by P. intermedium (Deacon, 1979).

1.6.3. Pythium dissotocum

Pythium dissotocum Drechsler was initially isolated by Drechsler in 1930 and 1940

from sugarcane (Saccharum oficinarum) in the USA (Drechsler, 1930; 1940). It is

considered a fast-growing Pythium sp., able to grow 13 mm per day, on potato-carrot

agar at 25 "C. Its optimal temperature range is 20 to 25 "C, with a minimum of 5 "C and

maximum of 35 "C; zoospores are formed at 5 to 20 "C (van der Plaats-Niterink, 1981).

P. dissotocum is one of many species where oogonia are often not observed in

culture; these are grouped according to the morphological characteristics of the

sporangia. According to this classification system, P. dissotocum belongs to 'Group F',

as it has filamentous, non-swollen sporangia (van der Plaats-Niterink, 198 1). As a result,

P. dissotocum is often not identified to species level, but partially identified to Group

level, and therefore the extent of its range and disease capacity may be greater than

presently understood. Nevertheless, only those isolates identified as P. dissotocum will

be presented here.

P. dissotocum has been isolated from soil, water and numerous plant species

throughout the world. It has been isolated from plant roots and has been reported to

cause pre- and post-emergence seedling damping-off (van der Plaats-Niterink, 1981).

However, a few researchers have investigated the effect that P. dissotocum has on older

plants. These studies are conducted on hydroponically-grown crops, especially those

where the nutrient solution is maintained between 20 and 25 "C. Inoculation experiments

conducted on established lettuce (Lactuca sativa cv. Salina), spinach (Spinacia oleracea

L.) and sweet peppers (Capsicum annuum L.) plants, showed that P. dissotocum can

cause significant root loss and yield reduction (Stanghellini and Kronland, 1986; Owen-

Going et al., 2003; Bates and Stanghellini, 1984). Effects on roots are dependent on

isolate, turning yellow to brown, as seen in sweet pepper (Owen-Going et al., 2003), or

roots appear white and healthy as was observed in lettuce (Stanghellini and Kronland,

1986).

1.6.4. Pythium intermedium

F'ythium intermedium de Bary was originally isolated from dead plant material, but

is found in soils throughout the northern hemisphere (van der Plaats-Niterink, 1981). It

grows very fast, extending 30 mm per day when grown on potato-carrot agar at 25 "C. Its

optimal temperature range is identical to P. dissotocum at 20 and 25 "C, with a minimum

of 5 "C and maximum of only 30 "C (van der Plaats-Niterink, 198 1).

The production of sporangia and zoospores by P. intermedium has only been

reported twice (Mattews, 193 1 ; Middleton, 1943), and they have not been observed since.

Oogonia are produced when compatible isolates are grown together; however, the species

is often identified morphologically by the abundant hyphal swellings which can form

dense chains (van der Plaats-Niterink, 198 1).

P. intermedium has been isolated from a range of plant species and is thought to

cause damping-off in many of them (van der Plaats-Niterink, 1981). As with P.

dissotocum, the number of infection experiments on established plants is limited;

however, it is shown to cause cavity spot in carrot (Howard et al., 1994; Allain-Boule et

al., 2004) and root-tip browning, root loss and yield reduction of hydroponically-grown

cucumbers (Stanghellini et al., 1 988).

1.6.5. Control methods for Pythium diseases

Pythium spp. are one of the primary groups of organism which cause pre- and post-

emergence damping-off and they are usually not host-specific (Hendrix and Campbell,

1973). Pythium spp. can also cause other diseases in established plants, as mentioned

previously. This disease dichotomy which is dependent on the plant's age determines the

timing of the control method to be implemented. Pythium control may occur before

planting and/or shortly after planting in order to decrease the losses caused by damping-

off. However, Pythium-related diseases on older plants may be controlled by treating an

established plant. Irrespective of the stage at which control occurs, they may be

chemical, biological effective cultural practices utilized.

Preplant treatment is usually implemented to decrease the likelihood of plants dying

due to damping-off (Cook, 2000). Soil steaming, pasteurization, or fumigation can

decrease Pythium populations to near zero. Often, pasteurization is preferred as it

maintains soil structure and does not kill all organisms, while excessive heat treatment

may yield the soil unfit for plant growth (Hendrix and Campbell, 1973). Fumigation with

various biocides, such as chloropicrin or methyl bromide, can be very effective in

decreasing initial Pythium populations (Hendrix and Campbell, 1973; Sutherland and

Shrimpton, 1989). Another preplant strategy is the application of fungicides to seeds,

which may allow plants to become established beyond the stage where they are

susceptible to damping-off. However, treatment may be phytotoxic (Sutherland and

Shrimpton, 1989).

Non-chemical pre-planting and post-planting control methods include soil

amendments, such as sawdust, bark, crop residues, green manuring, and compost. These

are effective as they encourage soil flora that are antagonistic to pathogenic Pythium spp.

(Hendrix and Campbell, 1973; Noble and Coventry, 2005). Alternatively, micro-

organisms commonly referred to as biocontrol agents, can be applied to growing medium

or plants at various times of the growing season in order to control pythiaceous species

(Table 1.1). The antagonistic effect that other organisms have on Pythium spp. can occur

through a variety of mechanisms including: production of antibiotic compounds,

secretion of enzymes, parasitism, competition for nutrients and infection sites,

interference with pathogenicity factors, or induced resistance in the plant host (Punja and

Utkhede, 2003). Post-planting chemical treatments involve the use of fungicides. Many

are available on the market (Table 1.2) with the most common active ingredient against

Pythium spp. being metalaxyl.

In addition to chemical and biological controls, good cultural practices can be

employed by growers in order to minimize the effects of Pythium spp. The use of

cultivars with increased resistance to Pythium diseases can be very successful, but is

dependent on availability. Probably the most effective cultural practice in reducing loss

due to Pythium spp. is the reduction of water availability (Hendrix and Campbell, 1973).

A film of water in the growing medium is required for zoospore motility and germination

(van der Plaats-Niterink, 1981); therefore, limiting the period where free-water is

available can decrease infection severity. However, this is only possible in the

greenhouse in a non-hydroponic growing system or in the field where rainfall is limited

and soil water-content can be controlled by irrigation (Hendrix and Campbell, 1973).

Tab

le 1

.1.

Bio

cont

rol a

gent

s av

aila

ble

for c

ontr

ollin

g di

seas

es c

ause

d by

Pyt

hium

spp

.

Pro

duct

trad

e na

me

Act

inov

ate

SP

Cill

us

Gre

en-a

ll G

K

odia

k M

ycos

top

* Sy

stem

3 (K

odia

k A

T)

Bin

ab T

Vec

tor

Bin

ab T

F B

inab

T W

G

Gre

en-a

ll T

WP

Plan

tshi

eld

HC

* Po

lyve

rsum

Pr

esto

p W

P Pr

imas

top

Roo

tshi

eld

Dre

nch

WP

* So

ilgar

d 12

G

Supr

evis

it T

-22

Gra

nule

s *

T-2

2 H

C

Tri

anum

P *

Tri

chod

ry

Tri

chof

low

T

rich

ogro

w

Tri

chop

el

Tri

chop

el R

Mic

roor

gani

sm(s

) con

tain

ed

Stre

ptom

yces

lyd

icus

WY

CD

108

B

acill

us s

ubtil

is

Bac

illus

sub

tilis

B

acill

us s

ubtil

is G

B03

St

rept

omyc

es g

rise

ovir

idis

K6

1 B

acill

us s

ubtil

is G

B03

Tr

icho

derm

a po

lysp

orum

and

Tri

chod

erm

a ha

rzia

num

Tr

icho

derm

a po

lysp

orum

and

Tri

chod

erm

a ha

rzia

num

Tr

icho

derm

a po

lysp

orum

and

Tri

chod

erm

a ha

rzia

num

Tr

icho

derm

a ha

rzia

num

GB

F-20

8 Tr

icho

derm

a ha

rzia

num

T-2

2(K

RL

-AG

2)

Pyth

ium

olig

andr

um

Glio

clad

ium

cat

enul

atum

J14

46

Glio

clad

ium

cat

enul

atum

J14

46

Tri

chod

erm

a ha

rzia

num

T-2

2(K

RL

-AG

2)

Glio

clad

ium

vir

ens

G-2

1

Tri

chod

erm

a ha

rzia

num

Tr

icho

derm

a ha

rzia

num

Rif

ai K

RL

-AG

2 Tr

icho

derm

a ha

rzia

num

Rif

ai K

RL

-AG

3 Tr

icho

derm

a ha

rzia

num

Rif

ai K

RL

-AG

2 Tr

icho

derm

a ha

rzia

num

Rif

ai K

RL

-AG

2 T

rich

oder

ma

harz

ianu

m R

ifai

KR

L-A

G2

Tric

hode

rma

spp.

Tr

icho

derm

a ha

rzia

num

and

Tri

chod

erm

a vi

ride

Tr

icho

derm

a sp

p.

Dat

a ob

tain

ed f

iom

Agr

icul

ture

and

Agr

i-Fo

od C

anad

a (K

abal

uk a

nd G

azdi

k, 2

005)

. * P

rodu

cts

regi

ster

ed fo

r use

in C

anad

a

2 1

Man

ufac

ture

r or

dis

trib

utor

Nat

ural

Ind

ustr

ies I

nc.

Gre

en B

iote

ch C

ompa

ny L

td.

Gre

en B

iote

ch C

ompa

ny L

td.

Gus

tafs

on, L

LC

V

erde

ra O

y H

elen

a C

hem

ical

B

inab

Bio

-Inn

ovat

ion

EFT

R A

E3

Bin

ab B

io-I

nnov

atio

n E

FTR

AE3

B

inab

Bio

-Inn

ovat

ion

EFT

R A

B

Gre

en B

iote

ch C

ompa

ny L

td.

Bio

wor

ks In

c.

Bio

prep

arat

y L

td.

Ver

dera

Oy

Ver

dera

Oy

Bio

wor

ks In

c.

Cer

tis U

SA, L

LC

B

orre

gaar

d B

iopl

ant

Bio

wor

ks In

c.

Bio

wor

ks In

c.

Bio

wor

ks In

c.

Bio

wor

ks In

c.

Bio

wor

ks In

c.

Agr

imm

Tec

hnol

ogie

s Ltd

. A

grim

m T

echn

olog

ies L

td.

Agr

imm

Tec

hnol

ogie

s Ltd

.

Tab

le 1

.2.

Exa

mpl

es o

f fu

ngic

ides

ava

ilabl

e fo

r co

ntro

lling

dis

ease

cau

sed

by Pythium s

pp

Pro

duct

trad

e na

me

Del

ta-C

oat@

D

exon

@

Div

iden

d@

Fena

mid

one@

Fo

re@

H

erita

ge@

* R

easo

n@ 50

0 SC

* R

idor

nil@

* Su

bdue

@ M

AX

X *

met

alax

yl-m

pr

opam

ocar

b m

efen

oxam

+ ch

loro

neb

fena

min

osul

f di

feno

cona

zole

im

idaz

olin

one

man

coze

b az

oxys

trob

in

fena

mid

one

met

alax

yl-m

m

etal

axyl

-m

terr

azol

e

Act

ive

ingr

edie

nt

Syng

enta

Cro

p Pr

otec

tion

Inc.

B

ayer

Cro

p Sc

ienc

e Sy

ngen

ta C

rop

Prot

ectio

n In

c.

Bay

er C

rop

Scie

nce

Gre

en A

gros

ino

Co.

, Ltd

. B

ayer

Cro

p Sc

ienc

e R

ohm

and

Has

s C

o.

Syng

enta

Cro

p Pr

otec

tion

Inc.

A

vent

is C

rop

Scie

nce

Syng

enta

Cro

p Pr

otec

tion

Inc.

Sy

ngen

ta C

rop

Prot

ectio

n In

c.

Che

mtu

ra U

SA C

orp.

Man

ufac

ture

r or

dis

trib

utor

R

efer

ence

(Gra

ham

, 200

3)

(Hsi

ang,

200

6)

(Keg

ley

et a

l., 2

007)

(K

egle

y et

al.,

200

7)

(Gra

ham

, 200

4)

(Mer

cer a

nd L

ator

se, 2

006)

(K

egle

y et

al.,

200

7)

(Hsi

ang,

200

6)

(Cha

put,

2002

) (G

raha

m, 2

003)

(G

raha

m, 2

003)

(K

egle

y et

al.,

200

7)

1.7. Pectobacterium carotovorum (syn. Erwinia carotovora)

1.7.1. Taxonomy

The genus Erwinia was named after the plant pathologist, Erwin F. Smith (De Boer,

2003) and when first described in 1917, it included members of the family

Enterobacteriaceae that were phytopathogenic (Toth et al., 2003). Erwinia bacteria are

gram-negative with peritrichous flagella, which allow bacteria to move in aqueous

environments where they are commonly found (Perombelon, 2002). Several species of

Erwinia are referred to as the soft-rot erwinias, which include E. carotovora, E.

chrysanthemi Burkholder, and E. cacticida Alcorn et al. (Gardan et al., 2003). They are

characterized by the production of extracellular pectolytic enzymes which can cause soft-

rot in a number of plants (Perombelon, 2002; Toth and Birch, 2005; Tournas, 2005). In

1998, Hauben et al. suggested, on the basis of 16s rDNA sequence analysis, that the soft-

rot erwinias be renamed into the resuscitated genus Pectobacterium (Toth et al., 2003).

Recently, this has been widely accepted by bacteriologists so that the soft-rot erwinias are

composed of P. carotovorum, P. chrysanthemi, and P. cacticida (Perombelon, 2002;

Gardan et al., 2003).

P. carotovorum is further divided into subspecies which are mostly named for the

plant species on which they are found and often differ according to their biochemical and

physiological characteristics. The exception is P. carotovorum subsp. carotovorum

(Jones) Bergey et al. which serves as a catch-all for strains that differ from other

subspecies (Fessehaie et al., 2002; Gardan et al., 2003).

1.7.2. Pectobacterium carotovorum subsp. wasabiae and subsp. carotovorum

Pectobacterium carotovorum subsp. wasabiae (Goto) (Pcw) was isolated from

Japanese-grown wasabi in 1986 (Goto and Matsumoto, 1986a) and was proposed as a

new subspecies in 1987. This was based on various physiological and biochemical

characteristics that distinguished it from P. carotovorum subsp. atrosepticum, Pcc, P.

carotovorum subsp. betavasculorum, P. chrysanthemi subsp. zeae and subsp.

chrysanthemi, and finally P. rhapontici (Goto and Matsumoto, 1987).

Both Pcw and Pcc have been isolated from wasabi (Goto and Matsumoto, 1986a)

and reportedly cause soft-rot on excised petioles, rhizomes and roots. They also cause

soft-rot in wasabi, tomato, and tobacco seedlings; however, Pcw is less virulent on intact

tissue than Pcc (Goto and Matsumoto, 1987).

Pcw and Pcc can be differentiated either biochemically or by sequence comparison.

Pcw fails to grow under anaerobic conditions in the presence of KCN (Cowan and Steel,

1974), unlike Pcc (Goto and Matsumoto, 1987). Probably the most accurate method to

differentiate between Pcc and Pcw is by amplifying the 16s-23s intergenic spacer (IGS)

region of the ribosomal gene using published primers (Jensen et al., 1993), followed by

sequencing the small (-510 bp) region using the same primers. A BLAST can then be

performed, comparing the unknown bacterium to sequences of the same region in

GenBank (Fessehaie et al., 2002).

Since the initial report of Pcw, no further isolation studies have been reported.

However, based on DNA-DNA hybridization studies, phenotypic and serological tests,

plus phylogenetic analysis of 16s rDNA sequences, it has been suggested the Pcw be

elevated to species level resulting in Pectobacterium wasabiae (Gardan et al., 2003).

1.7.3. Soft-rot disease caused by Pcc

Pcc causes soft-rot both pre- and post-harvest (Tournas, 2005) in a number of plant

species (Toth and Birch, 2005; Tournas, 2005). Plant infection may occur pre-planting,

in the ground, or post-harvest (Tournas, 2005; Perombelon, 2002). And despite timing of

infection, soft-rot can develop at any time post-infection, because Pcc is an opportunistic

pathogen (De Boer, 2003). For example, in potato tubers (Solanurn tuberosum L.), plants

may have up to -lo6 cellslg peel of bacteria but are asymptomatic until conditions for the

pathogen become favourable. This often occurs when plant resistance is impaired due to

localised anaerobic conditions, and oxygen dependent production of phytoalexins,

phenolics, free radicals, and cell wall lignin are hindered. Although anaerobic conditions

can cause the facultative anaerobe Pcc (De Boer, 2003) to switch from latent to active

infection, it is not a requirement and experiments demonstrate that sufficiently high

numbers of bacteria can overcome a plant under aerobic conditions (Perombelon, 2002).

Despite localised oxygen levels within plants tissue, Pcc numbers increase to very

high levels before switching from a latent to active infection, and allow the bacteria to

quickly overcome the plant host. This is achieved by quorum sensing, where Pcc uses

the production of N-acyl-homoserine lactone (AHL) to detect population levels within the

host. Once AHL production is greater than its loss, due to diffusion or inactivation,

which corresponds to lo6 to lo7 cells1mL in potato tubers (Perombelon, 2002), bacteria

release pectinase, pectate, cellulase, protease, and xylanase. These enzymes are then

produced exponentially due a positive feedback loop mediated by intermediate plant cell

wall compounds that are products of enzyme degradation (Barnard and Salmond, 2007).

1.7.4. Control methods for soft-rot diseases

Soft-rot diseases can result in large economic loss. In potatoes, the loss due to

Pectobacterium spp. is equivalent to the combined losses from all other potato diseases

(Tournas, 2005). As a result, many of the published methods prescribed for controlling

soft-rot due to Pectobacterium spp. are recommendations for potato growers.

Nevertheless, some of these methods could be carried over into controlling soft-rot in

other plant species. Control methods include chemical and cultural practices, while there

are no registered biological control products.

Cultural practices include strategies to limit the amount of free-water around the

plant, decrease wounding incidence, and good sanitation practices. Limiting free-water is

critical in controlling levels of bacteria. Therefore, if plants are field-grown, preparing

the soil so that it is well-drained may decrease the inoculum level around the plants. This

strategy is not possible in hydroponically-grown greenhouse plants; however, in

greenhouse plants grown in soil, water levels can be controlled to decrease free-water

(Johnson, 1999; Hamm and Ocamb, 2007). Decreasing the amount of wounding on

plants will decrease the infection rate by Pectobacterium spp.; therefore, care must be

taken during planting and weeding (Hartman and Nesmith, 1985). Measures should also

be taken to decrease the occurrence of wounding by feeding insects, nematodes, or by

fungal infection (Johnson, 1999; Hartman and Nesmith, 1985). As wounding is

sometimes inevitable during planting, disease severity can be decreased by planting when

moisture and temperature do not favour disease development (Johnson, 1999). Sanitation

can also reduce the spread of soft-rot bacteria (Hamm and Ocamb, 2007). This includes

disinfecting equipment and immediate removal of infected plants from the growing area

(Johnson, 1999). Finally, using disease-free material to propagate new plants is essential

in breaking the disease cycle (Hartman and Nesmith, 1985).

Chemical control methods to decrease soft-rot are varied and target various stages

at which Pcc can enter into the plant versus controlling the pathogen once soft-rot has set

in. This is most likely due to the rapid onset of soft-rot and the total tissue maceration

caused by the disease. If plant tissue is being used for propagation, washing plant

material with sodium hypochlorite at a rate of 75 ppm available chlorine can prevent

tissue infection prior to planting (Hartman and Nesmith, 1985; Hamm and Ocamb, 2007).

However, when seeds are used for propagation, treating them prior to planting with

fungicides to control pathogen infection can reduce the occurrence of bacterial infection

(Hartman and Nesmith, 1985). Although antibiotics and copper sprays can decrease

bacteria populations, the use of these is not recommended as they can impose a selective

pressure on bacteria thereby increasing bacterial resistance. In addition, the use of

antibiotics may be cost-prohibitive (Cervantes and Gutierrez-Corona, 1994).

1.8. Research objectives

Wasabi is currently being grown commercially in several parts of the lower

mainland and Vancouver Island of British Columbia (B.C.) to provide a quality product

for local use and export. Some limitations currently are availability of healthy planting

material and the occurrence of diseases that reduce plant growth and quality.

This research was conducted so that wasabi growers in B.C. could have access to

planting material that was pathogen-free, independent of seed-germination and seed-

production constraints. An additional benefit to these methods was they would allow

growers to grow specific plants with desired characteristics, such as increased levels of

isothiocyanates.

Wasabi growers require a high-value rhizome within a relatively short growing

period. Therefore, research was also performed to investigate the causal agent of root

loss in B.C., which was most likely decreasing nutrient uptake and slowing plant growth.

In addition, rhizome vascular blackening was investigated as an unknown problem

affecting the local industry. To accomplish this, my research objectives were:

1) To establish a micropropagation technique that can be used to obtain pathogen-

free plant material that can be repeatedly divided by means of axillary shoot

propagation.

2) To determine the causal agent of low root mass in B.C.-grown wasabi.

3) To determine the causal agent of rhizome vascular blackening observed in

wasabi grown in B.C.

CHAPTER 2: MICROPROPAGATION OF WASABI (WASABIA JAPONICA)

FROM NIERISTEM TIPS AND AXILLARY SHOOTS

2.1. Introduction

Wasabia japonica Matsumura, a member of the Cruciferae family, is a perennial

herb which is botanically related to nasturtiums (Chadwick, 1993). While the species is

native to Japan, it is being cultivated commercially in many countries, including North

Korea, New Zealand, Israel, Brazil, Thailand, Taiwan, Canada, and the USA (Follet,

1987; 1988; Palmer, 1990). Wasabi is grown for the food and pharmaceutical industries.

The rhizome, which takes approximately 18 - 24 months to develop, is often grated and

eaten as a condiment in Japanese cuisine, while the leaves can be eaten in salads.

Numerous allyl-isothiocyanates found in wasabi are believed to have health-promoting

properties (Jordt, 2004).

In order to initiate new plantings of wasabi, commercial producers use axillary

shoots with subtending leaves for vegetative propagation. This method is preferred since

establishing new plants from seeds is difficult because of an extended seed dormancy

period, a low germination rate, and high seedling mortality (personal communication

Oates, 1993). However, continuous vegetative propagation of wasabi can result in a

gradual decline in the growth rate (Chadwick, 1993), and also allows pathogens to build

up, thereby reducing plant survival and potentially spreading pathogens. Tissue culture

may provide a method by which wasabi growers can continually acquire large numbers

of disease-free planting stock.

Previous tissue culture work on wasabi has focused on development of somatic

embryos and micropropagation from zygotic embryos (Eun, 1995; 1996; 1999). Other

studies have evaluated apical meristems (Eun, 1997) and axillary buds (Hosokawa, 1999)

in order to micropropagate wasabi. The cultivar(s) of wasabi used in previous studies

was not stated. The media used were either Murashige and Skoog (MS) (Murashige,

1962), Gamborg's B5 (Gamborg, 1968), or a modified YK(N03) (Fukunaga, 1978), with

a range of growth regulators. Shoots derived from apical meristems were induced to

produce roots in vitro on MS supplemented with 0.01 mg/L IBA, and plantlets were

subsequently transferred to sterilized soil (Eun, 1997). However, a protocol for wasabi

micropropagation was difficult to establish based on previous published research due to a

number of reasons. The few published reports are written in Korean or Japanese, and

sufficient details regarding the methodology used and the reproducibility of the results

were not supplied. Hosokawa et al. (1999) reported results on axillary bud culture but the

sterilization protocols were not described. This is critical in order to successfully

establish explants from plants that have been grown under nonsterile conditions, such as

the greenhouse.

The objective of this research was to develop a method for micropropagation of

greenhouse-grown wasabi using axillary buds to produce pathogen-free plants. The

method was intended for commercial application, allowing growers access to a large

number of disease-free plants. We examined the effects of various tissue sterilization

procedures, effects of growth regulators (BA and kinetin) on shoot proliferation, and

various rooting media on root development of plantlets. Plantlets were also acclimatized

and transferred to the greenhouse.

2.2. Materials and methods

2.2.1. Plant material and meristem sterilization

Wasabi plants 'Daruma' were obtained from a commercial greenhouse located in

Richmond, British Columbia in October 2003. The plants were initiated from seed and

grown under semi-hydroponic conditions for 12 to 18 months. All of the plants used

were growing vigorously and appeared healthy. The efficiency of tissue sterilization was

investigated by sterilizing the dissected meristem-tip or alternatively by sterilizing the

axillary bud and then subsequently dissecting the meristem-tip aseptically. The axillary

bud region was trimmed from the plants and depending on the plant size, 10 buds were

obtained per plant. The explant was approximately 1 cm wide by 0.5 cm in length after

the leaves were trimmed off. The tissue containing the axillary bud was rinsed under

running tap water for five minutes. The tip with approximately five leaf primordia,

measuring between 1 and 1.5 mm, was excised from the axillary shoot and sterilized for

15 minutes in a 0.5% NaOCl solution containing Tween 20, subsequently rinsed four

times in sterile distilled water and finally soaked in sterile distilled water for five minutes.

Alternatively, the axillary bud was sterilized using the same methodology, and the

meristem-tip was then aseptically dissected from the bud in a laminar flow hood.

Twenty-five explants, originating from six plants, were used in each treatment.

The meristem-tips were individually placed in a 60 mm petri dish on Gamborg's B5

medium (Gamborg, 1968), which was used as the basal media (BM), solidified with 2.2

g/L phytagel (Sigma), supplemented with 0.5 mg/L ~ ~ - b e n q l a d e n i n e (BA), and pH was

adjusted to 6.7 - 6.8. The meristem-tips were incubated on a laboratory bench, where the

temperature was approximately 21-23 OC with a 16 hour photoperiod, under a light

intensity of 14 ~ r n ~ l r n ~ ~ s ~ ' provided by white fluorescent tubes.

The experiment was repeated in October 2005, using vegetatively propagated plants

or seed-derived plants, grown under semi-hydroponic conditions for 3 - 12 months. The

meristem-tip with approximately five leaf primordia was excised from the axillary bud

and sterilized for 15 minutes in a 0.5% NaOCl solution containing Tween 20,

subsequently rinsed four times in sterile distilled water and finally soaked in sterile

distilled water for five minutes. One hundred axillary meristem-tips originating from one

hundred plants, were sterilized and placed in individual 60 mm petri plates containing

Gamborg's B5 medium as described above and incubated under the conditions outlined

previously.

2.2.2. Shoot initiation

After the meristem-tips had grown in the petri plates for 10 days, those that were

free of contamination and appeared viable were transferred to 25 ml glass culture vessels

(Sigma) containing 20 ml of the same medium. The vessels were placed inside a Percival

growth chamber at 16 OC with a 16 hour photoperiod and light intensity of 14prn0lm'~s~'

provided by white fluorescent tubes. These meristems produced axillary shoots, which

were separated and established as main shoots and which subsequently grew to produce

additional axillary shoots (Fig. 2.1 .c, d). Shoot growth was monitored and shoot clusters

were divided every two months and transferred to fresh medium. A portion of the shoots

were used to determine ideal rooting conditions and the remainder of the shoots were

used to establish optimal shoot proliferation conditions.

Figure 2.1. Tissue culture and micropropogation of wasabi ( Wasabia japonica cv.

Daruma).

a) An 18 month old Wasabia japonica 'Daruma' plant. The arrow indicates the position of an axillary bud. b) A harvested wasabi rhizome, which is grated and eaten as a condiment. The arrow indicates the position of an axillary bud. c) Shoots developed from meristem-tips. d) Multiple axillary shoots (indicated by white arrows) developed from a main shoot (indicated by black arrow) after growing for two months on Gamborg's B5 at 18 OC with a 16 hour photoperiod and a light intensity of 10 pmolm-2s-' provided by white fluorescent tubes. e) Root development of the meristem tip derived shoots grown on perlite for two months with 10 ml BM, under the conditions described above. f) Root development of the meristem tip derived shoots grown on gravel for two months with 10 ml BM, under the conditions described above. g) Root development of the meristem tip derived shoots grown on 50 ml BM with 2.2 g/L phytagel for two months, under the conditions described above. h) Plantlets that were established from meristem tissue culture and grown hydroponically for two months, under the conditions described above. i) Plantlets that were established from meristem tissue culture and grown in steer manure for one month, under the conditions described above.

2.2.3. Shoot proliferation

Axillary shoots that had four leaves with petioles approximately 8 mm long were

separated and placed on BM solidified with 2.2 g/L phytagel (Sigma) and supplemented

with either 0.2, 0.5, 0.8, or 1.0 mg/L BA, or with 0.8, 1.0, 1.2, or 1.4 mg/L kinetin. Each

25 ml glass culture vessel (Sigma) contained three shoots and there were 10 culture

vessels per growth regulator treatment. The shoots were distributed randomly among the

different media, so that genotypic homogeneity within a treatment was avoided. All

shoots were placed within a growth chamber under the conditions described previously.

This axillary shoot became the main shoot for the new culture. After two months, the

new axillary shoots that developed were separated from the main shoot. One of the

axillary shoots with four petioles was transferred to fresh medium. The transfer was done

to avoid a carry-over effect from the medium on which the original axillary shoots had

developed. Two months later, the number of total shoots (axillary shoots and main shoot)

that had four leaves or more was determined. The experiment was done twice.

2.2.4. Root development

Axillary shoots that had been established on 0.5 mg/L BA were separated from the

main shoot and transferred to 7 different growing media as follows: perlite, vermiculite,

perlite : vermiculite mix at a ratio of 1 : I , or gravel, all of which received 10 ml of

hormone-free BM; BM solidified with 2.2 g/L phytagel supplemented with 0.2 mg/L

kinetin and 1.0 mg/L a-napthalene (NAA) or hormone-free; and a liquid hormone-free

BM was used where the shoot was suspended on a LifeRaftTM (Osmotek LifeLine).

There were five replicates per treatment, and plants were carefully distributed so that

each source plant was represented once on each type of media. Only shoots that had two

or more leaves measuring 5 mm or more were used. Each shoot was grown separately in

a Magenta vessel (Sigma-Aldrich). Cultures were placed inside a growth chamber under

the conditions described previously. After one month, the number of roots that

developed was counted and the plants were assessed for overall vigor.

2.3. Results and discussion

2.3.1. Meristem sterilization

The axillary buds that were sterilized in October 2003 produced 36 and 40 % sterile

meristems when they were sterilized before and after dissection, respectively. Therefore,

meristems used in October 2005 were dissected from the bud and then sterilized. This

resulted in 36 % clean and viable meristems after 10 days. The major problem resulting

in meristem contamination was high levels of bacteria, which was exacerbated by the

relatively high pH of the media; however, since wasabi prefers only slightly acidic

growing conditions (Chadwick, 1993), the pH was not reduced.

2.3.2. Shoot proliferation

Shoots that were placed on 0.5, 0.8, 1.0 mg/L BA and 1.4 mg/L kinetin produced

the most number of axillary shoots, ranging from 3.2 to 3.8 mean number of shoots

(Table 2.1). Shoots grown on 0.2 mg/L BA resulted in 3.0 shoots which was

significantly different from the highest shoot mean grown on 0.5 mg/L BA. On 0.8, 1.0,

1.2 mg/L kinetin and 0.2 mg/L BA the least number of mean shoots was observed. In

addition, shoots grown on I .O mg/L BA produced shoots which appeared mildly chlorotic

perhaps due to borderline phytotoxic effects. Shoots grown on different concentrations

of kinetin were highly variable in size producing large and many small shoots. We

selected 0.5 and 0.8 mg/L BA as the optimal cytokinin concentrations for obtaining

maximum shoot production.

Table 2.1. The effects of different cytokinins (kinetin and BA) on shoot proliferation on Gamborg7s B5 with 2.2 g/L phytagel. Shoots were grown at 16 OC with 16 hour day, under a light intensity of 14 pmolm-2s-' provided by white fluorescent tubes. The number of shoots were determined after 8 weeks and the average number of shoots is presented with the number in brackets being the standard error of the mean. Three shoots were grown per 25 ml glass culture vessels (Sigma-Aldrich), and ten culture vessels were used Der treatment.

Cytokinin

Kinetin 1 0.8

Average number of shoots generated

3.0 (*O. 14) bc

3.8 (50.13) a

3.6 (h0.12) ab

3.3 (h0.16) abc

Hosokawa et al. (1999) reported that the average number of shoots produced when

meristems were placed on YK(N03) with 1.0 mg/L BA for 8 weeks and a temperature of

20 "C was 3.6 * 0.8. These findings are similar to ours despite the different growing

conditions used. Although the shoots were grown for 8 weeks, it is difficult to make

further comparisons, since the gelling agent, light intensity, and cultivar of wasabi was

not specified. Furthermore, the shoots were not rooted and their survival in soil was not

specified. Eun et al. (1997) reported that 5.4 shoots were produced from one apical

meristem after 90 days of culture on MS supplemented with I .O mg/L BA. The exact

experimental conditions used could not be compared to those described here, since the

publication was in Korean and the cultivar used was not specified. We used axillary buds

as a source of meristems, and depending on the size of the donor plant used, up to 30

meristem-derived shoots could be established from one plant. This is highly beneficial

given the high initial cost of each wasabi plant. In addition, it would allow for up to 30-

fold increase of a desired plant at one time, allowing the establishment of plant lines to

occur rapidly.

2.3.3. Root development

The shoots that were grown in vermiculite or vermiculite and perlite (Fig. 2.1.f)

with BM failed to develop roots, while those placed in perlite (Fig. 2.1 .f) had one out of

five shoots with roots. These plants appeared water-soaked and some leaves were

chlorotic. Of the five shoots that were placed on gravel with liquid BM, only one

produced roots, and they were short. The plants suspended on the LifeRaftTM (Osmotek

LifeLine) over liquid BM did not survive. Two of the five plants that were grown on

media supplemented with NAA and kinetin developed very short and hairy roots;

however, the leaves drooped and failed to develop further. All of the plants grown on

hormone-free solid BM produced roots (Fig. 2.1 .c) which were long and the plants

appeared healthy.

All plantlets that rooted on hormone-free BM solidified with phytagel were

successfully transplanted into a hydroponic solution. Approximately 350 plants have

been obtained to date using this method. The success rate was almost 100 %. Similarly,

plants that were potted in steer manure (Home Depot) had a high survival rate.

Eun et al. (1997) reported rooting of wasabi shoots using 0.01 mg/L indole-3-

butyric acid (IBA) in solid MS after 15-30 days. The cultivar used was not specified and

the survival following transfer was unclear.

The methodology described here has been used to obtain wasabi plants that are

microbe-free at a high frequency. This can potentially be used by growers to replace

vegetative propagation which spreads disease and which can result in decreased growth

rate (Chadwick, 1993). In addition, by using axillary buds, a propagation program can be

established in which plant lines are selected for vigor, pathogen resistance, and taste, and

then micropropagated to reduce pathogens and to ensure quality and quantity of the

wasabi crop more readily.

CHAPTER 3: ROOT INFECTION OF WASABI

(WASABZA JAPONZCA) BY PYTHZUM SPECIES'

3.1. Introduction

Wasabi (Wasabia japonica Matsumura), a member of the crucifer family, is

cultivated in many regions of the world for its rhizome, which is used as a condiment in

Asian cuisine. Wasabi is currently grown commercially in Japan, North Korea, New

Zealand, Israel, Brazil, Thailand, Taiwan, Canada, and the USA (Follet, 1987; Mukogasa,

1988). The plant grows naturally in or near cool mountain streams in Japan and requires

cool, shaded, and humid conditions when grown under artificial cultivation (Chadwick

1993). Although wasabi can be grown in soil, high quality wasabi rhizomes are produced

on stream beds, or when plants are cultivated under hydroponic or semi-hydroponic

conditions (Chadwick 1993). Wasabi plants are usually propagated vegetatively by

excising vegetative offshoots from the rhizome and rooting them (Chadwick 1993).

Continuous vegetative propagation of plants can lead to a decrease in growth rate over

successive generations (Suzuki, 1976). The causes of this growth reduction are not

known.

The cool shaded growing conditions during wasabi culture are ideal for the

development of a range of fungal and bacterial plant pathogens, most of which have been

reported from Japan. Fungal pathogens include Phoma wasabiae Yokogi causing

' Published in: Canadian Journal o f Phytopathology (2007), 29(1): 79-83

40

necrosis of the vascular tissue (Adachi, 1987; Takuda and Hirosawa, 1975; Yokogi and

Notsu, 1933); Rhizoctonia solani Kuhn causing damping-off and foliar infection (Adachi

1987); Sclerotinia sclerotiorum (Lib.) de Bary causing cottony or watery soft-rot in

wasabi seedlings (Chawick 1993); Ascochyta brassicae Thuman (Chadwick 1993),

Septoria wasabiae Hara and Alternaria brassicae (Berkeley) Saccardo which cause leaf

spots (Kishi, 1988); and Albzigo wasabiae Hara and Peronospora alliariae f. sp.

wasubiae Gaumann which cause white rust (Adachi, 1987; Nozu et al., 1978) and downy

mildew (Chadwick 1993), respectively. Plasmodiophora brassicae Woronin also causes

club root in wasabi (Adachi 1987). Bacterial pathogens include Erwinia spp. which cause

soft-rot of the leaves and roots (Adachi 1987), while Pseudomonas spp. are thought to be

associated with "internal black rot syndrome" in wasabi (Goto and Matsumoto, 1986a;

1986b). Moreover, Corynebacterizun spp. are reported to cause vascular wilts, blights,

leaf spot and ring rot (Adachi, 1987; Matsumoto et al., 1985).

In British Columbia, there is limited commercial production of wasabi under semi-

hydroponic conditions in polyethylene greenhouses. The plants are grown for periods of

up to 18 months in gravel beds (Fig. 3.1 .a-c) with frequent daily mistings of water and

nutrient solution. The rhizomes develop to a size of up to 12 cm in length, and the lateral

roots and leaves are trimmed off prior to sale (Fig. 3.l.d-e). Currently, there are no

diseases reported on wasabi in Canada given the limited acreage and extent of

production.

During winter, 2004, wasabi plants harvested for sale were found to have limited

lateral root production, which made them easy to pull from the ground (Fig. 3.l.f).

Rhizome size was also reduced. These plants exhibited wilting during the summer, and

Figure 3.1. The effect of Pythiurn dissotocum and Pythiurn intermedium on wasabi

roots.

a) A gravel bed used for commercial production of wasabi that contains rooted seedlings that were vegetatively propagated. b) Mature wasabi plants after a year of growth. c) Close-up of wasabi seedling planted in a gravel bed. d) A healthy two-year-old wasabi plant. e) The whole (left) and sliced (right) rhizome from a healthy two-year-old wasabi plant. f) A healthy two-year-old wasabi plant after the side-shoots and leaves are trimmed off, revealing the rhizome. Note the masses of roots emerging from below the rhizome. g) A wasabi plant with very few roots collected in January 2004, displaying symptoms of severe root loss and browning. (h-j) Tissue cultured hydroponically grown plants inoculated with h) control; i) P. dissotocum, and j) P. intermedium. Photographs were taken two months after inoculation and incubation at 20 "C. (k-m) Root colonization of wasabi one week after inoculation with k) control, 1) P. dissotocum, and m) P. intermedium. Note mycelial colonization of roots.

in many cases succumbed to infection by other microorganisms, especially bacteria,

which resulted in plant mortality under very warm summer growing conditions. The

objective of this study was to determine the cause of root loss of wasabi, and the potential

role of soilborne fungi.

3.2. Materials and methods

3.2.1. Isolation and identification of fungi

In January 2004, wasabi plants exhibiting severe root loss were harvested from a

commercial greenhouse in Richmond, B.C. Epidermal tissues from rhizomes and roots

of symptomatic plants were surface-sterilized for 30 sec in 70 % ethanol and

subsequently for 1 min in 0.25% NaOCI. After blotting the excess moisture on filter

paper, tissues were plated onto water agar (WA) and incubated at room temperature (22

to 24 "C). Three days later, hyphal tips of each developing colony were transferred to

potato dextrose agar (PDA) containing 300 mg/L streptomycin sulfate (Sigma-Aldrich).

Cultures were identified to genus level using morphological criteria (Martin 1992).

3.2.2. Plant material

Rooted wasabi plants that were derived from meristem-tip culture and therefore free

of pathogens were used for all inoculation experiments. Meristem-tip derived plants

were obtained by excising axillary meristem-tips and sterilizing them in 0.5 % NaOCl for

15 min and subsequently transferring them to 60 mm diameter petri dishes (Fisher)

containing Gamborg's B5 medium (Gamborg et al. 1968) and 0.5 mg/L N~-benzyladenine

(BA), solidified with 2.2 g/L phytagel (Sigma-Aldrich). After 10 days of incubation

under laboratory conditions (22 - 24 "C), developing shoots were transferred to 25 mL

glass culture vessels (Sigma-Aldrich) containing 20 mL of Gamborg's B5 medium with

growth regulators as described above. The containers were placed inside a growth

chamber set at constant 16 OC with a 16 hr photoperiod and light intensity of 10 pmolm-

2 - 1 s provided by cool-white fluorescent lamps. After two months, developing plantlets

were transferred to the same medium without growth regulators, where they rooted after

an additional two months.

For inoculation experiments, tissue-culture derived plants were suspended in liquid

Gamborg's B5 medium by wrapping the base of the stem twice in a 2-cm-wide strip of

sponge and inserting it through a hole in the lid of an opaque 500 mL container (Sigma-

Aldrich). Air was continuously bubbled through the nutrient solution in each container;

the solution was changed every 6 weeks. The plants were grown in a growth chamber set

at 18 "C, 90 % relative humidity with a 16hr photoperiod with a light intensity of 10

pmolm-2s-' provided by cool-white fluorescent lamps for 6 months prior to inoculation.

3.2.3. Inoculation and root colonization experiments

The fungal isolates were grown on PDA with 300 mg/L streptomycin sulfate for

five days. Mycelial plugs (5 mm diameter) were added to the hydroponic solution, with

eight containers per isolate, each containing one plant. Control plants received no

inoculum. The temperature was increased to 20 "C after inoculation. Plants were

harvested after 2 months. Root segments, approximately 1 cm in length, were surface-

sterilized for 30 sec in 70% ethanol and then for 1 min in 0.25% NaOCI, rinsed in sterile

distilled water, blotted dry and plated onto PDA with 300 mg/L streptomycin sulfate. To

obtain root dry weight, the root mass of eight replicate plants was dried at 40 "C for 4

days and weighed. The standard error of the mean was determined. The experiment was

conducted twice. To study root colonization, plants were grown hydroponically as

described above and inoculated with mycelial plugs. One week later, root segments were

examined under the microscope at 10 x and 40 x magnification and evidence of mycelial

colonization was recorded.

3.3. Results and Discussion

3.3.1. Isolation and identification of fungi

In all isolations conducted from approximately 70 affected plants, which were

sampled at 6-8 week intervals during 2004, species of Pythiurn were recovered from most

wasabi root and rhizome epidermal tissues. No other fungal genera were observed,

except for an occasional isolate of Fusarium sumbucinum. At least two morphologically

distinct colonies of Pythitim were observed. Representative strains were identified by

DNA sequencing of the internal transcribed spacer region (Levesque and de Cock 2004).

P, dissotocum Drechsler and P, intermedium de Bary were represented by 1 and 4 out of

5 strains sequenced, respectively. A representative isolate of each species has been

deposited in the National Mycological Herbarium, Department of Agriculture, Ottawa,

Ontario. Approximately 70 cultures of these two species were obtained and they were

generally isolated from different plants, with P. dissotocum being the predominant

species.

3.3.2. Inoculation of wasabi plants and assessing root infection

Two months after inoculation, the roots on control (non-inoculated) plants appeared

white and healthy (Fig. 3.1.h) while the Pythium-inoculated plants had very poor root

development (Fig. 3.l.i, j). However, although the roots of inoculated plants were

significantly reduced, the foliage appeared healthy even after two months. Control plants

had an approximately six-fold higher root dry mass compared to the inoculated plants.

There was no significant difference between the root dry weights of plants inoculated

with either P. dissotocum or P, intermedium (Fig. 3.2). In all cases, Pythiurn spp. were

successf~illy reisolated from the inoculated plants.

The lack of foliar symptoms on inoculated plants may be due to the age at which

the plants were inoculated and the cool growth conditions in hydroponics. If younger

seedlings had been inoculated, the Pythiutn spp. may have caused damping-off.

Damping-off is reported to be a very common problem for wasabi growers in British

Columbia, causing an average of 50 % seedling loss, although under certain conditions,

100 % seedling mortality can occur. Secondly, wasabi is an aquatic plant, and despite

being infected with P. dissotocum or P, intermedium and experiencing severe root loss, it

likely can transport sufficient water to the leaves under the cool growing temperatures

preferred by the plant (Chadwick 1993) and therefore could survive and continue to

grow. It has often been observed by growers that even though wasabi plants growing in

hydroponic or semi-hydroponic conditions have very few roots, the foliage of the plant

seems unaffected.

Many Pythium species are ubiquitous organisms that grow well in running surface

water and often are the causal agents of damping-off and root rot. In most cruciferous

species, P. debaryanum and P. ultimum Trow are the primary pathogens that cause

damping-off in seedling-trays in the greenhouse and in direct-seeded field plantings

Howard et al. 1994). P. dissotocum, often partially identified as Pythium group F, is a

common pathogen of greenhouse-grown lettuce, spinach, tomatoes, and peppers, causing

Control P. dis. P. int.

Treatment

Figure 3.2. Root dry weights from 8-month-old wasabi plants inoculated with Pythium SPP. Tissue culture derived plants were grown hydroponically for 6 months and inoculated with Pythium dissotocum (P. dis.) or P. intermedium (P. int.). Roots were collected after 2 months and dried at for 4 days at 40 O C . Data are the mean from eight replicate plants. Vertical bars represent the standard error. The experiment was conducted twice.

damping-off in seedlings. It attacks the small feeder roots of older plants, causing a

reduction in root mass, browning of tips and lower yield (Howard et al. 1994). P.

intermedizm is found in soils worldwide and has a very wide host range (van der Plaats-

Niterink 1981), which includes carrot (Howard et al. 1994) and cucumbers, where it

causes root infection and decay (Stanghellini et al. 1988). P. dissotocum and P.

intermedizrm both grow rapidly at temperatures between 20 and 25 OC; however, their

minimum temperature for growth is 5 OC, while their upper range is 35 and 30 OC,

respectively (van der Plaats-Niterink 198 1).

The use of vegetative propagation of wasabi for many generations has been noted to

decrease growth rate (Suzuki 1976). This reduced vigour could be due to pathogen

persistence on roots and rhizomes over the generations, with subsequent offspring

yielding a higher proportion of plants infected with pathogens such as Pythi~lm. In

comparison, plants derived by meristem-tip propagation when grown over successive

generations in tissue culture do not show this reduction in growth and vigour (G.

Rodriguez and Z. K. Punja, unpublished). In addition, Pythium species can be introduced

into the greenhouse via equipment, which is frequently shared with other greenhouse

growers, or by runoff water from neighbouring farms. Samples of growing medium

tested prior to planting wasabi and water used for irrigation were free of Pythium

inoculum; however, all growing medium that contained established vegetatively

propagated wasabi plants had Pythizim inoculum present (G. Rodriguez and 2. K. Punja,

unpublished). This suggests that inoculum could be reintroduced to new plantings either

through propagation of diseased material or contamination from external sources.

In addition to the Pythium spp. which caused decreased root mass and reduced

nutrient transport to the growing plant, plants with reduced root mass can wilt during the

warm summer months as well as become predisposed to infection by other

microorganisms, especially bacteria. In Pythium-infected plants, for example, the

occurrence of Erwinia carotovora subspecies carotovora is enhanced, resulting in

softening and blackening of the rhizome (Rodriguez and Punja 2006). Therefore,

sublethal infection of wasabi by Pythium species can lead to other significant problems if

not controlled through proper sanitation, propagation using disease-free material, and

fungicide applications.

CHAPTER 4: RHIZOME BLACKENING OF WASABI CAUSED BY

PECTOBACTERIUM CAROTOVORUM SUBSP. CAROTOVORUM

4.1. Introduction

Wasabi (Wasabia japonica) is a perennial herb which is native to Japan and the

freshly-ground rhizome has been eaten as a condiment since the tenth century (Chadwick,

1993). This high-value crop is currently also being processed for the pharmaceutical

industry. Wasabi can be grown in soil but high-quality rhizomes for the fresh market are

traditionally cultivated in stream beds or in hydroponic or semi-hydroponic conditions.

Wasabi grows best at air temperatures between 8 and 18 OC and growth ceases at

temperatures above 28 O C (Follet, 1986). Aquatically-grown plants prefer a water

temperature of approximately 10 - 13 "C year-round, and although they will grow in

water of higher temperature, rhizome growth can be compromised and plants become

more susceptible to disease (Hodge, 1974). Good quality rhizome production requires

water with a high level of dissolved oxygen (Follet, 1986).

In order to initiate new plantings of wasabi, commercial producers use axillaly

shoots with subtending leaves for vegetative propagation. This method is preferred since

establishing new plants from seeds is difficult because of an extended seed dormancy

period, a low germination rate, and high seedling mortality. However, continuous

vegetative propagation of wasabi can result in a gradual decline in the growth rate

(Chadwick, 1993), and also allows pathogens to build up, thereby reducing plant survival

and potentially spreading pathogens.

In Japan, about 65 % of wasabi grown in an aquatic environment is sold on the

fresh market (Follet, 1986), and a major factor which reduces rhizome quality is a

visually unappealing internal blackening of the rhizome. These rhizomes are often sold

for processing or discarded and make up the remaining 35% of the crop harvested.

Japanese reports describe growers abandoning fields in cases where symptoms are severe

(Chadwick, 1993). The problem has been reported to occur in various regions where the

crop is cultivated, including Japan (Goto and Matsumoto,

1933; Lo et al., 1 WO), New Zealand (Broadhurst and Wright,

2006) and Taiwan (Lo and Wang, 2000b).

1986a; Yokogi and Notsu,

1998), Australia (Sparrow,

Rhizome blackening symptoms are reported to occur in the cortex, vascular tissue,

epidermis, andlor pith (Adachi, 1987; Chadwick, 1993; Sparrow, 2006; Goto and

Matsumoto, 1986a). The problem has been variously referred to in the literature as

blackleg (Chadwick, 1993; Yokogi and Notsu, 1933), streak disease (Lo et al., 1990;

Wang et al., 1992), internal black rot syndrome (Goto and Matsumoto, 1986a), or black

rot disease (Lo and Wang, 2000b). In many studies, it is reported that the Ascomycete

Phoma wasabiae is the causal agent of rhizome blackening symptoms as well as

blackening symptoms appearing on leaves, petioles and roots (Yokogi and Notsu, 1933;

Chadwick, 1993; Adachi, 1987; Sparrow, 2006; Goto and Matsumoto, 1986a; Lo et al.,

1990; Wang et al., 1992; Lo and Wang, 2000a).

In New Zealand, microbes isolated from blackened wasabi rhizomes included

Phoma wasabiae, Phoma chysanthemicola, Leptosphaeria maculans, Erwinia spp.

(possibly syn. Pectobacterium spp.) and Psetidomonas spp. Inoculation experiments

were conducted with fungal isolates only. Plants inoculated with P. wasabiae and L.

maculans developed leaf spots and blackened petioles, while occasionally plants wound-

inoculated with P. wusabiae developed vascular blackening in the rhizome (Broadhurst

and Wright, 1998). Studies in Japan reported a range of bacteria recovered from

blackened cortical, epidermal, root and vascular tissues of the rhizome including;

Pectobacterium carotovorunz subsp. carotovorum and subsp. wasabiae, Pectobacterium

rhapontici, Pseudomonas marginalis, and Pseudomorzas viridrflava. In addition, P.

wasabiae was often isolated from these tissues (Goto and Matsumoto, 1986a; 1987).

However, rhizome inoculation experiments were not conducted to confirm pathogenicity

of these bacteria.

Wasabi is currently grown in British Columbia (B.C.), Canada, and blackened

vascular rhizome tissue was observed in both cultivars Daruma and Mazuma during

2005. In the absence of a confirmed causal agent, the objective of this research was to

identify microbes from affected rhizomes and to complete Koch's postulates to establish

the causal agent.

4.2. Materials and methods

4.2.1. Microbial isolation and microscropy

Wasabi cultivar Daruma plants grown in Richmond, B.C. were collected

approximately every 10 weeks from November, 2005 to August, 2006 and those with

vascular blackening were identified (Fig. 4.1 .a-d). The leaves were trimmed off and the

upper and lower 1 cm of the affected rhizome was discarded. The remainder of the

rhizome was rinsed in running water for 15 min. The pith, vascular, cortex, and

epidermal tissues were sectioned (2 x 2 mm), surface-sterilized in 70 % ethanol for 30 sec

Figure 4.1. Wasabi rhizomes with vascular blackening. (a) Healthy plants. (b) Healthy rhizome. (c, d) Plants with symptoms ol'vascular blackening in longitudinal section (c) and in cross- section (d). (e-h) Microscopic images of wasabi rhizome exhihiling rhizome vascular blackening. (e) Cross-section ol'blackened rhizome vascular tissue slained with toluidine blue; (f) as i n (e), but longi~udinal-section. (g) Cross-section of symptomatic [issue under phase- contrast illumination: gels ( G ) and tylose (T). ( h ) Close-up of xylem parenchyma cells with bacteria (B).

followed by 0.25 % NaOCl for 1 min and plated onto water agar (WA, Anachemia) with

300 mg/L streptomycin sulphate (Sigma-Aldrich) or onto nutrient agar (NA, Difco)

without antibiotics. The dishes were incubated at room temperature (21-23 OC) for 1 to 2

weeks. Fungi growing on WA were transferred to potato dextrose agar (PDA) and

identified to genus level using morphological criteria. Bacteria recovered on NA were

grouped as gram-positive or negative according to the KOH method (Fluharty and

Packard, 1967). Representative colonies of gram-negative bacteria were identified using

BiologTM (Biolog Inc., U.S.A.). Cultures were stored in 80 % glycerol at -80 OC

For light microscopy, segments of wasabi rhizomes with blackening symptoms in

the vascular tissues were fixed in formaldehyde-acetic acid (FAA) for 48 hr and rinsed in

70 % ethanol. Sections (3 pm thick) were cut, immediately stained with 0.1 % toluidine

blue for 30 sec and examined under a light microscope at various magnifications. The

presence of fungal hyphae and bacterial cells was noted.

4.2.2. Plant material for pathogenicity tests

Wasabi plants cv. Daruma were obtained from a commercial greenhouse located in

Richmond, B.C., the leaf petioles were trimmed from the rhizome to expose the axillary

buds (Fig. 4.1 .b) and rinsed under running tap water for 5 in. The meristem-tip measuring

between 1 and 1.5 mm with approximately five leaf primordia, was excised from within

the axillary bud and sterilized for 15 min in a 0.5% NaOCl solution containing Tween 20,

rinsed four times in sterile distilled water, and finally soaked in sterile distilled water for

5 min. Meristem-tips were placed in 60 mm petri dishes containing Gamborg's B5

medium (Gamborg et al., 1968) with 0.5 mg/L N~-benqladenine (BA)(Sigma-Aldrich),

solidified with 2.2 g/L phytagel (Sigma-Aldrich), The meristem-tips were incubated on a

laboratory bench, where the temperature was approximately 21-23 "C with a 16 hour

photoperiod, under a light intensity of 14 pmolm'2s-' provided by white fluorescent tubes.

After 10 days meristem-tips that were free of contamination and appeared viable, were

transferred to 100 mL glass plant tissue culture vessels (Sigma-Aldrich) containing 20

mL of the same medium. The vessels were placed inside a Percival growth chamber set

at 16 "C with a 16 hour photoperiod and light intensity of 14 ymolm-2s-' provided by

white fluorescent tubes. These meristems produced axillary shoots, which were

aseptically separated every two months and transferred to fresh medium (Fig. 4.2.b). To

determine if these in vitro plants were free of facultative microbial contamination, shoots

were aseptically macerated using a mortar and pestle and suspended in phosphate

buffered (0.01 M, pH 7.1) saline (0.85 %) solution (PBS) and 10 pL was plated onto NA

and PDA and incubated under laboratory conditions for two weeks. No bacterial or

fungal growth occurred. Individual axillary shoots were then rooted on hormone-free

Gamborg's B5 (Gamborg et al., 1968) solidified with phytagel (Sigma-Aldrich) in 77 mm

x 77 mm x 97 mm Magenta0 vessels (Sigma-Aldrich) placed inside a growth chamber

under the conditions described previously. After two months, the plants were used for

inoculation experiments (Fig. 4.2.c).

4.2.3. Fungal inoculations

The pathogenicity of several Pythium and Fusurium isolates recovered from

wasabi roots was evaluated using the tissue culture-derived hydroponically-grown plants

following the procedure described by Rodriguez and Punja (2007). All isolates were

grown on PDA for two weeks and 5 x 5 mm mycelial plugs were added to the hydroponic

Figure 4.2. Reproduction of wasabi vascular blackening symptoms. (a, b) Meristem-tip derived palhogen-Free tissue culture plants [ha[ were inocula~ed with Pectobacterium carotovorun~ subsp. car-otovorum (Pcc). (c) Whole plant eight weeks after inoculation. (d, e, f) Rhizomes in which plant roots were stabbed (d) and cut (e, f) and then inoculated with Pcc blackened vascular tissues can be seen. (g-k) Microscopic images of blackened vascular rhizome tissue depicting gels (G) and bac~eria (B) i n xylem parenchyma (h, j) and xylem tracheid (k) cells, following artificial inoculation.

solution. Symptoms of root rot or blackening were assessed after 12 weeks at 22 "C.

Fungi were reisolated from plant roots by taking three root sections of 1 cm length,

surface-sterilizing as described previously, and plating on WA with 300 mg/L

streptomycin sulfate. The experiments were performed twice with 10 plants per fungal

isolate.

4.1.4. Bacterial identification

A number of bacterial species, including Agrobacterium tumefaciens, Enterobacter

intermedius, Peetobacterium carotovorum subspecies carotovorum (Pcc), Pseudomonas

fluorescens, and Psez~dornonasfulva were recovered from wasabi rhizomes and identified

by BiologTM (Biolog Inc., U.S.A.) with a probability of 100, 100, 99, 96, and 97%,

respectively. Pathogenicity tests were conducted using these bacteria by infiltrating the

mesophyll space of a green cabbage leaf (purchased at a local store) with a 1 0 % f d m l of

bacteria suspension in PBS (Schaad, 1988). None of the isolates appeared pathogenic

(data not shown), since they did not produce a hypersensitive response or cause infection.

This test is invalid for Pcc (Schaad, 1988) and therefore the pectolytic enzyme activity of

was determined according to Schaad (1988). Potato Yukon Gold (purchased at a local

store) slices were inoculated with 1 o4 to 10' cfdrnL of Pcc suspended in PBS. Similarly,

wasabi rhizomes were sliced longitudinally, surface-sterilized, and inoculated. Tissues

were incubated in plastic containers with moist Filter paper under ambient conditions and

examined after 48 hr for evidence of decay. Pcc isolated was found to be highly virulent

and therefore were included in the following experiments.

For the KCN growth test (Goto and Matsumoto, 1987), Pcc was grown on NA and

then suspended in tryptic soy broth (Difco) to 0.5 McFarlands (Cowan and Steel, 1974;

Goto and Matsumoto, 1987). KCN broth or broth base was prepared according to Cowan

and Steel ( 1 974) and 3 mL aliquots were placed in 14 mm round bottom snap cap Falcon

tubes (BD Biosciences). One pL of suspended bacteria was added to KCN broth and to

the broth base and 1 mL of sterile paraffin was pipetted on top to provide anaerobic

treatments, while the aerobic controls were left open to the air. The cultures were

incubated at 30 OC for 2 weeks. Growth was compared to that of confirmed Pcc and Pcw

strains (provided by S. DeBoer, Canadian Food Inspection Agency, Table 4.1).

Table 4.1. Reference strains of Pectobacterium carotovorum ssp carotovorum (Pcc) and P. carotovorum subspecies wasabiae (Pcw) used in this study

Pcc 1 5 17 1 potato I U.S.A. (M. Powelson) 1 - I

Bacterium

PCC

In order to conduct a sequence comparison of the -510 bp IGS region, Pcc was

Strain

155

PCW

PCW

grown for 2 days in tryptic soy broth and the DNA extracted according to Lee et al.

(1 993). The previously published primers G l f: GAAGTCGTAACAAGGTA and L l r:

Host of origin

potato

9 1

92

CAAGGCATCCACCGT were used to amplify the 16s-23s rDNA IGS regions (Jensen

et a]., 1993) according to Fessehaie et al. (2002) except with an annealjng temperature of

Country of origin

Canada (B. Copeman)

wasabi

wasabi

55 "C and 31 cycles. PCR reactions were performed in a GeneAmp PCR System 9700.

GenBank accession number 16s / Small IGS

AF373 188

Fifteen pL of product was run on a 2 % agarose gel (BioShop) in a Tris-Acetate-EDTA

Japan (M. Goto (9 1 ))

Japan (M. Goto (92))

(TAE) buffer at 80 V for 1.5 hours. The smaller DNA band (-510 bp) was amplified

AF373 193

AF373 194

using the "band stab" method (Wilton et al., 1997) with the G l f and L l r primers. The

product was sequenced by Macrogen Inc. (Seoul, South Korea) and the DNA sequence

was compared to other Pcc and Pcw IGS sequences in GenBank.

4.2.5. Bacterial inoculations

To determine whether the isolated Pcc strain could induce rhizome vascular

blackening, the tissue-culture derived plants were used and the roots were trimmed and a

portion of the rhizome base was cut off with a scalpel. Rhizomes were soaked in PBS

containing bacteria ( lo8 cfu/mL) for 30 min and transferred to potting mix (steer manure,

Home Depot, Coquitlam, B.C.) that had been autoclaved three times. Controls included

plants which had not been wounded, and plants receiving buffer only. The plants were

grown for 8 weeks in a Percival growth chamber set at 22 "C, 90 % relative humidity with a

14 hr photoperiod and a light intensity of 10 pmolm-2s-' provided by cool-white fluorescent

lamps. The experiment was performed twice, with 10 plants per treatment.

To determine the effect of temperature on symptom development, rhizomes were

wounded by stabbing with a needle followed by Pcc inoculation as above and grown in

potting mix for 8 weeks. Plants were then transferred to 10, 22 or 27 "C and incubated for

an additional 8 weeks. The presence or absence of blackened vascular tissue was assessed

visually and microscopic examination of the tissues was performed as described

previously. The experiment was performed twice, with 10 plants per treatment. Bacterial

isolation was attempted from asymptomatic and symptomatic plants (with blackening) by

surface-sterilizing rhizome sections and plating onto NA. Recovered bacteria were

suspended in 9 mL PBS and 10 pL was pipetted and spread onto crystal violet pectate

(CVP) medium in a 60 mm petri dish (VWR) (Schaad, 1988). Colonies exhibiting

pectolytic activity were selected and picked with a sterile pipette tip and lysed in 5 pL of

sterile water by placing at 95 "C for 5 min. The PCR reaction was performed as described

above.

4.2.6. Bacterial plus fungal inoculations

To determine the potential interaction between Pythium spp. and Pcc, plants were

grown in a hydroponic system as described previously (Rodriguez and Punja, 2007).

Mycelial plugs of Pythium species were added to the hydroponic solution and plants were

grown as before, but at 18 "C. One week later, plants were removed and inoculated with

lo8 c fdmL Pcc bacteria by soaking them for 30 min. An additional set of plants was

wounded by stabbing and inoculated with the bacteria as described. All plants were grown

at 22 "C for 8 weeks and then harvested and presence of rhizome blackening was noted.

The experiment was performed twice with 10 plants per treatment.

4.2.7. Identification of inoculum sources

Potting mix used for vegetative propagation of wasabi was supplied by the grower,

and consisted of composted manure which was taken from unused bags. One gram of

each sample was placed in PBS and dilution-plated (up to 1 0 . ~ ) onto CVP plates. After 4

days, two colonies with pectolytic activity were selected from the 10-hilution plate and

amplified using PCR as described for those colonies obtained from blackened rhizome

tissue. Water samples were taken in July, 2005 and February, 2006 directly from the

irrigation heads at the greenhouse, and dilution plated up to 10.' on NA and PDA.

4.3. Results

4.3.1. Microbial isolation and microscopy

Wasabi plants with internal vascular blackening appeared visibly healthy when

viewed externally (Fig. 4.1 .a, b). From 60 plants sampled, various bacteria were isolated

from the epidermis and vascular regions at 100 and 80% frequency, respectively. The

cortex and pith were free from bacterial infection.

Pythium dissotocum Drechsler and Pythium intermedium de Bary were recovered

from blackened wasabi rhizome epidermal tissues (Rodriguez and Punja, 2007).

Fusurium sambticinum was occasionally recovered from these tissues. Fungi were not

recovered from the cortex, vascular region, or pith of the symptomatic rhizomes.

When symptomatic tissue was observed under the microscope, only the xylem

tracheid cells appeared black (Fig. 4.1 .e-g). Gels and tyloses were observed in the xylem

tracheids (Fig. 4.1 .g), which are indicative of systemic infection by microbes (Beckman,

2000). Bacteria were often visible in and between parenchyma cells that were in close

proximity to the blackened xylem tracheids (Fig. 4. I .h). Only one of the eight samples

examined microscopically did not contain visible bacteria despite being symptomatic.

Bacteria were always visible in epidermal tissues. Fungal mycelia were not observed

near the vascular tissue of the symptomatic rhizomes; however, they were visible on the

epidermis.

4.3.2. Fungal inoculations

The Pythiurn spp. caused significant root loss and were successfully reisolated from

root tissues. F. sumbucinum caused black lesions to develop in most plants where the

petioles were in contact with the hydroponic solution which contained the inoculum.

However, F. sambucinum did not infect root tissues as it had no negative effect on root

mass (data not shown), and was not reisolated from root tissue.

4.3.3. Bacterial identification

When Pcc was placed on potato and wasabi sections, tissue maceration occurred.

Potato tissue was degraded when bacteria were at a concentration 2 lo5 cfulml, while

wasabi maceration occurred when bacteria were at a concentration of 10' and lo8 cfulml.

Results of the KCN test confirmed that the bacterium originally isolated from wasabi was

Pcc and not Pew. The isolated bacterium and the confirmed Pcc control isolates (strains

155 and 157) grew in both aerobic and anaerobic conditions in the KCN broth, while the

confirmed Pew isolates (strains 92 and 94) only grew under aerobic conditions. This

identity was further confirmed when the IGS-PCR products were shown to consist of two

bands of -5 10 and -550 bp, which is typical of Pectobacterium carotovorum subspecies

carotovorum, subspecies atroseptica, and subspecies betavasculorum (Fessehaie et al.,

2002; Toth et al., 2001). The smaller IGS-PCR band (-510 bp) was sequenced, and

compared to other sequences in GenBank. It exhibited 99% homology with Pcc strain

El61 (GenBank accession number AF373189.1), and 96% homology with Pcw strain

ATCC 433 16 (GenBank accession number AF232679).

4.3.4. Bacterial inoculations

Pathogen-free wasabi plants from tissue culture (Fig. 4.2.a, b) when inoculated with

Pcc developed blackening in the rhizome vascular tissue (Fig. 4.2.d-f), despite appearing

outwardly healthy (Fig. 4.2.c). Of the wound-inoculated plants, 70 % developed black

streaks in the vascular tissue and three plants (15 %) exhibited soft-rot symptoms and

died. Only one plant which was a wound-control (by cutting) developed blackening of

the vascular tissue. None of the uninoculated control plants developed vascular

blackening. Pcc was successfully reisolated from the inoculated plants. Although

bacteria were reisolated on NA from the wound-control and control plants, pectolytic pits

were not formed on CVP, indicating they were not Pcc. These bacteria were not

identified further.

When 8 week-old wasabi plants were wound-inoculated with Pcc and later incubated

at 10, 22 or 27 OC, they developed rhizome vascular blackening or died due to soft-rot.

Rhizome vascular blackening developed in 50% or more of plants that were wounded by

cutting and then inoculated. Those that were stabbed and inoculated developed blackening

at a 25 to 50% frequency (Table 4.2). Soft-rot occurred in some of the inoculated plants

that were wounded and later incubated at 22 or 27 "C, irrespective of wounding method.

Soft-rot did not develop on those plants that were inoculated and later placed at 10 OC

(Table 4.2). The plants that were incubated at 27 OC developed blackening in the vascular

tissue of many petioles and along the leaf veins. Microscopic examination of rhizome

vascular blackening tissues revealed blackened xylem tracheids with bacteria in close

proximity either within or between xylem parenchyma cells (Fig. 4.1 .h, j).

4.3.5. Bacterial plus fungal inoculations

Plants that were wounded or inoculated with P. dissotocum or P. intermedium, and

subsequently inoculated with Pcc, developed rhizome vascular blackening in 9/20, 11/20,

and 8/20 plants, respectively. None of the control plants or the plants inoculated with

Pythium spp. alone were symptomatic. Pcc, P, dissotocum, and P. intermedium was

reisolated from all inoculated plants.

Table 4.2. Incidence of rhizome vascular blackening (rvb) and soft-rot (rot) in wasabi plants that were inoculated with Pcc and grown in potting mix at different temperatures.

I Incubation temperature ('C) after 8 weeks at 22 'C I

Note: - indicates asymptomatic plants, + indicates less than 25 % o f plants exhibited symptoms, ++ indicates between 25 and 50 % of plants were symptomatic, +++ indicates that 50 % or more of plants developed symptoms. Symptoms included rhizome vascular blackening (rvb) or soft-rot (rot) where Pcc was reisolated fiom infected tissue. Data represents two experiments that were combined (n=20).

Treatment

Cont ro l

W o u n d by s tabb ing

W o u n d by cu t t ing

W o u n d by s tabb ing + Pcc

W o u n d by cu t t ing + Pcc

4.3.6. Identification of inoculum source

Two bacterial colonies isolated from potting mix were subjected to colony-IGS-

PCR and exhibited the same banding pattern to that of the Pcc isolates from rhizomes

10

(Fig. 4.3). The -510 bp IGS region from these bacteria showed 100 % homology to the

rvb

++ +++

same region to Pcc that was obtained from the blackened rhizome vascular tissue in

rot

22

greenhouse-grown wasabi plants. Water samples obtained from the irrigation heads in

2 7

rvb

+++

the greenhouse did not yield bacteria or fungi.

rvb

+ ++ +++

rot

+ +

4.4. Discussion

rot

+ ++

Our observations indicate that rhizome vascular blackening of wasabi is not only

due to infection by Phomu wasabiue as reported in the literature. Microscopic

observations of symptomatic greenhouse-grown plants and inoculated tissue culture-

derived plants suggest that bacteria are also associated with rhizome vascular blackening.

Figure 4.3 Identification of Pectobacteriurn carotovorurn subspecies carotovorum using primer set L l r and G l f. Fragment sizes were -5 10 and -550 bp. Lanes M1 to M5 represent isolates which exhibited pectolytic activity after dilution-plating of potting mix onto crystal violet pectate medium (CVP). M I , M2 and M4 exhibited the same banding pattern as Pcc, the positive control isolate originally obtained from symptomatic wasabi, while M3 and M5 showed pectolytic activity on CVP but were not Pcc as demonstrated by PCR. Water was included as a negative control (H20).

In both instances, blackened xylem tracheids in the rhizome were surrounded by xylem

parenchyma tissue which contained bacteria both inter- and intra-cellularly. The

blackening symptom is likely a tissue response to microbial invasion, with accumulation

of oxidised phenolic compounds as a host defence response (Mace et al., 1972; Matsuki,

1996; Ploetz, 2005). P. wasabiae is reported to be a systemic pathogen (Lo and Wang,

2000b), but microscopic examination of symptomatic plants did not show mycelia

beyond the epidermal tissue and to date, P. wasabiae has not been recovered from wasabi

rhizomes grown in B.C. Previous isolation experiments in New Zealand showed that P.

wasabiae was recovered from symptomatic rhizomes in 9 out of 192 plants, indicating

that other agents could be involved in blackening symptoms (Broadhurst and Wright,

1998). This was also suggested by Goto and Matsumoto (1986), who found that only 14

out of 149 plants with vascular rhizome blackening symptoms yielded P. wasabiae.

This is the first report describing inoculation studies conducted using bacteria

isolated from wasabi, although previous studies from Japan (Goto and Matsumoto,

1986a) and New Zealand (Broadhurst and Wright, 1998) reported the presence of Pcc

and Erwinia spp. (possibly syn. Pectobacterium spp.) and Pseudomonas spp. in

symptomatic tissue. We also recovered Pseudomonas spp. in our study, as well as

Agrobacterium tumefaciens, Enterobacter intermedius, but pathogenicity tests on

cabbage leaves showed that none of them were pathogenic (data not shown). Inoculation

studies conducted on pathogen-free tissue culture-derived wasabi plants indicated that

Pcc is a likely cause of rhizome vascular blackening. Inoculated plants exhibited

identical symptoms to symptomatic greenhouse-grown plants, both at the macroscopic

and microscopic levels, and Pcc was successfully reisolated from inoculated plants. The

involvement of Pcc in this disease syndrome may explain the lack of success that

fungicide spray programs have had in Asia in reducing the incidence of rhizome

blackening (Lo et al., 2002), presumed to be due to P. wasabiae.

In addition to rhizome vascular blackening, soft-rot was occasionally observed

when temperatures were increased (plants grown at 22 and 27 "C) in potting mix but not

under hydroponic conditions. In potting mix, the moisture content was high to prevent

plant wilting and this may have caused localised regions of depleted oxygen which would

have been optimal for Pcc, an opportunistic pathogen and a facultative anaerobe (De

Boer, 2003). Perombelon (2002), in studies on potatoes, showed that Pcc, although often

associated with asymptomatic plants, can induce rot once conditions become favourable

or host resistance is impaired. Favourable conditions include temperatures around 27 "C

and sufficient moisture (De Boer, 2003; Perombelon, 2002; Smadja et al., 2004).

Impairment of host resistance often occurred when localised soil conditions become

anaerobic and oxygen-dependent production of phytoalexins, phenolics, free radicals, and

cell wall lignin are hindered (Perombelon, 2002). Bacteria can release large amounts of

cell-wall degrading enzymes to induce soft-rot (Barnard and Salmond, 2007), and soft-rot

symptoms were observed on wasabi rhizomes and foliage during the warm summer

months on plants grown in a commercial greenhouse.

A primary inoculum source is likely the potting mix as results showed that the

growth medium used by growers to root offshoots during vegetative propagation

contained Pcc. Experiments in which plants were wounded by cutting followed by

inoculation had higher vascular blackening after incubation than those that were wounded

by stabbing. Vegetative propagation likely spreads Pcc since vegetative offshoots are cut

from the main rhizome using knives, providing wounds for colonization by bacteria. In

addition to the potting mix, vegetatively propagated plants, even asymptomatic plants,

may contain Pcc (Goto and Matsumoto, 1986a). This may explain why the

recommended use of only two successive generations of vegetatively propagated plants

has been reported to decrease the frequency of symptom development (Adachi, 1987;

Chadwick, 1993). In addition to infection via vegetative cuttings, Pcc may be infecting

wasabi through entry points created by infection with Pythium spp. This explains the

occurrence of rhizome vascular blackening in commercial plants initiated from seed, and

indicates that in order to decrease the incidence of rhizome vascular blackening, damage

by Pythium spp. must be reduced.

The use of pathogen-free plants derived from meristem-tip culture may be one

avenue to reduce rhizome blackening, provided there is no inoculum being introduced

from the potting mix. Tissue cultured plants would also not incur wounding, thus

reducing the amount of direct entry into the rhizome vascular tissue by Pcc and likely

reducing the extent of blackening. These should be tested to confirm the absence of

microbes within tissues by plating onto media or using serological test kits prior to use

(Falkiner, 1997).

The results from this study show that Pcc is a probable cause of rhizome vascular

blackening of wasabi and that symptoms occur when wounded roots or cut rhizomes are

colonized by bacteria.

CHAPTER 5: GENERAL DISCUSSION AND CONCLUSIONS

5.1. Tissue culture of wasabi

5.1.1. Pathogen-free material

This research successfully determined a micropropagation technique which can be

used to establish pathogen-free wasabi plants (cv. Daruma). This is different from the

protocol recently outlined by Hung et al. (2006) in which larger explants containing the

axillary buds were used to establish the aseptic culture and antibiotics were used (5OmgIL

of streptomycin) to suppress bacterial growth. Under these conditions, it is probable that

bacterial pathogens have not been excluded, as bacterial contaminants which may remain

latent within the tissue and do not necessarily produce visible growth on tissue culture

media. Latent infection may only be detected using 'indexing media' (Sigma-Aldrich) or

alternatively using broad-spectrum media or commercial serological test kits for specific

plant pathogenic bacteria (Leifert C., 1992).

5.1.2. Acclimatisation

Approximately 1500 rooted plants have been obtained using the micropropagation

protocol described here. In March 2007, 60 rooted plants were taken from tissue culture

and transplanted directly into the greenhouse. Despite the absence of an acclimatisation

period, all plants survived and continued to grow and produced new leaves. Therefore,

an additional stage of acclimatisation appears to be unnecessary in early spring, which is

highly desirable from a commercial standpoint due to reduced labour and material costs.

However, direct transfer from tissue culture may not be possible during the hot summer

months or very cold winter months, restricting the direct transfer of tissue culture

material to the spring and fall seasons only.

The apparent lack for the need of an acclimatisation stage is probably a function of

two factors. Firstly, using the protocol described here, in vitro plants were grown under

cool conditions at 18 "C. When tissue culture plants are grown under warm conditions,

typically 25 * 3 "C (Pollard and Walker, 1990), plants often develop stomata which

respond poorly to stimuli that would normally induce closure. In addition, tissue-cultured

plants often have poor epicuticular wax development (Pollard and Walker, 1990). The

placement of culture vessels under cooler temperatures is sometimes used to acclimatise

plants prior to removal from in vitro culture (Maene and Debergh, 1986). The cooler

temperature lowers the humidity level around the leaves within the culture vessel and

these leaves have stomata that are more responsive and may have improved cuticular wax

development (Pollard and Walker, 1990). Secondly, wasabi is grown under high

humidity levels within the greenhouse. Placing plants within a humid environment after

removal from in vitro culture is common practice for acclimatising plantlets (Scott,

1986), which decreases the water loss experienced by the plant due to poor stomata1

response and poor epicuticular wax development (Pollard and Walker, 1990).

5.1.3. Commercialisation

While the meristem-tip and axillary bud multiplication protocol established here has

been successful, the original purpose was for use on a commercial scale so that growers

would have access to large quantities of pathogen-free planting material. In order to

achieve this, the protocol must be commercialised. This involves increasing the volume

of cultures and decreasing the production cost, minimising cost per unit production.

Strategies for material and labour cost reduction should be investigated to achieve this.

All chemicals used in the production of tissue culture plants, except for sucrose,

were tissue culture grade and therefore more expensive than laboratory grade chemicals.

By changing the grade of chemicals used, commercial costs may be decreased; however,

the efficacy would have to be compared to tissue culture grade chemicals. The long-term

goal of this procedure was always in mind during its development and therefore, only the

effects of relatively inexpensive hormones were investigated. BA was determined to be

the most effective hormone at inducing shoot proliferation, and is one of the most

inexpensive cytokinins available (Sigma-Aldrich).

In addition to decreasing material costs, the removal of the rooting stage should be

examined. Costs could be significantly reduced if unrooted shoots are directly

transplanted into the greenhouse as labour, material costs, and overhead costs are all

reduced. Although highly desired by commercial producers of tissue cultured plants, this

technique is limited to particular plants and growing conditions (Hu and Wang, 1983;

Pollard and Walker, 1990). However, given the high humidity levels in the greenhouse

and the ease at which axillary shoots from established plants are routinely propagated, it

may be successful.

In order to accurately schedule availability of pathogen-free planting material,

commercial producers should determine whether the number of axillary shoots obtained

from each division decreases over time according to age of original meristem-tip explant,

and if so how many division cycles can be obtained from each meristem-tip. Repeated

cell division may cause the accumulation of genetic aberrations (Pollard and Walker,

1990). However, this is a problem more closely associated with plants that are derived

through callus proliferation, while axillary shoot proliferation is relatively resistant to

genetic aberrations (Morel, 1960a; Sagawa, 1966). The data obtained suggests that the

number of plants obtained from an axillary shoot stabilised at 3.4 to 3.8 shoots per

axillary shoot division, over a two year period (or 12 axillary shoot division cycles).

Commercial propagators need to determine whether this decreases and the point at which

it decreases, in order to accurately plan when new aseptic cultures should be established.

5.2. Low root mass in wasabi

5.2.1. Pythium spp. control

Following the isolation, identification and confirmation of pathogenicitiy of

Pythium spp., wasabi growers implemented a fungicide spray control program to target

this pathogen. Following the initiation of this program, plant growth rate and root mass

have increased significantly. Additional studies may be required in order to secure

registration for fungicide use on wasabi, as current use is based on fungicides used in

other Brasssicaceae crops.

In order to produce wasabi plants that are certified organic, the efficacy of

biocontrol agents should be investigated. Products such as MycostopTM, Rootshield

Drench WPTM, and Prestop WPTM (Table 1.1) could be investigated to determine their

effectiveness against P. dissotocum and P. intermedium in wasabi. MycostopTM and

Rootshield Drench WPTM contain Streptomyces griseoviridis and Trichoderma

harzianum as the active ingredient, respectively. They are currently registered in Canada

for use on many plant species, including many members of the Brassicaceae family

(Kabaluk and Gazdik, 2005); therefore, if effectiveness is established, registration for use

on wasabi may be easier. Prestop WPTM contains the microorganism Gliocladium

catenulatum which is currently awaiting registration for use in greenhouse crops by the

Pest Management Regulatory Agency. As it represents an alternative biological agent, it

may display significant differences in controlling P. dissotocum and P. intermedium in

wasabi.

5.3. Rhizome vascular blackening

5.3.1. Pcc isolation variation and the possible involvement of additional microbes

Research conducted indicates that Pcc can cause rhizome vascular blackening.

However, isolation studies from Japan reported 30% of symptomatic plants contained

Pcc (Goto and Matsumoto, 1986a), while only 18% of symptomatic plants sampled in

New Zealand contained Erwinia spp. (possibly Pcc)(Broadhurst and Wright, 1998). This

variation could be due to differences in surface-sterilisation, conducting studies when

temperatures are low, or rhizome vascular blackening caused by infection by other

microorganisms.

Pcc may occur at low levels when temperatures are low because growth is often

depressed under these conditions, until optimal conditions are present and bacteria

increase in number. One of the parameters defining optimal conditions is ambient

temperature, which is 27 "C for Pcc (PCrombelon, 2002; Smadja et al., 2004). To test

whether low isolation levels of Pcc corresponds to temperature an epidemiological study

would need to be conducted. Symptomatic plants may be sampled continuously, perhaps

during each harvest. Isolation procedures should by followed as outlined here, so as not

to over-sterilise tissue, and isolation data correlated to temperature within the

greenhouses.

As blackened vascular tissue is a symptom of a plant's response to systemic

infection (Mace et al., 1972; Matsuki, 1996; Ploetz, 2005), other microbes in addition to

Pcc, may cause rhizome vascular blackening. Pcc secretes pectolytic enzymes during

plant attack and other bacteria which secrete these enzymes could elicit rhizome vascular

blackening in wasabi similar to the response seen when wasabi is infected with Pcc.

Fungi that are systemic pathogens may too elicit a similar response by the plant to

infection. Both pectolytic enzyme-secreting bacteria, namely Pseudomonas flourescens

Migula and systemic fungi, namely P. wasabiae and L. maculans, have been previously

isolated from symptomatic tissue in Japan and New Zealand (Goto and Matsumoto,

l986a; Broadhurst and Wright, 1998). Additional microbe isolation studies should be

performed in B.C. from greenhouse-grown plant material in conjunction with the

epidemiological studies mentioned previously. In particular, isolation of pectolytic-

enzyme-producing bacteria should be performed by surface-sterilizing the symptomatic

tissue and plating directly on CVP. Isolation of P. wasabiae and L. maculans should also

be focused on. Inoculation studies could then be done with isolated microbes under

similar conditions described here.

Isolation of Phoma spp. was successful in May 2007, when black lesions on the

leaves, measuring up to 1 cm across, developed on greenhouse-grown plants in

Aldergrove, B.C. Discolouration in the rhizome tissue is currently being monitored in

these plants; however, an increase in rhizome vascular blackening has not been reported

by the grower. Inoculation experiments are currently being conducted to determine the

isolates' ability to cause leaf lesions, and to examine the possibility of induction of

rhizome vascular blackening in the plant.

5.3.2. Defining symptoms and causal agents

Since the isolation of P. wasabiae in 1933 by Yogoki, it has traditionally been

regarded as the causal agent of rhizome blackening (reviewed in Chadwick, 1993).

However, isolation studies report highly variable rates of Phoma spp. associated with

blackened rhizome tissue and inoculation studies have not always been performed (Goto

and Matsumoto, l986a; Broadhurst and Wright, 1998; Rodriguez and Punja, 2006; 2007).

Also, the types of rhizome tissues that are blackened are rarely distinguished. The

prevailing belief that P. wasabiae is the causal agent of any rhizome blackening was

recently demonstrated in a Tasmanian study (Sparrow, 2006), where blackening in the

rhizome pith was attributed to P. wasabiae. Yet, isolation experiments were not

performed and the symptoms shown in photographs are quite unlike the original report by

Yokogi in 1933. The lack of inoculation studies and the combining of all rhizome

blackening symptoms together has led to failure in controlling rhizome blackening as

reported by Lo et a1 (2002) and has not only resulted in financial loss by growers

(Chadwick, 1993), but exacerbated the loss as growers unsuccessfully apply fungicides to

their crop (Lo and Wang, 2001).

Blackening in the rhizome pith, cortex or epidermis (Lo and Wang, 2000b; Adachi,

1987; Chadwick, 1993; Sparrow, 2006; Goto and Matsumoto, 1986a; Lo and Wang,

2000a; 2001), and blackening of petioles or leaf veins (Lo and Wang, 2001; Chadwick,

1993; Adachi, 1987) need to be separated. Studies here demonstrated that Pcc can cause

blackening of the rhizome vascular tissue, but did not appear to cause any other

symptoms. However, wounding and temperature stress did appear to cause tissue

discolouration, while nutrient deficiency may also induce tissue blackening. This

blackening was not the same as that induced by inoculation with Pcc. Instead, it was a

non-specific tissue discolouration, which is expected when phenolic production by a plant

is in response to an abiotic stress (Matsuki, 1996).

During tissue culture experiments, shoots with small rhizomes sometimes

developed blackening in the pith. Plating of this tissue after maceration showed that it

was free from microbe infection. In addition, shoots usually blackened at the base where

they were aseptically cut from the mother-shoot; and experiments with callus often

produced large amounts of phenolic compounds. These reactions are most likely due to

wounding, and therefore some of the non-specific tissue blackening in the rhizome,

observed in greenhouse grown plants, may also be due to wounding.

In addition to wounding, high temperature also caused tissue discolouration.

During Pcc inoculation experiments conducted here, leaf veins and petioles turned black

at 27 "C, irrespective of whether they had been inoculated. Interestingly, reports of P.

wasabiae infection in wasabi often describe blackened leaf veins and petioles as a

symptom of infection (reviewed in Chadwick, 1993).

Calcium and boron deficiencies are also reported to cause rhizome blackening (Lo

and Wang, 2001). Although the exact tissue was not specified in the study, boron

deficiency occurred in plants grown in Richmond, B.C. and caused rhizome pith and

vascular tissue to discolour.

These examples demonstrate the need for symptom differentiation in wasabi. This

together with extensive isolation and inoculation studies, will help provide the

information needed to successfully decrease all types of tissue discolouration.

5.4. The interaction between Pcc and wasabi

A question that arises when studying the interaction of wasabi and Pcc is: why does

Pcc rarely induce soft-rot in wasabi? When inoculated plants were placed under

controlled optimal conditions for the bacteria, rates of soft-rot were surprisingly low.

And, bioassay inoculations with Pcc on wasabi versus potato also revealed that wasabi

appears to be more resistant to soft-rot by Pcc. These observations may by due to one or

both of the following reasons; wasabi is somewhat resistant to soft-rot caused by Pcc, or

Pcc is not the necrotroph that we traditionally have regarded it to be.

The rhizome vascular blackening that is problematic to wasabi growers appears to

be due to phenolic production by the plant to reduce infection by Pcc. Under these

conditions, phenolic compounds are oxidized within the plant as a mechanism of vascular

defence and are often visualised as discolouration in the vascular tissue, the formation of

gels and tyloses and increased lignification of cell walls (reviewed in Beckman, 2000).

Under controlled inoculation conditions, soft-rot often did not develop, therefore the

pathogen Pcc is largely ineffective in causing disease in wasabi. The black

discolouration within the rhizome vascular tissue indicates that the phenolic compounds

produced may allow wasabi to withstand the bacterial attack and possibly explain the

tolerance level of wasabi to soft-rot even under optimal temperatures for the bacteria.

Therefore, studying the plant's response to attack by Pcc may reveal a key feature which

could later be used in Pcc soft-rot control. Further research may be conducted to study

which plant genes are involved in the response to Pcc infection. In particular, it would be

interesting to distinguish if wasabi up-regulates genes apart from those that are generally

involved in pathogen control, when infected plants are placed under optimal conditions

(27 "C) for bacterial growth (Smadja et al., 2004). This could be determined by

subtractive hybridization, where the plant genes that are up-regulated during infection

under optimal conditions for the bacteria are "subtracted" from those that are up-

regulated during infection at lower temperatures (10 "C), when bacterial infection is

latent.

Pcc, along with other soft-rot erwiniae, have traditionally been regarded as strict

necrotrophs, due to their ability to cause rapid maceration of tissue and subsequent plant

death, by releasing large amounts of pectolytic enzymes in a highly regulated manner

(Toth and Birch, 2005). However, research where pathologists are attempting to

determine how soft-rot erwiniae interact with the plant host is challenging this

necrotrophic ideology. Studies involving soft-rot erwiniae are exposing components of

these pathogens which are similar to biotrophic phytobacteria. For example, transcription

factors in the plant which are elicited by soft-rot erwiniae, appear to elicit not only the

expected jasmonic acidjethylene-dependent pathway, but sometimes an exclusively

salicylic-dependent response, usually reserved for response to biotrophs. Soft-rot

erwiniae also possess type I11 secretion systems, some aspects of which down-regulates a

plant's response to infection (Toth and Birch, 2005); this too is typical of a plant's

response to a biotroph. Further studies may reveal more information on how Pcc remains

latent within a plant, and thus help explain the observed lack of soft-rot under optimal

conditions.

5.5. Conclusions

A micropropagation technique was established where pathogen-free plant material

could be repeatedly divided by means of axillary shoot propagation, allowing a

multiplication rate of 3.8*0.13 per division of axillary shoot. Further studies need to be

conducted in order to establish if this technique is commercially cost-effective. P.

dissotocum and P. intermedium were determined to be the causal agents of reduced root

mass and suppressed growth rate of commercially-produced greenhouse-grown plants.

Finally, research showed that Pcc could induce rhizome vascular blackening; however,

further research should be conducted to determine the distribution of the pathogen and

the ability of other systemic pathogens to cause the same symptom. Accurately

describing the various rhizome blackening symptoms and linking these to either

particular abiotic or biotic stresses would significantly aid the growers in decreasing the

occurrence and economic losses associated with this problem.

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