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Effect of Plasmodiophora brassicae resting spore concentration and
crop rotation on growth of clubroot-resistant crops
By
Jill A. Dalton
A Thesis
Presented to
The University of Guelph
In partial fulfillment of requirements
for the degree of
Master of Science
in
Plant Agriculture
Guelph, Ontario
© Jill A. Dalton, January 2016
ABSTRACT
Effect of Plasmodiophora brassicae resting spore concentration and
crop rotation on growth of clubroot-resistant crops
Jill Allison Dalton Advisors:
University of Guelph Dr. Mary Ruth McDonald
Dr. Bruce D. Gossen
Clubroot caused by Plasmodiophora brassicae Woronin is an important threat to production of
canola (Brassica napus L.) and Brassica vegetables in Canada and worldwide. To help
understand the role of crop rotation in integrated clubroot management, this research examined
the pattern of decline in resting spores and the influence of spore concentration on growth and
development of clubroot-resistant crops. It was found that a portion of the resting spore
population is long-lived, but most (>99%) spores survive for only 1–2 years. Higher
concentrations of resting spores resulted in reduced plant growth and delayed development in
resistant canola and napa cabbage. However, the growth response was inconsistent across studies
and repetitions, and may be influenced by other factors such as soil type and crop species. For
canola growers, a ≥ 2-year break from canola, in combination with a clubroot-resistant cultivar, is
recommended as a clubroot management strategy wherever clubroot is found.
iii ACKNOWLEDGEMENTS
I would like to express my sincere gratitude to my co-advisors Dr. Mary Ruth McDonald
and Dr. Bruce D. Gossen for providing me with this opportunity and for their guidance, patience,
insight and support throughout this process. Thank you for helping me to strengthen my skills in
scientific analysis, writing and public speaking. I would also like to recognize the advice and
encouragement of committee members Dr. Katerina Serlemitsos Jordan and Dr. Sean Westerveld.
Thank you all for your time and expertise.
I appreciate the financial support provided by the Canola Council of Canada, Agriculture
and Agri-Food Canada through the Canola Science Cluster of Growing Forward 2, and the
University of Guelph Department of Plant Agriculture.
Thank you to Dr. Abhi Deora, Dr. Fadi Al-Daoud, Travis Cranmer and many other lab
colleagues for technical advice, support and encouragement. I am very grateful to Kevin Vander
Kooi, Laura Riches, Shawn Janse, Misko Mitrovic, Dennis Van Dyk and summer staff at the
Muck Crops Research Station for all of your help with the 2014 and 2015 field trials. I would
also like to acknowledge Godfrey Chu, Ken Bassendowski, Denis Pageau and others at AAFC
and University of Guelph for providing materials and support for the crop rotation studies.
I am very grateful to my parents for encouraging me to continue learning, to always
do my best and to believe in myself. I would like to say a special thank you to my sister for
teaching me to think critically. I appreciate the tremendous amount of support I have received
from my friends and family, especially Jack & Wilda Mardlin, Elizabeth Fraser, Gabe Sawhney,
Ashleigh Dalton, Laura Dalton, Martin Dalton and, finally, John Jeffrey Mardlin, for truly
going above and beyond a reasonable amount of support.
iv TABLE OF CONTENTS
CHAPTER ONE – LITERATURE REVIEW ......................................................................................................... 1 1.1 Brassica crops and their relatives ...................................................................................................... 1 1.1.1 Canola and other oilseed Brassica crops ................................................................................ 1 1.1.2 Brassica vegetables .......................................................................................................................... 2 1.1.3 History ....................................................................................................................................................... 4 1.1.3 Economic importance of Brassica crops in Canada ........................................................... 4 1.1.4 Pests and diseases in Brassica crops ........................................................................................ 5
1.2 Clubroot of Brassica ................................................................................................................................. 7 1.2.1 Host range and history of clubroot ........................................................................................... 7 1.2.2 Causal agent and taxonomy .......................................................................................................... 8 1.2.3 Disease cycle ....................................................................................................................................... 9 1.2.4 Pathotypes ........................................................................................................................................ 12 1.2.5 Incompatible Interactions .......................................................................................................... 13
1.3 Factors influencing infection and development ........................................................................ 14 1.3.1 Soil moisture .................................................................................................................................... 15 1.3.2 Temperature .................................................................................................................................... 15 1.3.3 Soil pH ................................................................................................................................................. 16 1.3.4 Resting spore concentration ..................................................................................................... 17
1.4 Disease management ............................................................................................................................ 17 1.4.1 Cultural control ............................................................................................................................... 17 1.4.2 Synthetic fungicides and surfactants ..................................................................................... 19 1.4.3 Biological control and biofungicides ..................................................................................... 21 1.4.4 Host plant resistance .................................................................................................................... 23 1.4.5 Metabolic cost of resistance ...................................................................................................... 25
1.5 Summary and objectives ..................................................................................................................... 26 CHAPTER TWO -‐ EFFECT OF RESTING SPORE CONCENTRATION ON GROWTH OF CLUBROOT-‐RESISTANT BRASSICA CROPS ................................................................................................. 28 2.1 Introduction .............................................................................................................................................. 28 2.2 Materials and Methods ......................................................................................................................... 33 2.2.1 Controlled environment study – canola ............................................................................... 33 2.2.2 Large pot studies – outdoors .................................................................................................... 35 2.2.3 Controlled environment study of canola, cabbage and napa cabbage .................... 36 2.2.4 Field trials ......................................................................................................................................... 37 2.2.5 Controlled environment pH study .......................................................................................... 39 2.2.6 Statistical analysis ......................................................................................................................... 40
2.3 Results ......................................................................................................................................................... 41 2.3.1 Controlled environment study ................................................................................................. 41 2.3.2 Large pot studies – outdoors .................................................................................................... 44 2.3.3 Controlled environment study – canola, napa cabbage and cabbage ..................... 45 2.3.4 Field trials ......................................................................................................................................... 46 2.3.5 pH study ............................................................................................................................................. 52
2.4 Discussion .................................................................................................................................................. 54
v CHAPTER THREE -‐ DECLINE IN RESTING SPORES AND EFFECT OF CROP ROTATION FOLLOWING A SUSCEPTIBLE CROP ............................................................................................................... 62 3.1 Introduction .............................................................................................................................................. 62 3.2 Materials and Methods ......................................................................................................................... 67 3.2.1 Decline in resting spores over time following susceptible canola ............................ 67 3.2.2 Effect of resistant canola on the concentration of resting spores in soil ............... 70 3.2.3 Controlled environment study – crop rotation and spore concentration ............. 71 3.2.4 Statistical analysis ......................................................................................................................... 73
3.3 Results ......................................................................................................................................................... 74 3.3.1 Decline in resting spores over time following susceptible canola ............................ 74 3.3.2 Effect of resistant canola on the concentration of resting spores in soil ............... 75 3.3.3 Weather .............................................................................................................................................. 77 3.3.4 Controlled environment – crop rotation and spore concentration .......................... 78
3.4 Discussion .................................................................................................................................................. 81 CHAPTER FOUR -‐ GENERAL DISCUSSION ................................................................................................... 91 LITERATURE CITED .............................................................................................................................................. 97 APPENDIX 1: SUPPLEMENTARY TABLES FOR CHAPTER TWO ...................................................... 116 APPENDIX 2: SUPPLEMENTARY TABLES FOR CHAPTER THREE ................................................. 172
vi LIST OF TABLES
Table 2.1 Soil properties of field trial sites at the Muck Crops Research Station at Holland Marsh, Ontario in 2014 and 2015. ........................................................................................................................... 38
Table 2.2 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible canola inoculated with 1 x 106 spores ml-1 under controlled conditions. ................................................... 42
Table 2.3 Clubroot incidence (CI) and severity (disease severity index, DSI) in canola breeding line ACS-N39 (susceptible check) and resistant cultivar 45H29 in an outdoor trial using large pots near Bradford, ON, 2014 and 2015. ..................................................................................... 44
Table 2.4 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible canola, napa cabbage and cabbage inoculated with 1 x 106 spores ml-1 P. brassicae, under controlled conditions. .................................................................................................................................... 45
Table 2.5 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible canola and napa cabbage grown in field soil at Muck Crops Research Station, 2014. ......................... 46
Table 2.6 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible and resistant canola cultivars, grown in field soil at sites with high, lower and no measurable concentration of resting spores at the Muck Crops Research Station, 2015. .............................. 47
Table 3.1 Quantification of resting spore concentration using qPCR after crop rotation treatments including fallow (F), resistant (R) and susceptible canola (S), sampled in spring and fall of 2014 and spring of 2015 at Normandin, Quebec. ............................................................................. 777
Table 3.2 Comparison of soil locations used in crop rotation and inoculum trials, under controlled environment in Guelph, Ontario, 2014. ............................................................................. 71
Table 3.3 Quantification of resting spore concentration using qPCR after crop rotation treatments including fallow (F), resistant (R) and susceptible canola (S), sampled in spring and fall of 2014 and spring of 2015 at Normandin, Quebec.. ............................................................................... 76
Table 3.4 Clubroot incidence (CI) and severity (disease severity index, DSI) in canola breeding line ACS-N39 (susceptible check) and the resistant cultivar 45H29, inoculated with P. brassicae and grown under controlled conditions .......................................................................... 78
Table 3.5 Effect of crop rotation and spore concentration on biomass (dry shoot weight, g) of clubroot-resistant canola, under controlled conditions in field soil from Elora, ON, and Scott, SK, 2014. ............................................................................................................................................... 81
vii LIST OF FIGURES
Figure 2.1 Plant height of clubroot-resistant canola at 6 weeks after inoculation with increasing concentrations of Plasmodiophora brassicae resting spores under controlled conditions (two repetitions, n=8, P = 0.005). ........................................................................................................................ 43
Figure 2.2 Plant maturity at harvest in canola grown in controlled conditions in response to increasing concentration of Plasmodiophora brassicae resting spores (P = 0.007) in the first repetition. ........................................................................................................................................................... 43
Figure 2.3 Plant height of three resistant canola at the Muck Crops Research Station, Ontario, at sites with high, lower and undetectable resting spore concentrations, 2014 and 2015. .......... 49
Figure 2.4 Plant height at 8 weeks after seeding (WAS), biomass at harvest and maturity at 8 WAS for resistant canola cultivars grown in field soil with 7 x 105 spores g-1 soil (Low) and 7 x 106 spores g-1 soil (High) at Muck Crops Research Station, 2014. Bars with the same letter do not differ at P = 0.05 based on Tukey’s multiple means comparison test. ................ 50
Figure 2.5 Plant height and maturity at 8 weeks after seeding (WAS), biomass at harvest for resistant canola cultivars grown in field soil with <1000 spores g-1 soil (BDL), 3 x 106 spores g-1 soil (Low) and 1 x 107 spores g-1 soil (High) at Muck Crops Research Station, 2015. Bars with the same letter do not differ at P = 0.05 based on Tukey’s multiple means comparison test. ....................................................................................................................................................................... 50
Figure 2.6 Leaf length and biomass of napa cabbage at sites with 7 x 105 spores g-1 soil (Low) and 7 x 106 spores g-1 soil (High) in field trials at Muck Crops Research Station, Ontario, 2014. Bars with the same letter do not differ at P = 0.05 based on Tukey’s multiple means comparison test. ............................................................................................................................................... 51
Figure 2.7 Leaf length and biomass of napa cabbage at sites with <1000 spores g-1 soil (BDL), 3 x 106 spores g-1 soil (Low) and 1 x 107 spores g-1 soil (High) in field trials at the Muck Crops Research Station, Holland Marsh, Ontario, 2015. Bars with the same letter do not differ at P = 0.05 based on Tukey’s multiple means comparison test. .......................................................... 52
Figure 2.8 Clubroot incidence (CI) in susceptible canola grown under controlled conditions at a range of pH (5.5 to 7.5) and concentrations of resting spores of P. brassicae at 22 days after inoculation (DAI). .......................................................................................................................................... 53
Figure 3.1 Decline in resting spore concentrations of Plasmodiophora brassicae over time following susceptible canola at Normandin, Quebec, assessed in 2014. ..................................... 75
Figure 3.2 Monthly rainfall (bars) and mean monthly air temperature (line) at Normandin, Quebec, for May to September, 2007-2013. .......................................................................................... 78
Figure 3.3 Effect of crop rotation and inoculation with Plasmodiophora brassicae on plant height (area under growth stairs, AUGS) of clubroot-resistant canola, under controlled conditions in field soil from Elora, ON, and Scott, SK in 2014. .......................................................................... 80
1 CHAPTER ONE – LITERATURE REVIEW
1.1 Brassica crops and their relatives 1.1.1 Canola and other oilseed Brassica crops
The family Brassicaceae (formerly Cruciferae) is composed of angiosperms in the order
Brassicales, distinguishable by their corolla, made up of four flower petals shaped like a cross
(Cartea et al., 2011). Plants in the Brassicaceae have six stamens, and the four inner stamens are
shorter than the two outer stamens (Franzke et al., 2011). The family contains 338 genera and
3709 species, including many important crops used for a variety of purposes including vegetables
for human consumption, oils for cooking and industrial use, condiments and livestock fodder (Al-
Shehbaz et al., 2006). There are three important diploid Brassica species, B. oleracea L., B. rapa
L. and B. nigra L. (Dixon, 2007). Three amphidiploid species, B. napus L., B. carinata L. and
B. juncea L., have emerged from crosses of diploid species (U, 1935). Arabidopsis thaliana (L.)
Heynh, which is a member of the family Brassicaeae, is an important model organism used in the
study of angiosperms (Arabidopsis Genome Initiative, 2000; Al-Shehbaz et al., 2006).
Canola is a quality standard for oilseed rape developed from B. napus (rapeseed) in the
1970s by Canadian plant breeders from Agriculture and Agri-Food Canada and the University of
Manitoba. By definition, canola produces seed with oil that contains less than 2% erucic acid and
less than 30 μmol g-1 of glucosinolates (Canola Council of Canada, 2011). Other names for
oilseeds with a comparable nutritional profile include Double Zero (00) Rapeseed and LEAR
(Low Erucic Acid Rapeseed) oil (Health Canada, 2003). The majority of canola cultivars grown
in Canada are B. napus, but canola-quality cultivars of B. rapa and B. juncea have also been
released (Canola Council of Canada, 2011).
2 Canola may be fall-seeded or spring-seeded, but almost all of the canola grown in western
Canada is spring-seeded. Like all Brassica crops, canola plants emerge quickly, with a pair of
cotyledons developing 4-7 days after seeding (Lamb, 1989). After the first pair of true leaves
develop, a rosette of broader leaves is formed. At this point, the fall-seeded winter annual enter a
dormancy state, whereas the spring-seeded plants continue growth. The next stage of growth is
stem elongation, followed by flower bud production (Hayward, 2012). Flowers are produced in
racemes, with one raceme developing at the very top of the plant, followed by axillary racemes.
The corolla consists of four yellow petals, which open beginning at the base of each raceme.
Petals drop after fertilization, and a cylinder-shaped pod is formed, called a silique. Upon
maturation, this pod contains between 15 and 40 seeds. Leaf senescence begins as the pods
mature, starting with the leaves at the base of the plant (Lamb, 1989).
Brassica napus and B. rapa make up a large majority of the global Brassica production
(Hayward, 2012). B. rapa, one of the original crossing parents of B. napus, contains the most
cold-tolerant cultivars of Brassica oilseeds. These are popular in areas where early maturity is
important due to the colder climate, including parts of Canada, Sweden and Finland. In India,
B. juncea makes up 80% of the oilseed production because of its heat and drought tolerance
(Hayward, 2012). In East Africa, B. carinata is cultivated and has recently been developed into a
biofuel (Abukutsa and Onyango, 2005; Hayward, 2012).
1.1.2 Brassica vegetables
Brassica vegetables are also known as cruciferous vegetables, cole crops, cabbage family
vegetables or mustard family vegetables. They are morphologically diverse, and are cultivated for
various edible parts of the plant, including roots, stems, leaves, seeds, flowers and sprouts
3 (Dixon, 2007). They are cultivated in many countries around the world, most commonly in
temperate climates of the Northern Hemisphere. Europe is a major center of diversity for
B. oleracea, or European Brassica vegetables, while Asia is the center of diversity for B. rapa, or
Asian Brassica vegetables (Cartea et al. 2011). Brassica oleraceae includes cabbage, broccoli,
cauliflower, kale, kohlrabi and Brussels sprouts. Brassica rapa includes turnip and Asian
vegetables that are leafy (such as pak choy) and headed (napa cabbage, also called Chinese
cabbage). Rutabaga is the only common B. napus vegetable. Brassica juncea crops include
mustard seed for condiment mustard and Asian greens such as red leaf mustard and mizuna.
Culinary vegetables in this family outside of the Brassica genus include Eruca sativa (arugula)
and Raphanus sativus (radish) (Tsunoda, 1980; Appel and Al-Shehbaz, 2003). Plants become
woody and less flavourful during the reproductive stage, and therefore non-bolting varieties are
popular for vegetable production. Most are biennial in nature, but are grown as annuals when
cultivated.
There has been increased attention to the health benefits of eating Brassica vegetables.
Kale and broccoli have been reported to reduce the risk of age-related chronic and degenerative
diseases and several types of cancer (Traka et al., 2010; Soengas et al., 2011). Health-promoting
phytochemicals in Brassica greens include vitamin C, carotenoids and DL-a-tocopherol (Singh et
al., 2007). Glucosinolates, such as glucoraphanin, glucoiberin and glucoraphanin, have received
special attention for their antioxidant and anticarcinogenic properties (Cartea et al., 2011).
Brassica greens are also a source of dietary fiber, which interacts with phytochemicals in the
digestive system (Palafox-Carlos et al., 2011).
4 1.1.3 History
Wild species of Brassica can be found across Europe and Central Asia, and the origin of
Brassica species is believed to be in the Mediterranean region (Tsunoda, 1980). Brassica rapa is
native to the highlands, rather than the coastal areas of the Mediterranean, which led it to adapt
relatively quickly to Scandinavia, Eastern Europe and China (Tsunoda, 1980). However, Ignatov
et al. (2010) has suggested that China is a centre of origin for the B. rapa Asian subgroup. Wild
types of B. oleracea with distinctive phenotypes are spread throughout Europe and the
Mediterranean, in small, isolated areas (Rakow, 2004).
1.1.3 Economic importance of Brassica crops in Canada
Oilseed rape is the second most economically important oilseed crop worldwide
(Snowdon et al., 2007). It is second only to soybean, and has been since the 1980s. As of 2008,
Canada was the largest producer of oilseed rape in the world (FAOSTAT data 2008, cited in
Hayward, 2012). Canola has the highest farm cash receipts of any agricultural crop in Canada
(Statistics Canada, 2013), and total production and farm cash receipts have increased from 12.9
million tonnes ($5.1 billion) in 2009 to 14.7 million tonnes ($8.2 billion) in 2013. In 2011, the
total acreage of canola production increased. However, there was an 8% decrease in canola
production due to a 17.5% decrease in yield per hectare (Statistic Canada, 2013). Production of
Brassica vegetable crops is listed in Table 1.1.
5 Table 1.1. Production (metric tonnes) and value of Brassica vegetables in Canada in 2012
(Statistics Canada, 2013).
Farm cash receipts (millions $ Canadian)
Production (thousand metric tonnes)
Cabbage 62.5 165
Broccoli 40.2 35
Cauliflower 23.9 31
Rutabaga and turnip 21.5 50
Radish 10.8 12
Brussels sprouts 7.5 6
1.1.4 Pests and diseases in Brassica crops
There are many diseases that affect Brassica crops in Canada, caused by bacteria, fungi,
viruses and other pathogens. Sclerotinia sclerotiorum (Lib.) Massee, a fungal stem rot, affects
both oilseed and vegetable crops worldwide. It is one of the most devastating diseases of
B. napus (rapeseed) crops in China (Zhao and Meng, 2003). Blackleg, also known as phoma stem
canker or dry rot, is caused by Leptosphaeria maculans (Desmaz.) Ces & De Not. This disease
affects oilseed crops as well as vegetable crops (Fitt et al., 2006). Alternaria leaf spot, caused by
Alternaria brassicae (Berk.) Sacc. and A. brassicicola (Schwein.) Wiltshire, attack oilseed,
vegetable and condiment brassica crops worldwide (Humpherson-Jones and Phelps, 1989).
Pathogens that cause damping off in Brassica crops include Fusarium spp., Rhizoctonia solani
(Jensen et al., 1999), and Pythium ultimum Trow and P. debaryanum Auct. non R. Hesse
(Abdelazher, 2003; Tanina et al., 2004). Downy mildew, caused by Peronospora parasitica, is an
oomycete pathogen of Brassica crops (Jensen et al., 1999). Clubroot, caused by Plasmodiophora
brassicae Woronin, is an economically important disease all over the world (Garber and Aist,
6 1979; Howard et al., 2010; Karling, 1968; Voorrips, 2003), but has the largest impact in
temperate countries (Karling, 1968).
Bacterial diseases are more problematic for vegetable crops than for oilseed crops. The
bacterial disease of greatest economic importance is black rot, caused by Xanthomonas
campestris pv. campestris (Pammel) Dowson (Williams, 1980; Berg et al., 2005). Another
important bacterial disease in vegetables is bacterial soft rot, caused by Erwinia carotovora var.
carotovora (Jones) Bergey et al. and Pseudomonas marginalis pv. marginalis (Brown) Stevens
(Ren et al., 2000; Charron and Sams, 2002).
Cauliflower mosaic virus and cabbage black ring spot are two viruses of Brassica
vegetables that are transmitted by the aphid species Myzus persicae Sulz. and Brevicoryne
brassicae L. (Broadbent, 1957). Another virus that affects B. rapa (turnip) crops is turnip yellow
mosaic virus, transmitted by flea beetles in the genus Phyllotreta (Broadbent, 1957).
Brassica crops are subject to herbivory by insects. Many species in the genus Lepidoptera
are herbivores of Brassica crops in Canada. Major pests of Brassica vegetable crops in Ontario
include caterpillar pests, such as imported cabbageworm (Pieris rapae L.), cabbage looper
(Trichoplusia ni Hübner), diamond-back moth (Plutella xylostella L.) and cabbage maggot (Delia
radicum L.). Mammestra configurata, also known as bertha armyworm, is a major oilseed crop
defoliator. Flea beetles (order Coleoptera, genus Phyllotreta) are important leaf feeders,
especially on seedlings (Lamb, 1989). Lygus spp. (L. elisus, L. lineolaris) and aphids
(M. persicae and B. brassicae) cause damage but generally have a minor impact on yield (Lamb,
1989). Swede midge (Contarinia nasturtii Keiffer) is an important pest of Brassica crops in
Ontario and an emerging pest in parts of western Canada. Native to Europe and Asia, adults
emerge from soil cocoons in mid May until mid June, with peak emergence occurring in the first
7 week of June (Allen et al., 2008). A second generation emerges 7-21 days later, depending on
climatic conditions, and there are generally four to five overlapping generations throughout the
season. Control of Swede midge relies on careful timing of insecticide application, based on
pheromone traps and field scouting (Hallett et al., 2009).
1.2 Clubroot of Brassica 1.2.1 Host range and history of clubroot
Clubroot is caused by the obligate Rhizarian pathogen Plasmodiophora brassicae
Woronin. Severe clubbing symptoms reduce the uptake of nutrients and water in Brassica crops
(Burki et al., 2010; Cavalier-Smith et al., 2013). Clubroot can infect most or all cultivated and
non-cultivated species of the family Brassicaceae, including all of the crops previously listed
(Sections 1.1.1, 1.1.2), and cruciferous weeds, such as Capsella bursa-pastoris (L.) Medik.
(shepherd's-purse), Sinapsis arvensis (L.) DC. (wild mustard), Diplotaxis muralis L. (annual
wall-rocket), Sisymbrium officianale (L.) Scop. (hedge mustard) and Arabidopsis thaliana (L.)
Heynh. (Buczacki and Ockendon, 1979). Clubroot was introduced to North America from
Europe, as a result of cross-Atlantic transport of livestock and clubroot-infected animal fodder. In
the 1920s, clubroot was reported in British Columbia, Ontario, Quebec and the Maritimes
(Drayton et al., 1926). Cabbage, cauliflower and rutabaga were the three main crops affected at
that time. Clubroot was first confirmed in canola fields in Alberta in 2003 (Tewari et al., 2005),
in Saskatchewan in 2009 (Dokken-Bouchard et al., 2010) and in Manitoba in 2013 (Strelkov and
Hwang, 2014). Previously, there were unpublished reports of clubroot in vegetable gardens in
western Canada (Tewari et al., 2005). In the United States, clubroot was confirmed on canola in
North Dakota in 2013 (Chittem et al., 2014).
8 The characteristic symptoms of clubroot are abnormal swelling of the roots caused by
hyperplasia and hypertrophy (Karling, 1968). Nutrient and water transport are progressively
inhibited as the clubs develop (Voorrips et al., 2003). This leads to the first noticeable,
aboveground symptoms of affected plants, which are wilting, yellowing and stunting. In severe
infections, plants die prematurely (Karling, 1968). Clubroot reduces both the yield and the quality
of the crop (Wallenhammar et al., 1999; Strelkov et al., 2005; Tewari et al., 2005; Dixon, 2009).
1.2.2 Causal agent and taxonomy
The causal agent of clubroot is P. brassicae, described by Mikhail Woronin in 1878.
P. brassicae is an obligate, biotrophic, soil-borne pathogen that reproduces in the cytoplasm of
the host plant cell (Karling, 1968; Williams and McNabola, 1967). The pathogen can survive
without a host as a dormant resting spore in soil for many years (Wallenhammar, 1996).
P. brassicae is classified in the order Plasmodiophorales, which consists of plant pathogens with
a multinucleate vegetative form, called a plasmodium (Karling, 1968). Characteristics of this
order include life stages of both resting spores and motile zoospores with two whiplash flagella
of unequal length. The family Plasmodiophoraceae consists of nine genera and 35 species. The
genera are distinguishable by the organization of cysts and sporangia. All of the species in the
genus Plasmodiophora have a plasmodial stage within the cell of the host plant, and they all
cause malformation due to hypertrophy (Karling, 1968). As scientific knowledge of clubroot has
developed over the past 100 years, the taxonomic classification of P. brassicae has evolved.
Plasmodiophorales, now classified in the kingdom Chromista, were previously classified as fungi
and protozoans (Cavalier-Smith, 2013). One example of a structural difference that distinguishes
9 chromists from fungi is the production of flagellated zoospores. Chromists have tubular ciliary
hairs, which protozoans lack (Cavalier-Smith, 1993).
1.2.3 Disease cycle
The lifecycle of P. brassicae begins with the germination of resting spores, followed by
two infection phases (Tommerup and Ingram, 1971). Resting spores are round and 2.8-5.0 µm in
diameter (Ayers, 1944; Ingram and Tommerup, 1972). Filamentous materials connect young
resting spores, while mature resting spores develop spines (Ikegami et al., 1978). During resting
spore formation, the nucleus and nucleoli become much smaller and undergo meiosis I and II,
characterized by two nuclear divisions (Garber and Aist, 1979). Cell walls of resting spores
contain chitins, proteins and lipids, and are resistant to microbial degradation (Buczacki and
Moxham, 1983). Resting spores are uni-nucleate and contain no nucleoli, but do contain
complete organelles (Williams and McNabola, 1967). The decay of clubs is aided by soil micro-
organisms (Ingram and Tommerup, 1972). The decay releases resting spores into the soil. Resting
spores can survive in a dormant state for a long period of time. The half-life of a soil’s infective
capacity has been calculated at 3.6 years in Sweden (Wallenhammar, 1996) and 4.4 years in
western Canada (Hwang et al., 2013). Under favourable conditions, resting spores germinate to
produce primary zoospores. Germination can take place with or without a host plant present
(Ingram and Tommerup, 1972; Friberg, 2005), but the frequency of germination is increased in
the presence of a compatible host (Macfarlane, 1970). Environmental influences on germination
include pH, soil moisture, soil temperature, and calcium ion concentration (Kageyama and
Asano, 2009). The likelihood of germination also increases with age and the level of decay of
spore-containing clubs (Macfarlane, 1970).
10 Root hair infection (RHI) occurs when primary zoospores move through the soil via films
of water and then penetrate root hairs of plants (Kageyama and Asano, 2009). Ayers (1944)
reported that primary zoospores are 2.8–5.9 µm in length. Primary zoospores are spindle-shaped
or ovate, with two flagella of unequal length (Ayers, 1944). The flagella are angled at 45 degrees
from each other, and consist of two micro-tubules and nine outer doublets (Aist and Williams,
1971). Zoospores attach to the cell wall of host root hairs in preparation for encystment. Flagella
coil around the zoospore body so that the zoospore is slightly flattened against the host cell wall,
and the axonemes are retracted so that the sheath membrane is no longer surrounding them (Aist
and Williams, 1971). During encystment, an enlarged adhesorium is formed by swelling of a
cylindrical structure called a satchel, which is 700 nm in length and packed with Golgi vesicles
and condensed cytoplasm (Aist and Williams, 1971). The adhesorium punctures the cell wall of
the root hair following adhesion to the cell wall (Aist and Williams, 1971). Host wall degradation
does not appear to be a factor in root hair penetration (Aist and Williams, 1971). RHI can occur
within 24 hours of resting spore germination (Dobson and Gabrielson, 1983). Penetration of the
cell wall takes 2.5-3.5 h. After a zoospore has entered the host cell, it becomes amoeboid and
forms a multinucleate thallus (Ayers, 1944). As thalli develop, they become detached from the
point of penetration in the root hair cell wall. Primary plasmodia undergo synchronous mitotic
divisions to form zoosporangia, each containing 4-16 secondary zoospores (Ayers, 1944; Ingram
and Tommerup, 1972). At favourable temperature conditions of 20-25°C, zoosporangial numbers
increase significantly between 3 and 4 days (Dobson and Gabrielson, 1983). At the optimum
temperature of 25°C, secondary zoospores are fully developed after approximately 5 days, and
are then released into the soil (Sharma et al., 2011a). There are no clubbing symptoms or crop
losses due to colonization of root hairs (Macfarlane, 1952; Kageyama and Asano, 2009).
11 Cortical infection, or secondary infection, follows RHI (Donald et al., 2008). Secondary
zoospores enter and infect root cortical cells (Kageyama and Asano, 2009). Aist and Williams
(1971) suggest that a similar process to the penetration of root hairs may occur in cortical
infection. The size of secondary zoospores after release is 9.6-14.4 µm in diameter (McDonald et
al. 2014). The mean diameter of encysted zoospores is 21.7 µm. Secondary zoospores are much
smaller when located inside root hairs (2.7-5.0 µm in very early stages, 3.3-4.5 µm in early
stages, and on average 8 µm later on) (Ikegami et al., 1978). The size difference between
secondary zoospores located in root hairs vs. released may be caused by higher osmotic or
physical pressure on secondary zoospores located inside root hairs (McDonald et al., 2014). In
the very early stage of development, secondary zoospores are spherical, with a smooth surface
and a membranous envelope (Ikegami et al., 1978), while in later stages, the surface of the
zoospore becomes rougher and the shape may range from spherical to ovate. Ikegami et al.
(1978) proposed that morphological differences between secondary zoospores may be related to
the nutrients available within the host cell tissue. Secondary zoospores are initially uni-nucleate
(McDonald et al., 2014), which indicates that fusion of zoospores may not be required for
secondary infection to take place, as was suggested in previous studies (Ingram and Tommerup,
1972). Under favourable conditions of 20-25°C temperature and high soil moisture, cortical
infection by secondary zoospores can occur in 3 hours (Dobson and Gabrielson, 1983).
Clubs are formed on plant roots, as the result of redirection of carbohydrates from the rest
of the plant to infected roots caused by the pathogen (Evans and Scholes, 1995). Cellular
multiplication, elongation, hypertrophy and hyperplasia take place at this stage, leading to visible
clubbing symptoms (Karling, 1968; Deora et al., 2012). Morphological changes are linked to the
disruption of plant regulators such as cytokinins and auxins, and an increase in host metabolism
12 (Muller and Hilgenberg, 1986; Ludwig-Muller, 1993; Jameson, 2000). Production of amino
acids, proteins, starches and lipids all increase during club development. Increases in
glucosinolate content, myrosinase and nitrilase also occur during club formation (Ludwig-Muller
et al., 1999; Grsic-Rausch et al., 2000).
Plasmodia form an envelope with seven layers and uniform thickness, mostly made up of
cells from the host, which surrounds each plasmodium during vegetative growth (Williams and
McNabola, 1967). Vegetative plasmodia consist of many spherical bodies with smooth surfaces,
which together create a sponge-like structure (Ikegami et al., 1978). Growth stops when
sporogenesis begins (Williams and McNabola, 1967). At this time, the envelope that surrounds
each plasmodium gets thinner, shedding its outer layers (Williams and McNabola, 1967). By the
time the vegetative growth period is ending, the plasmodia take up most of the host cell, and the
cytoplasm has been forced to the periphery of the cell (Williams and McNabola, 1967).
Eventually the clubs begin to decrease their production of sugars, RNA and starch, and the
plasmodia divide into millions of resting spores. The transition to resting spores happens very
quickly (Williams and McNabola, 1967). As the roots break down and decay, the plant loses
what is left of its ability to take in water and nutrients and can no longer support itself (Williams
and McNabola, 1967; Voorrips et al., 2003).
1.2.4 Pathotypes
Pathogen specialization in P. brassicae, historically known as ‘races,’ are now called
‘pathotypes’ (Williams, 1966; Ayers, 1972). Pathotype is based on virulence, whereas race
categories are based on genes, alleles or chromosome structure (Sturhan, 1985). In Canada, the
Williams pathotype classification system is typically utilized (Williams, 1966). In this
13 classification system, some populations may have only one phenotype present for virulence,
while others have several phenotypes in different frequencies (Sturhan, 1985).
Pathotypes 2, 3, 5, and 6 are most prevalent in Canada. Pathotype 6 is most common in
Ontario, and also occurs in British Columbia and Quebec (Reyes, 1974; Williams, 1966).
Pathotype 2 is most prevalent in Quebec, but is also found in the Atlantic provinces (Ayers,
1972). Pathotypes 3 and 5 have been found on canola in Alberta since 2003, and presumably the
other prairie provinces (Ayers, 1972; Cao et al., 2007). In addition, studies of single-spore
isolates indicate that pathotype 8 may be present near Edmonton, Alberta and Orton, Ontario
(Xue et al., 2008). In 2013, a novel strain of the pathogen, initially called 5x, was identified near
Edmonton, Alberta. A range of genotypes of the pathogen have since been identified in fields
where resistance has been overcome. The various genotypes that were subsequently identified
were virulent against all commercially available canola cultivars in Canada (Cao et al., 2015;
S. Strelkov, unpublished). The new pathotypes are referred to collectively as pathotype X until a
more definitive identification is possible.
1.2.5 Incompatible Interactions
An incompatible interaction takes place between a host plant that is resistant and a
virulent strain of the pathogen. In most incompatible interactions with P. brassicae, clubroot
symptoms do not develop and resting spores are not produced (Ludwig-Muller et al., 1999; Feng
et al., 2012a). Root hair infection can occur on host and non-host plants (Dixon, 2006).
Secondary plasmodia have been observed in clubroot-resistant canola cultivars (Donald et al.,
2008; Gludovacz, 2013; McDonald et al., 2014). However, pathogen development is inhibited
before completing its lifecycle and clubroot symptoms are not observed (Morgner et al., 1995;
14 Hwang et al., 2011b). During this stage, auxin levels increase in resistant plants (Ludwig-Muller
et al., 1993). Park et al. (2007) suggested that an antagonistic interaction between growth
hormones such as auxin and salicylic acid may lead to reductions in plant growth during disease
resistance.
1.3 Factors influencing infection and development
The small but abundant resting spores of P. brassicae are readily transported anywhere that
contaminated soil or water can go. For example, spores can be moved on Brassica seedlings for
transplanting, hand tools, trucks, farm machinery, livestock, compost, harvest bins, boots, gloves
and other items of clothing of farm workers. Transportation of livestock fodder, such as turnip,
may also transport the disease (Donald et al., 2006). Clubroot spread from one localized area to
an entire region in the Holland Marsh, Ontario, following flooding associated with Hurricane
Hazel in 1954 (Conners et al., 1956). This indicates that flooding may be responsible for
movement of resting spores up to 8 km. Resting spores may also be present in airborne dust and
may be dispersed up to 2 km by wind-mediated soil erosion. However, it is not known whether
those spores remain viable and it is unlikely that enough windborne resting spores travel
distances that are long enough to establish new infestations (Rennie et al., 2015).
The physical condition of the soil has a great influence on clubroot incidence and severity.
Important influences on clubroot development include soil moisture, temperature, pH,
concentration of resting spores, and the level of host plant resistance or susceptibility (Murakami
et al., 2002; Dixon, 2006). Micronutrient levels such as calcium and boron can also have an
effect on clubroot development (Webster and Dixon 1991a, 1991b).
15 1.3.1 Soil moisture
High soil moisture is favourable for clubroot development (Karling, 1968). Soil moisture
is generally measured as gravimetric water content (mass of water per mass of soil) or volumetric
water content (volume of water per volume of soil) (Or and Wraith, 2002). Resting spore
germination is delayed at soil moisture levels below 30% moisture content (Macfarlane, 1952),
though continuous levels of soil moisture are not required. Short intervals of high moisture
content following heavy rains are sufficient for infection, especially in the 2-3 weeks following
seeding (Thuma et al., 1983). Moisture levels affect the motility of zoospores within the soil
(Colhoun, 1973). However, pathogen development can be lower in saturated soils, when all
available pore spaces are filled with water (Gludovacz, 2013). There is a positive correlation
between total rainfall and clubroot incidence and severity (Gossen et al., 2012). Cortical infection
requires higher levels of soil saturation than is required for root hair infection (Dobson et al.,
1982). The interaction between soil moisture and temperature also plays a key role in clubroot
development (Donald et al., 2006).
1.3.2 Temperature
Temperature has a major influence on clubroot development, with a soil temperature of
18-25°C being most favourable for pathogen development, especially in the first 2-3 weeks of
plant growth (Colhoun, 1953; Buczacki et al., 1978; McDonald and Westerveld, 2008; Sharma et
al., 2011a, 2011b; Gossen et al., 2012). Soil temperature at 5-cm depth has a strong correlation
with clubroot severity and incidence (McDonald and Westerveld, 2008). Air temperature may
also be used as an effective indicator of clubroot development, in particular during the last 10
days before harvest for fast growing Asian Brassica vegetable crops such as Shanghai pak choy
16 (McDonald and Westerveld, 2008). The optimal temperature for germination of resting spores is
also 25°C, the temperature at which activity of the gene product Pro1 is highest and other
germination factors are stimulated (Dixon, 2009; Feng et al., 2010). Clubroot development is
very slow at air temperatures below 17°C and above 30°C (Sharma et al., 2011a, 2011b). The
role of temperature in infection and symptom development has been confirmed in seeding date
trials that provide a range of temperatures for clubroot development under field conditions
(Gossen et al., 2012). Seeding date also affected clubroot development in short-season Brassica
vegetables, such as Shanghai pak choy, with the lowest incidence in May and September
plantings when the mean air temperature was lower during the early stages of plant growth
(Gossen et al., 2012). Although canola is a long-season crop with limited seeding date flexibility,
research has shown that seeding 1-3 weeks earlier in spring can decrease clubroot severity
(Hwang et al., 2011a, 2012a) because plants can become established during temperatures that are
too low for clubroot development.
1.3.3 Soil pH
Increased soil pH is directly correlated with decreased clubroot incidence and severity,
particular when soil pH is above 7.0 or 7.2 (Webster and Dixon, 1991a, 1991b; Donald and
Porter, 2004). Indeed, clubroot mitigation recommendations for vegetable crops often
recommend raising soil pH to 7.2 or above (OMAFRA, 2010). With adequate soil moisture and
optimal temperatures for clubroot development, however, moderately severe clubroot symptoms
developed up to a pH of 8 (Gossen et al., 2013). At temperatures below 17°C, there is a weaker
correlation between pH level and clubroot incidence and severity.
17 1.3.4 Resting spore concentration
There is a positive correlation between resting spore levels in the soil and clubroot
severity in susceptible Brassica crops (Dixon, 2006; Hwang et al., 2011a, 2012b). Depending on
environmental conditions, an inoculum concentration of 1000 spores g-1 dry soil is considered to
be the threshold for clubroot development in most susceptible cultivars (Murakami et al., 2002;
Donald and Porter, 2009; Faggian and Strelkov, 2009). The effect of resting spore concentration
on the severity of clubroot symptoms varies depending on the interaction with other
environmental factors, such as pH (Colhoun, 1953). Resting spores were distributed throughout
the soil profile to depths of 102 cm (Cranmer, 2015), although a previous report did not find
resting spores deeper than 45 cm below the soil surface with more than 97% of spores in the top
0-5 cm of soil (Kim et al., 2000).
1.4 Disease management 1.4.1 Cultural control
No single method provides substantial, consistent clubroot reduction, so an integrated
approach is essential for effective clubroot management (Donald et al., 2006; Diederichsen et al.,
2009). Recommended practices for sanitation and prevention of pathogen spread include nursery
hygiene for transplanted crops, avoidance of contaminated irrigation sources, and prevention of
cross-contamination by the movement of farm equipment from infested to non-infested fields
(Donald et al., 2006). Liming is a form of cultural control that has been commonly recommended
against clubroot in Brassica vegetable crops (Colhoun, 1953; Murakami et al., 2002). Small
increases in pH can suppress clubroot. However, at temperature and moisture levels that are
optimal for clubroot development, changes in pH are less effective at suppressing clubroot
18 (Gossen et al., 2013). Furthermore, liming is not an economically viable solution for canola
producers due to the high cost (Howard et al., 2010), and the effect can be inconsistent from year
to year (McDonald et al., 2004; McDonald and Westerveld, 2008). Increasing the pH level may
also have negative impacts on uptake of other nutrients (Hwang et al., 2008; Gossen et al., 2012)
and so may not be suitable for other crops in a rotational system (Hildebrand and McRae, 1998).
Boron and calcium have been found to suppress clubroot in Brassica vegetables (Webster
and Dixon, 1991a, 1991b), with a positive correlation between the rate of drench application of
boron and clubroot suppression (Deora et al., 2011). Field trials in high organic content soil
(Holland Marsh, Ontario) demonstrated that boron applied at a rate of 4 kg B ha-1 reduced
clubroot severity by 64% relative to the control (Deora et al., 2011). This was the most effective
rate with no phytotoxic effects, which occurred at rates above 2 kg B ha-1 in sand under
controlled conditions and 48 kg B ha-1 or greater in field conditions (Deora et al., 2011).
Phytotoxic effects of boron include chlorosis and necrosis. Boron is active against both the
primary and secondary stages of infection (Webster and Dixon, 1991b; Deora et al., 2011).
However, drench applications of boron are not considered to be economical for canola producers
due to the high water volume required for root drenches. Other application techniques may be
feasible, for example in combination with other fertilizers at sowing (Deora et al., 2011).
Calcium cyanamide (nitrogen and calcium oxide) has been used to suppress clubroot for
approximately 60 years, in addition to its uses as a slow-release nitrogen fertilizer and herbicide.
Calcium cyanamide (Perlka, 50% calcium oxide, 19.8% nitrogen, 1.5% magnesium oxide) was
effective in suppressing clubroot on Asian Brassica vegetables (McDonald et al., 2004). High
levels of limestone or wood ash (± quintozene fungicide) also suppressed clubroot, but the
19 authors concluded that soil amendments to change pH were not a practical or economically viable
option for canola producers (Hwang et al., 2011c).
Calcium application can delay pathogen maturation and reduce the total pathogen biomass
in root hair cells. At pH 5.5, a delay in pathogen development was observed only at the highest
rate of calcium application, compared to the control, while at pH 6.5, a delay was observed at all
rates of calcium (Webster and Dixon, 1991a). The effectiveness of calcium amendments
decreased as the pH increased above 6.5 to 8.0 (Donald and Porter, 2004).
Production of susceptible crops in a short- to mid-term (2-5 year break from canola) crop
rotation is not a viable management option. After a single year of cropping with a susceptible
cultivar in a soil already infested with 1 x 106 resting spores per mL, the increase in spore load
was more than 60-fold (Hwang et al., 2013). In fields with high clubroot infestation, it took 17.3
years for the pathogen level to decrease below a detectable level (Wallenhammar, 1996). There
are, however, benefits to implementing short-term rotations in combination with clubroot-
resistant cultivars. A break from canola for two or more years reduced the concentration of
P. brassicae resting spores in the soil up to 10-fold and increased yield in resistant canola (Peng
et al., 2013). The concentration of P. brassicae resting spores declined by 98% from the initial
concentration following a 2-year break from susceptible canola (Peng et al., 2015). Increasing
cropping diversity in canola production in the absence of P. brassicae results in a yield increase
of 22% in a crop rotation with canola 1 in 6 years (Harker et al., 2014).
1.4.2 Synthetic fungicides and surfactants
Numerous fungicides have been tested against clubroot, including fluazinam (3-chloro-N-
[3-chloro-2,6-dinitro-4-(trifluoromethyl)phenyl]-5-(trifluoromethyl)-2-pyridinamine) (Allegro®
20 500F, ISK Biosciences Corporation), which is registered for management of clubroot on Brassica
vegetables in Canada, and cyazofamid (4-chloro-2-cyano-N,N-dimethyl-5-p-tolylimidazole-1-
sulfonamide) (Ranman® 400SC ISK Biosciences Corporation). Cyazofamid directly inhibits
resting spore germination, which in turns inhibits both root hair infection and pathogen
development (Mitani et al., 2003). Inhibition of root hair infection increases as the incubation
period with cyazofamid increased from 1 to 10 days (Mitani et al., 2003). Fluazinam disrupts the
oxidative phosphorylation metabolic pathway in mitochondria (Guo et al., 1991). Fluazinam and
cyazofamid provided 100% clubroot control on Shanghai pak choy under controlled conditions
(Adhikari, 2010). However, fluazinam and cyazofamid were not effective under high pathogen
populations in field trials (Tanaka et al., 1999; Saude et al., 2012; Peng et al., 2014), possibly due
to insufficient coverage over the entire soil volume, or degradation / leaching from the root zone
(Peng et al., 2014).
There are currently no fungicides registered in Canada for management of clubroot on
canola, and drench application would not be economically viable for canola producers (Howard
et al., 2010). Opportunities for fungicidal control, however, could include seed treatments or
fumigants, used only to treat localized areas (Donald and Porter, 2009), such as low-lying areas
with a higher spore concentration due to increased moisture, compaction and / or activity from
equipment carrying contaminated soil. Various surfactants have been evaluated for use against
clubroot (Humpherson-Jones, 1993; Hildebrand and McRae, 1998) though none have been
registered in Canada, possibly due to phytotoxicity (Howard et al., 2010).
21 1.4.3 Biological control and biofungicides
There are no proven, cost-effective biological controls for clubroot management at the
current time. However, there are some promising biological control agents (BCA) and
biofungicides (Peng et al., 2011; Kasinathan, 2012; Lahlali et al., 2013). A BCA is a living
organism that can be used to suppress pathogens by natural competition, parasitism and / or as a
microbial antagonist. A biofungicide is either a living organism or a compound derived from a
living organism that is toxic to fungal pathogens (Pal and McSpadden Gardener, 2006).
Biological control agents have been identified that cause induced systemic resistance
(ISR) to clubroot and other diseases. These BCAs protect the plant for a longer period of time
than synthetic fungicides (Peng et al., 2011). Microorganisms with activity against clubroot
include fungi such as Trichoderma, and bacteria such as Streptomyces (Cheah et al., 2000).
Commercial biofungicides with activity against clubroot available in Canada include Mycostop
(S. griseoviridis) (Verdera Oy, Finland), Actinovate (S. lydicus) (Natural Industries, USA), Root
Shield (T. harzianum) (BioWorks Inc. USA), Prestop (Gliocladium catenulatum syn.
Clonostachys rosea f. catenulate) (Verdera Oy) and Serenade (Bacillus subtilis QST713) (Bayer
CropScience, Germany).
Bacillus subtilis effectively colonized canola roots, a critical component to induced
resistance and antibiosis (Lahlali et al., 2013). B. subtilis resulted in small reductions in clubroot
severity as a soil drench and as a seed treatment in some, but not all, studies under controlled
conditions (Peng et al., 2011). Drench applications of Serenade and Prestop were moderately
effective in canola under controlled conditions (Kasinathan, 2012). Both Serenade and Prestop
were more effective than indigenous BCA isolates and as effective as synthetic fungicides under
low pathogen pressure (Peng et al., 2011). Neither Serenade nor Prestop effectively suppressed
22 clubroot in soil with high concentration of resting spores in field trials (Kasinathan, 2012; Peng et
al., 2014). Adequate soil moisture and distribution are required for B. subtilis to effectively
suppress clubroot. Depending on temperature in field conditions, primary and secondary
infections could take place over a period of 3 weeks in spring. Therefore, it can be challenging to
ensure adequate coverage of B. subtilis during the period of potential infection (Adhikari, 2010;
Peng et al., 2014).
Initial testing indicated that Actinovate was only effective at low inoculum levels, while
Mycostop was more consistently effective across inoculum levels (Adhikari, 2010). The efficacy
of each of these biofungicides varied greatly among trials and sites (Peng et al., 2014).
Acremonium alternatum is an endophytic fungus that enters the host plant through roots
and colonizes root and leaf cells. When applied to Arabidopsis thaliana, it reduced clubroot
severity by 50% and delayed development of P. brassicae (Jaschke et al., 2010). Heteroconium
chaetospira (Grove) M.B. Ellis is another endophyte that reduced clubroot severity by 52-97% in
field trials (Narisawa et al., 2000). A high rate of H. chaetospira reduced severity on canola
exposed to low levels of the pathogen, but was not effective when pathogen pressure was high.
When applied in combination with P. brassicae inoculation, H. chaetospira increased the activity
of phenylalanine ammonia lyases (PAL), and the expression of genes involved in the biosynthesis
of jasmonic acid, ethylene, auxin and PR-2 proteins (Lahlali et al., 2014). In broccoli
(B. oleracea var. italica), stimulation of defense response with salicylic acid activated
pathogenesis-related (PR) genes and reduced clubroot symptoms (Lovelock et al., 2013).
Two organic plant growth stimulants, Fructigard and PlasmaSoil (TILCO, Biochemie),
consisting of algal extracts, amino acids and phosponate, are effective against clubroot on napa
cabbage (B. rapa subsp. pekinensis) and canola (B. napus), but ineffective on Arabidopsis
23 thaliana. Plants with induced resistance exhibited the following differences: cells were more
densely packed in secondary phloem, large cells were retained in root cortex, and the outer
protective cell layer was thicker and impermeable (Kammerich et al., 2014). Individual
components of the formulation did not have the same effect.
1.4.4 Host plant resistance
Resistant cultivars are an important component of effective, integrated clubroot control in
the Brassicaceae family (Diederichsen et al., 2009). Clubroot resistance genes have been found
in B. napus, B. oleracea and B. rapa (Hirai, 2006; Piao et al., 2009). The CRa gene in B. rapa
was the first clubroot resistant gene to be identified (Ueno et al., 2012). European fodder turnips
(B. rapa ssp. rapifera) carried multiple genes for resistance (Delourme et al., 2012). It is
common, however, for a single dominant resistance gene to be introduced to another crop, for
example in B. napus Swedish winter oilseed rape cultivar Tosca (Delourme et al., 2012). A single
major gene generally provides resistance to one specific pathotype, and resistance has been
overcome over time (Kuginuki et al. 1999; Dederichsen et al., 2009; Ueno et al., 2012). In
B. napus, resistance genes have been named Pb-Bn and PbBn. (Piao et al. 2009). To avoid
confusion, Piao et al. (2009) suggested standardized loci nomenclature for B. napus (PbBn),
B. rapa (PbBr) and B. oleracea (PbBo).
Canola cultivars with high levels of resistance to clubroot that are currently commercially
available in Canada include 45H29 (Pioneer Hi-Bred, Mississauga, ON), D3152 (DuPont
Canada, Mississauga, ON), Proven 9558C (Viterra, Regina, SK), 1960 and CS 2000 (Canterra
Seeds, Winnipeg, MB), 73-67 and 73-77 (Monsanto, Winnipeg, MB), 6056 CR (Brett Young,
Winnipeg, MB), V12-3 (Cargill Limited, Winnipeg, MB) and 14H1176 (Syngenta Canada, Inc.,
24 Guelph, ON). Resistance to pathotypes 2, 3, 5 and 6 was demonstrated in four commercial
cultivars of canola. These cultivars all had uniform responses at both the root hair and cortical
infection stages (Deora et al., 2013). Cultivar 45H21 (Pioneer Hi-Bred, Mississauga, ON) was
found to be resistant to pathotype 6 only. Resistance to pathotype 3 is currently essential for
commercial cultivars in many areas of Alberta, and effective resistance to the new strains of
P. brassicae in western Canada will be essential for clubroot management in this region over the
next decade.
Integrated management strategies for clubroot can improve the durability of genetic
resistance by reducing selection pressure. Durability of resistance refers to the effectiveness of
genetic resistance over time during prolonged and widespread use in the presence of the pathogen
under favourable environmental conditions (Johnson, 1984). In smaller pathogen populations, the
ability to adapt to plant host resistance is more limited because the likelihood of new genetic
variations arising from mutations or recombination is smaller (Fisher, 1930). Population size may
be reduced in a number of ways: decreasing the initial inoculum load, routine reductions, such as
fungicide applications, or limiting the movement of genotypes among pathogen populations,
which decreases the spread of mutant alleles and genotypes (McDonald and Linde, 2002). More
durable resistance can be developed through efforts focused on developing broad-spectrum
resistance (Piao et al., 2009). Broad resistance to clubroot is defined as resistance against the
majority of P. brassicae pathotypes (Fu et al., 2011). This strategy could incorporate disrupting
pathogen selection for virulence by rotating cultivars with different major resistance genes over
time, when available or pyramiding several resistance genes in one cultivar (McDonald and
Linde, 2002; Ueno et al., 2012). For example, the new canola cultivar PV580GC (Crop
25 Production Services Canada, High River, AB), registered in October 2015, will be the first
multigenic clubroot-resistant canola cultivar available in Canada (CPS Canada, 2015).
1.4.5 Metabolic cost of resistance
A metabolic cost of disease resistance is the negative effect on growth and development
that occurs when a plant’s resources are reallocated toward defense in response to pathogen
recognition (Brown and Rant, 2013). It is also called a fitness cost of disease resistance (Brown
and Rant, 2013). Continuous low levels of expression of a major resistance gene can cause
slightly decreased productivity in plants even in the absence of the pathogen, although more
significant fitness costs are incurred when presence of the pathogen leads to induction of a
resistance response (Bergelson and Purrington, 1996). Induced resistance is the activation of
previously inactive resistance mechanisms, rather than the fabrication of new modes of resistance
that did not previously exist in that plant host, and relies on signals effectively triggering the
plant’s defensive response against an attacking pathogen (Van Loon et al., 1998). The cost of
disease resistance can be seen as a trade-off, because there is a greater cost of not expressing a
defense mechanism in the presence of a virulent pathogen.
In canola, root hair infection plays an important role in recognition and induction of
resistance to P. brassicae (Feng et al., 2012a). The metabolic cost of clubroot resistance may
result in decreasing vegetative growth or seed production, or delayed maturity (Hwang et al.,
2011a; Deora et al., 2012b; Peng et al., 2014). In western Canada, delayed maturity of canola is a
particular concern because it can result in increased susceptibility to frost damage, with
associated loss in quality as well as yield.
26 1.5 Summary and objectives
Clubroot is an economically important disease of canola and Brassica vegetable crops in
Canada. Fungicides and biofungicides are not currently used in commercial canola production
because results have been inconsistent and they are not economically viable. Therefore, genetic
resistance is essential for clubroot management. The mechanisms of resistance to clubroot are not
well understood, and resistance has often been quickly overcome by the pathogen. Previous
research under controlled conditions has indicated that there may be a metabolic cost of
resistance to clubroot. In addition, the effect of resting spore concentration on the severity of
clubroot symptoms varies depending on the interaction with other environmental factors. Also,
plant growth and yield increase in response to increased diversity in crop rotation, compared to
continuous canola. More research on the effect of short- to mid-term (2-5 year break from canola)
crop rotation on concentration of resting spores in the soil and on metabolic cost of resistance
could have important implications for the development of integrated clubroot management
systems for canola.
The objectives of this research were to:
1. Examine the effect of concentration of P. brassicae resting spores on growth and
development of clubroot-resistant canola and Brassica vegetables, to identify if there is a
metabolic cost of resistance.
2. Examine the effect of crop rotation on the rate of decline of resting spores over time and
the concentration of resting spores in soil.
3. Examine the interaction between crop rotation and inoculation with P. brassicae on
growth of clubroot-resistant canola.
27 4. Assess the interaction between pH and inoculum concentration of P. brassicae on
clubroot incidence and severity of susceptible canola.
The following hypotheses were tested:
1. Plant height and biomass are reduced and development is delayed by increasing
concentration of resting spores in resistant plants.
2. The concentration of resting spores in soil declines exponentially with increasing length
of break interval following a susceptible canola crop.
3. The interaction of crop rotation and inoculation with P. brassicae results in a greater cost
of resistance to clubroot in soil with a history of continuous canola compared to a more
diverse crop rotation.
4. At low concentrations of resting spores, clubroot incidence is reduced at pH >7.0
compared with pH 6.0 to 7.0, but there is no effect of pH at high concentrations of spores.
28 CHAPTER TWO
EFFECT OF RESTING SPORE CONCENTRATION ON GROWTH OF
CLUBROOT-RESISTANT BRASSICA CROPS
2.1 Introduction
Inoculum concentration in soil is an important factor influencing infection by soil-borne
pathogens (Richardson and Munnecke, 1964). Disease severity increases with increasing
inoculum concentration in many susceptible plant-pathogen interactions, including Fusarium wilt
of lettuce (Lactuca sativa L.) caused by Fusarium oxysporum f. sp. lactucae (Scott et al., 2012),
root rot in lentil (Lens culinaris Medikus) caused by Rhizoctonia solani Kühn (Chang et al.,
1998) and pre-emergence damping off in field pea (Pisum sativum L.) caused by Pythium
irregulare Buisman (Richardson and Munnecke, 1964). Reduction of inoculum concentration in
soil is the focus of many disease management strategies, including soil solarization, fumigation
and crop rotation with non-host or bait crops. For example, soil solarization reduces fusarium wilt
of lettuce by up to 91% (Matheron and Porchas, 2010). Fumigation with methyl bromide plus
chloropirin reduces Verticillium dahliae Kleb. microsclerotia by 93% (Short et al., 2015) and
crop rotation reduces inoculum concentration of F. oxysporum f. sp. lactucae by 86% within 12
months in the absence of a host crop (Scott et al., 2012).
Genetic resistance is essential for management of many soil-borne pathogens, including
F. graminearum in wheat (Triticum aestivum L.) (Buerstmayer et al., 2009), Sclerotinia
sclerotiorum (Lib.) de Bary in soybean (Glycine max (L.) Merr.) (Cober et al., 2003) and
P. brassicae in canola (Cao et al., 2009). However, resistance to plant pathogens can be
associated with a metabolic cost to the plant, also called the fitness cost of disease resistance.
This can result in a negative association between disease resistance and other traits, such as plant
29 maturation and yield. The negative effect of the reallocation of resources from growth to defense
during the disease resistance response can be quantified using measurable indicators of plant
fitness. Bergelson and Purrington (1996) reviewed 88 published studies and found that overall
about 50% of plant species surveyed had a significant fitness cost of disease resistance.
The effect of inoculum concentration on resistant crops is generally not well understood
because fitness costs of disease resistance are challenging to identify and to quantify. There may
be several physiological and genetic factors contributing to the overall cost of resistance (Brown
and Rant, 2013). Yield is determined by numerous traits and genetic components (Campbell and
Kondra, 1978; Foulkes et al., 2000). Indicators such as vegetative growth, leaf area and pod
development are useful in the assessment of cost of resistance because they make important
contributions to yield of canola (Campbell and Kondra, 1978; Freyman et al., 1973; Krogman
and Hobbs, 1975, Chongo and McVetty, 2001). A fitness cost can be incurred either from the
presence of the resistance gene, or from expression of defense by the resistant plant upon
recognition of the pathogen (Brown and Rant, 2013). Constitutive defenses, which are always
present in the plant, have continuous low levels of expression. This leads to a mean fitness cost of
3.5% reduced productivity compared to susceptible plants, averaged across many different crops
(Bergelson and Purrington, 1996). Induced resistance is believed to incur less metabolic cost
overall, compared to constitutive resistance, because the costs are reduced (though not
eliminated) in the absence of pests (Purrington, 2000; Walters et al., 2013). The overall cost of
resistance can be seen as a trade-off, because there is a greater cost of not expressing a defense
mechanism in the presence of a virulent pathogen.
The trade-off between plant growth and defense against pathogens involving the
jasmonante-signaling pathway is a widely conserved defense strategy in angiosperms. It has been
found that gibberellic acid activates defense and also mediates a delay in the degradation of
30 growth-response proteins in rice (Oryza sativa L.) and Arabidopsis thaliana (L.) Heynh. (Yang et
al., 2012). In Arabidopsis, the PMR6 gene codes for a protein required for susceptibility to
powdery mildew (Erysiphe cichoracearum DC.) and may also influence plant growth and
development and pectin degradation (Vogel et al., 2002). A pmr6-1 mutant with loss of function
for the PMR6 gene (resistant to powdery mildew) is 23% smaller in rosette diameter and has
increased pectin in cell walls relative to the susceptible control. Pectin in cell walls may play a
role in resistance to powdery mildew (Vogel et al., 2002). Model plant-pathogen systems using
mutant constructs with loss of function for an identified resistance gene could improve
understanding of metabolic cost of resistance to clubroot.
In commercial canola cultivars, a major challenge for comparison of fitness costs is that
near-isogenic lines, differing only in presence or absence of resistant gene, are not available.
Instead, comparison of fitness must be conducted between induced and non-induced plants to
assess the overall costs of induced resistance (Purrington, 2000). Studies of other plant-pathogen
interactions demonstrate that induced resistance costs are commonly related to a hypersensitive
response (HR) (Boyd et al., 1995; Tian et al., 2003). However, in commercial canola cultivars,
HR is not the mechanism of clubroot resistance (Deora et al., 2013).
In response to clubroot infection, resistant canola (B. napus) roots form a ring of
concentrated reactive oxygen species (ROS) in the inner cortex. ROS accumulation is a response
to wounding, which reinforces cell wall proteins and limits P. brassicae invasion of vascular
tissues during secondary colonization (Deora et al., 2013). Enzymes that metabolize ROS, such
as copper / zinc superoxide dismutase cytochrome c oxidase, decreased in susceptible canola
within the first 12 hours after inoculation, and then increased 24-72 hours after inoculation (Cao
et al., 2008). Indole-3-acetic acid (IAA) increases continually in resistant cultivars of napa
cabbage (B. rapa subsp. pekinensis) (Ludwig-Muller et al., 1993) and canola (Feng et al., 2012).
31 Root hair infection plays an important role in recognition and induction of resistance to
P. brassicae in host plants (Siemens et al., 2002; Feng et al., 2012; McDonald et al. 2014).
Clubroot symptoms do not develop and resting spores are not produced during an incompatible
interaction (Ludwig-Muller et al., 1999; Feng et al., 2012). However, root hair infection may take
place on host, non-host and resistant plants (Feng et al., 2012).
Increasing the resting spore concentration of P. brassicae from 1 x 105 to 1 x 108 spores
g-1 soil increases clubroot severity and decreases plant height and seed yield in susceptible canola
(Brassica napus L.) cultivars (Hwang et al., 2011a). In a recent study, four clubroot-resistant
canola cultivars (45H29, 73-67, 73-77 and Proven 9558C) inoculated with P. brassicae exhibited
reductions in plant height, number of flowers and pods, and delayed progression from the
vegetative to reproductive stage (Deora et al., 2013). These findings were consistent with two
previous studies showed that seedling emergence, height and yield were negatively correlated
with increasing resting spore concentration in a resistant canola cultivar (Hwang et al., 2011a;
Deora et al., 2012b). In western Canada, delayed maturity can lead to late canola crops, which
may then be damaged by frost.
In a susceptible cultivar, 1000 spores g-1 soil is commonly cited as the infection threshold
(Donald and Porter, 2009). However, the effect of resting spore concentration on the severity of
clubroot symptoms varies depending on the interaction with other environmental factors, such as
pH (Colhoun, 1953). Root hair infection (RHI) declines above pH 6.5 (Myers and Campbell,
1985; Webster and Dixon, 1991; Donald and Porter, 2004; Gossen et al., 2013). Development of
primary plasmodia is inhibited at pH ≥7.2 prior to secondary zoospore release (Myers and
Campbell, 1985). The interaction between pH and resting spore concentration is not well
understood. Maximum root hair infection in slightly acidic soil (pH 6-6.5) occurs at temperatures
near 25°C (Gossen et al., 2013). Severe clubbing occurs at resting spore concentrations as low as
32 103 at pH 6.3, but resting spore concentration is positively correlated with clubroot incidence in
alkaline soil (pH 7.8) (Colhoun, 1953). Increased understanding of this interaction could make it
easier to predict clubroot severity and improve recommendations regarding use of resistant
cultivars or other management practices.
Clubroot management that is sustainable for several years requires effective strategies for
reducing the concentration of resting spores in soil. To date, the implementation of cultural,
biological and chemical strategies have been ineffective (Ahmed et al., 2011) or not cost-
effective (Porter et al., 1991; White and Buczacki, 1977). In fields with a high initial level of
inoculum, the concentration of resting spores may remain above the infection threshold for a
susceptible crop for many years (Wallenhammer, 1996), even though there has been a significant
reduction in the number of resting spores (Peng et al., 2014).
There were two objectives of this research. The first was to identify and quantify the
metabolic cost of resistance to P. brassicae by examining the effect of the concentration of
resting spores of P. brassicae on growth and development of clubroot-resistant canola and
Brassica vegetable cultivars. It is hypothesized that increasing inoculum concentration will
reduce plant growth in canola, napa cabbage and cabbage plants and delay development in
canola. The second objective was to assess the interaction between pH and resting spore
concentration of P. brassicae on clubroot incidence and severity in susceptible canola. It is
hypothesized that pH will not affect clubroot incidence at high concentrations of resting spores,
but clubroot incidence at low concentrations of resting spores will be reduced at pH >7.0
compared with pH 6.0 to 7.0.
33 2.2 Materials and Methods 2.2.1 Controlled environment study – canola
Growth room studies were conducted using canola cultivar 45H29 (Pioneer Hi-Bred Ltd,
Chatham, ON), which is resistant to pathotype 6 (Deora et al., 2013) of P. brassicae (Williams’
system). The seeds were sown in tall plastic pots (conetainers) filled with soil-less media
(Sunshine Mix #4, Sun Gro Horticulture Canada Ltd., Agawam, MA). Two seeds were planted
per pot and thinned to one seedling per pot. Four replicates of 10 pots each of the clubroot-
susceptible canola line ACS-N39 (AAFC, Saskatoon SK) were included as a susceptible control
(Gludovacz, 2013). The plants were maintained at 25°/20° C day/night, with 16-hr photoperiod
and 65% relative humidity, and fertilized with 1 g L-1 N-P-K (20-20-20) and 1 g L-1 magnesium
sulphate solution at 2–3 day intervals. The plants were watered with deionized water adjusted to
pH 6.0 using commercial white vinegar. Each 9-day-old seedling was inoculated with 5 mL of
resting spore suspension of P. brassicae pathotype 6 in the following treatments: 0, 1 x 104,
1 x 105, 1 x 106, 1 x 107 and 1 x 108 spores mL-1. Inoculum was prepared following a standard
protocol (Sharma et al., 2011). Briefly, clubbed roots were washed and soaked in deionoized
water, and 10 g of gall was homogenized in 300 mL deionized water for 2 min in an electronic
blender. The mixture was filtered though eight layers of cheesecloth and a haemocytometer was
used to estimate the resting spore concentration. The resulting spore suspension was diluted to
create the required treatment concentration.
In each replicate, the height of each plant was assessed at 7-day intervals from 2 to 6
weeks after inoculation (WAI). In repetition 2, the height measurement for week 5 was missed by
mistake. At 10 weeks after planting, plants were harvested and weighed, and roots were assessed
for clubroot incidence (%) and severity using the standard 0–3 rating scale, where 0 = no
34 clubbing, 1 < 1/3 of root clubbed, 2 = 1/3–2/3 of roots clubbed and 3 > 2/3 of roots clubbed
(Strelkov et al., 2006). A Disease Severity Index was calculated according to Crete et al. (1963):
DSI = ∑ [(c [(class no.)(no. of plants in each class)] x100 [(total no. plants per sample)(no. classes - 1)]
Plant height was measured from hypocotyl (at soil surface) to shoot apex. Plant height
measurements from the controlled environment studies were compared at individual time-points
and also for overall growth over time. Growth over time was analyzed using area under the
growth stairs (AUGS). The original model for this analysis is area under the disease progress
curve (AUDPC), which is commonly used as a quantitative measure of disease over time (Van
der Plank, 1963). It is calculated from multiple disease assessments over time (Jeger and
Viljanen-Rollinson, 2001). In the current study, plant height assessments were used instead of
disease assessment at each time-point. In area under the growth curve (AUGC), the first and last
time-points are only weighted at 50%, because the trapezoid only extends in one direction from
the midpoint. Area under the Growth Stairs (AUGS) was used in this study, which provides equal
weighting of all time-points to summarize season-long growth into a single value, calculated as
follows:
AUGS =Y! + Y!!!
2 !!!
!!!
× t!!! − t! +Y! + Y!
2 × t! − t!n− 1
where Yi is plant height in cm at the ith observation, ti is time in days after inoculation at the ith
observation, and n is the total number of observations (Simko and Piepho, 2012).
35 2.2.2. Large pot studies – outdoors
Outdoor studies in large pots were conducted as a randomized complete block design
(RCBD) with four replicates per treatment at Bajar Farm, near the Muck Crops Research Station,
King, Ontario. The clubroot-resistant canola cv. 45H29 (Pioneer Hi-Bred, Caledon, ON) and
ACS-N39 (AAFC, Saskatoon, SK), a susceptible canola check, were seeded into 200-cell plug
trays on 16 July 2014 and grown in a greenhouse. The seedlings were hand transplanted into
plastic pots (30 cm × 27.5 cm dia.) containing dark grey gleysol-Granby sandy loam soil
(Westerveld, 2005) from the Bajar farm site on 06 August 2014, at 3 weeks after seeding. When
the trial was conducted in 2015, one repetition was seeded on 09 June and another on 24 June.
The seedlings were hand transplanted into pots containing 75% mineral soil from the Bajar farm
and 25% soil-less media (Sunshine Mix #4, Sun Gro Horticulture Canada Ltd., Agawam, MA) on
23 June and 08 July, at 2 weeks after seeding. The soil-less media was mixed into the field soil
sample using an electrical cement mixer.
The soil pH in each pot was lowered from 7.8 to 6.4 by application of 7.3 g 90% sulphur
chips, 2.9 g nitrogen sulphate, and 1.0 g magnesium sulphate and 400 mL water pH adjusted to
4.0 with phosphoric acid. The pots were watered once per week with water adjusted to pH 6.0
using commercial vinegar and N-P-K (20-20-20) solution. Inoculum was prepared as described
previously (Sharma et al., 2011). Each pot was inoculated at transplanting with 450 mL of spore
suspension of pathotype 6 to produce a final concentration of 0, 1 x 103, 1 x 104, 1 x 105, 1 x 106
and 1 x 107 resting spores g-1soil in the top 15 cm of the pot. In 2014, each pot was inoculated at
6 weeks before transplanting and again at transplanting. In 2015, each pot was inoculated one
time at transplanting.
The height of each resistant plant was assessed at weekly intervals starting at 2 WAI. At 8
WAI, the developmental stage (vegetative, bud, flowering, pod development) of each plant was
36 assessed. Plants were then harvested, weighed and roots assessed for clubroot incidence (%) and
severity using a standard 0-3 rating scale, as previously described.
Each experimental unit consisted of two pots with five plants per pot, except for the
treatments in the fourth replicate in 2014, which only had one pot per unit because there were not
enough seedlings for transplant. Biomass assessments included five plants per experimental unit.
Resistant canola plants that exhibited clubbing symptoms were removed from the biomass
assessments in the second repetition in 2015, but were included in all of the plant height
assessments in both years and in the biomass assessments for the first repetition in 2015.
2.2.3 Controlled environment study of canola, cabbage and napa cabbage
Three clubroot-resistant canola cultivars, 45H29, 73-67 (Monsanto Canada Inc.,
Winnipeg, MB) and 73-77 (Monsanto Canada Inc.), and the susceptible line ACS-N39 were
sown in tall plastic conetainers filled with mineral soil from Elora Research Station (Elora, ON).
In companion studies, three clubroot-resistant napa cabbage cultivars, China Gold (Sakata Seed
Corporation, Morgan Hill, CA), Yuki (Sakata Seed Corporation) and Emiko (Bejo Seeds Inc.,
Oceano, CA), and the susceptible cultivar Mirako (Bejo Seeds Inc.) and three clubroot-resistant
cabbage cultivars, Kilaherb (Syngenta Seeds, Inc., Minnetonka, MN), Kilaton (Syngenta Seeds,
Inc.), and Tekila (Syngenta Seeds, Inc.), and the susceptible cultivar Bronco (Bejo Seeds Inc.)
were assessed.
For each study, two seeds were planted per pot and thinned to one seedling per pot. Four
replicates of 10 pots were included for each treatment. The plants were maintained at 25°/20° C
day/night, with 16-hr photoperiod and 65% relative humidity, and fertilized with 1 g L-1 N-P-K
(20-20-20) and 1 g L-1 magnesium sulphate at 2–3 day intervals. The plants were watered with
deionized water adjusted to pH 6.0 using commercial white vinegar. Each 10-day-old seedling
37 was inoculated with 5 mL of resting spore suspension of 1 x 106 resting spores mL-1 of
P. brassicae pathotype 6. Inoculum was prepared as described above. In each replicate, the height
of each plant was assessed at 7-day intervals starting 2 WAI. Each study was repeated with the
following modifications: 25% soil-less media (Sunshine Mix #4, Sun Gro Horticulture Canada
Ltd., Agawam, MA) was incorporated into the soil blend with 75% mineral soil, and the mixture
was treated with water adjusted to pH 4.0 using phosphoric acid to lower the pH of the soil at 2
weeks prior to planting.
2.2.4 Field trials
Field trials were conducted to compare the growth of clubroot-resistant canola and napa
cabbage cultivars at two adjacent sites at the Muck Crops Research Station, Holland Marsh,
Ontario. Both sites were on organic soil, a typic humisol-muck (McDonald et al., 2008), and
received the same agronomic treatments (tillage and fertilization) and had a similar crop history,
but differed in concentration of resting sproes of P. brassicae (Table 2.1).
In one trial, three clubroot-resistant canola cultivars, 45H29, 73-67 and 73-77, and the
susceptible line ACS-N39 were sown on 26 June, 2014. In a companion trial, three clubroot-
resistant napa cabbage cultivars, China Gold, Yuki and Emiko, and the susceptible cultivar
Mirako were sown on 16 July, 2014. Each trial was laid out in a randomized complete block
design with four replicates. Both crops were direct seeded at a rate of approximately 18 seeds per
m of row using an Earthway push seeder fitted with an Earthway 1002-9 disc. Each canola plot
was 6 m in length and each napa cabbage plot was 3 m in length. Napa cabbage plants were
thinned to approximately 25 cm apart within rows. Each plot consisted of a single row, spaced
~26 cm apart. In each replicate, a representative sample of 10 consecutive plants starting 2-3
plants in from the beginning of the row was measured repeatedly at 7-day intervals starting 4
38 weeks after seeding. Plant growth was assessed in canola by measuring plant height from the
hypocotyl to the shoot apex. Napa cabbage plant growth was assessed by measuring the length of
the third and fourth youngest leaves. AUGS was calculated as previously described.
Table 2.1 Soil properties of field trial sites at the Muck Crops Research Station at Holland
Marsh, Ontario in 2014 and 2015.
Site ID Spores
g-1 soil pH Soil analysis Texture
P K Mg OM
2014-Low 7 x 105 6.3 67 240 381 78 Muck
2014-High 7 x 106 6.5 52 195 408 75 Muck
2015-Low 3 x 106 6.1 79 291 493 77 Muck
2015-High 1 x 107 6.3 84 100 463 79 Muck
2015-BDL1 BDL 6.0 112 548 495 82 Muck
1BDL = Below Detection Limit
At 9 weeks after planting, the developmental stage (vegetative, bud, flowering, pod
development) of each canola plant was assessed. Plants were harvested and weighed, and roots
were assessed for clubroot incidence (%) and severity using a standard 0-3 rating scale, as
previously described.
Both trials were repeated in 2015, with the addition of an adjacent site in each trial with
inoculum concentration below detectable levels (Table 2.1). All plots were 6 m in length, and
each plot for both canola and napa cabbage was seeded on 09 July 2015 to allow for comparison
39 across crop species. Also, the susceptible napa cabbage cultivar Mirako was replaced with
Suzuko B-2961 (Bejo Seeds, Inc.) due to discontinuation of Mirako.
2.2.5 Controlled environment pH study
Growth room studies were conducted using the susceptible line ACS-N39 in a factorial
design with two factors. One factor was inoculum concentration (0, 1 x 103, 1 x 105, 1 x 107
spores mL-1) and the other factor was pH (5.5, 6.0, 6.5, 7.0, 7.5). The experimental design was a
RCBD with three replicates. Three plants were grown per experimental unit as outlined in the
protocol by Kasinathan (2012) for assessment of root hair colonization. Plant culture followed the
protocol outlined by Kasinathan (2012), which were adapted from Donald and Porter (2004).
Seeds were sown in moist sand in Petri dishes at 20°C with 16-h photoperiod. When the
cotyledons were fully expanded, individual seedlings were transplanted into 50-mL Falcon tubes
(Fisher Scientific, Markham, ON) containing autoclaved, non-calcareous sand at pH 6.5
(Hutcheson sand, Hutcheson Sand Mixes, Huntsville, ON). The plants were fertilized with
15:15:18 N:P:K and ammonium sulphate. A stock solution was prepared using 40 g of N:P:K and
20 g of ammonium sulphate in 1 L of water. A 5-mL aliquot of stock solution was added to 1 L
of deionized water and the pH was adjusted before watering as described below. The plants were
watered with deionized water adjusted to the desired pH using biological buffers (Myers and
Campbell, 1985): PIPES [piperazine-N, N’-bis-(2-ethanesulfonic acid,) monosodium salt, mono
hydrate], MES [2-(N-morpholino) ethanesulfonic acid, sodium salt] and HEPES [N-2-
hydroxyethylpiperazine-N’-2-ethanesulphonic acid] (Robiot Canada, Toronto, ON). Stock
solutions were prepared and stored at 5°C. The pH meter (Hanna instruments, Woonsocket, RI)
was calibrated before each use, using standard buffer solutions (Fisher Scientific, Nepean, ON).
40 To achieve the desired pH of the sand growth medium, solution of the target pH was added to
each Falcon tube. The water that drained from the Falcon tubes was collected and the pH of the
drain solution was measured for each treatment. Based on this result, the pH of the watering
solution was adjusted up or down and the watering solution was re-applied until the desired pH
was attained in the drain solution. Drainage holes were made at the bottom of each Falcon tube to
avoid water logging.
Each 10-day-old seedling was inoculated with 1 mL of resting spore suspension.
Inoculum was prepared following the protocol outlined by Sharma et al. (2011) as described
above. Plants were harvested at 8, 15 and 22 days after inoculation (DAI). Roots (including root
hairs) were sampled from all three plants in one experimental unit for one biological replicate.
Clubroot incidence and severity were assessed using the 0–3 scale previously described. The
roots were rinsed with deionized water, followed by surface sterilization in 10% bleach, rinsed
three times in deionized water to remove debris from the root surface, cut into 1-cm pieces,
frozen in liquid nitrogen, and stored at -80oC until use, as described in Lahlali et al. (2011). This
study was not repeated.
2.2.6 Statistical analysis
All analyses were conducted using SAS version 9.3. The data for the controlled
environment study and large pot outdoor study were analyzed in a mixed model analysis of
variance using single-degree of freedom contrasts using PROC MIXED and PROC GLM. Spore
concentration was a fixed effect and block and repetition were random effects.
The data for field trials and studies of canola, napa cabbage and cabbage were analyzed as
a mixed model analysis of variance using single degrees of freedom and the slice option to
41 examine the interaction between cultivar and site (spore concentration) with PROC MIXED and
PROC GLM. Levene’s test was also used to test for homogeneity of variance across sites.
Variance in plant height, biomass and maturity of resistant canola across sites within each year
was homogeneous (Tables A1.35-37, A1.41-43, A1.50-52), so comparisons were made among
sites for those response variables. Variance in plant height was homogenous across sites in 2015
but not in 2014. Therefore, plant height, biomass and maturity provide a more robust indication
of cost of resistance to clubroot at various concentrations of resting spores than plant height in the
2014 field trial. There was homogeneity of variance in CI and DSI for susceptible canola among
sites within year. Variance was not homogeneous across years for any variable, so each year was
analyzed separately. Means separation were conducted using Tukey’s test at P = 0.05 level of
significance. The fixed effects in the mixed model analysis were cultivar, crop and spore
concentration and random effects were block and repetition.
The data for the pH study were analyzed in a mixed model analysis of variance, with
spore concentration and pH as fixed effects, and block and repetition as random effects. Clubroot
incidence and disease severity index for the pH treatments at each concentration of resting spores
were compared using Tukey’s procedure as described by Kasinathan (2012).
2.3 Results 2.3.1 Controlled environment study
There were high rates of clubroot incidence and severity in the susceptible control and no
clubroot symptoms in resistant canola plants in both repetitions under controlled conditions
(Table 2.2). One additional repetition was conducted, but it was not included in the analysis due
to inconsistent plant growth and signficant block effect (P <0.0001) (see data in appendix).
42 Table 2.2 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible
canola ACS-N39 inoculated with 1 x 106 spores mL-1 under controlled conditions1.
Repetition CI DSI
1 100 ns2 71 b3
2 100 95 a 1 N.B. There were no clubbing symptoms in any resistant plants (data not shown).
2 ns = not significant. 3 Columns with the same letter do not differ at P = 0.05, based on Tukey’s multiple means
comparison test.
Plant height of canola cv. 45H29 (resistant) at 6 WAI declined in a quadratic relationship
(y = -0.086x2 + 0.129x + 31.7, R² = 0.21) with increasing concentration of resting spores
(P = 0.005) across the two repetitions (Fig. 2.1). Plant height was reduced by approximately 13 %
at conecentrations > 1 x 106 spores mL-1 compared to the non-inoculated control at 6 WAI, based
on the regression equation. There was no repetition effect on plant height at 6 WAI and therefore
data were analyzed across repetitions.
The biomass of resistant canola (dry shoot weight) declined in a weak linear relationship
(y = 0.189x + 6.47, R2 = 0.07, P = 0.03) with increasing inoculum in the first repetition. The
proportion of resistant canola plants at the seedpod development stage at 8 WAI declined in a
quadratic relationship (y = -1.63x2 + 9.37x + 78.2, R2 = 0.60) with increasing concentration of
resting spores (P = 0.007) in the first repetition (Fig. 2.2). The regression was not significant for
biomass or maturity in the second repetition.
43
Figure 2.1 Plant height of clubroot-resistant canola at 6 weeks after inoculation with increasing
concentrations of Plasmodiophora brassicae resting spores under controlled conditions (two
repetitions, n=8, P = 0.005).
Figure 2.2 Plant maturity at harvest in canola grown in controlled conditions in response to
increasing concentration of Plasmodiophora brassicae resting spores (P = 0.007) in the first
repetition.
y = -‐0.086x2 + 0.129x + 31.7 R² = 0.21, P = 0.005
0
5
10
15
20
25
30
35
40
0 1 2 3 4 5 6 7 8
Plant height (cm
)
Resting spore concentration, log scale (spores mL-‐1)
y = -‐1.63x2 + 9.37x + 78.2 R² = 0.60, P = 0.008
0 10 20 30 40 50 60 70 80 90 100
0 1 2 3 4 5 6 7 8
Plants at pod stage (%
)
Resting spore concentration, log scale (spores mL-‐1)
44 2.3.2 Large pot studies – outdoors
Clubroot incidence and severity in the susceptible control was low in 2014, but high in
both repetitions in 2015 (Table 2.3). There were no clubbing symptoms in resistant canola in
2014, but there were low levels of clubroot incidence and severity in both repetitions in 2015
(Table 2.3). There was no effect of inoculation on plant height, maturity or biomass among the
treatments in 2014 or 2015 (data not shown).
In the 2 weeks following transplanting of canola in 2014, mean air temperature was 25°C
maximum and 12°C minimum. In 2015, mean air temperature was 23°C maximum and 15°C
minimum in the first repetition and 27°C maximum and 14°C minimum in the second repetition.
Table 2.3 Clubroot incidence (CI) and severity (disease severity index, DSI) in canola breeding
line ACS-N39 (susceptible check) and resistant cultivar 45H29 in an outdoor trial using large
pots near Bradford, ON, 2014 and 2015.
ACS-N39 45H29
Assessment CI DSI CI DSI
2014 35 b1 16 c 0 b 0 ns2
2015 Repetition 1 98 a 78 b 8 ab 6
2015 Repetition 2 100 a 97 a 15 a 12 1 Means in a column followed the same letter do not differ at P = 0.05, based on Tukey’s multiple
means comparison test. 2 ns = not significant.
45 2.3.3 Controlled environment study – canola, napa cabbage and cabbage
Clubroot incidence and severity were low in canola and moderate in napa cabbage and
cabbage in the first repetition of the study under controlled conditions. Clubroot symptoms were
low in cabbage and moderate in canola and napa cabbage in the second repetition (Table 2.4).
In resistant napa cabbage, inoculation with P. brassicae resting spores reduced leaf length
at 5 WAI by 11.4% (±6.5) compared with the non-inoculated control (P = 0.005) across
repetitions, but there was no effect of inoculation on plant height of resistant canola or leaf length
of resistant cabbage. There were no differences in the timing of maturity in resistant canola or in
biomass of any crop species (data not shown).
Table 2.4 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible
canola, napa cabbage and cabbage inoculated with 1 x 106 spores mL-1 P. brassicae, under
controlled conditions. 1
Repetition
Canola Napa cabbage Cabbage
CI DSI CI DSI CI DSI
1 18 b2 6 b 39 a 20 ns3 52 a 20 a
2 63 a 35 a 30 ab 21 23 b 10 ab 1 N.B. There were no clubbing symptoms in the non-inoculated control or in any resistant plants
(data not shown). 2 Columns with the same letter do not differ at P = 0.05 based on Tukey’s multiple means
comparison test. 3 ns = not significant.
46 2.3.4 Field trials
The concentration of resting spores site at the low spore concentration site in 2014 was
90% lower than the concentration of resting spores at the high spore concentration site. However,
spore concentrations at both sites were much higher than the infection threshold for susceptible
cultivars (103) (Faggian and Strelkov, 2009). In 2014, there was 100% CI and DSI in the
susceptible canola at both sites. Susceptible napa cabbage had 23% CI and 13% DSI at the low
spore concentration site, and 90% CI and 44% DSI at the high spore concentration site (Table
2.5). There were no clubbing symptoms in resistant canola or napa cabbage in 2014.
Table 2.5 Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible
canola and napa cabbage grown in field soil at Muck Crops Research Station, 2014.
Canola Napa cabbage
Site ID Spores
(g-1 soil)
CI DSI CI DSI
2014-High 7 x 106 100 ns1 100 ns 90 a2 44 a
2014-Low 7 x 105 100 100 23 b 13 b 1 N.B. There were no clubbing symptoms in the non-inoculated control or in any resistant plants
(data not shown). 2 ns = not significant. 3 Columns with the same letter do not differ at P = 0.05, based on Tukey’s multiple means
comparison test.
In 2015, there was 100% CI in the susceptible canola cultivar at the high and low spore
concentration sites, and 45% at a site with spore concentration below detectable limits. DSI in
susceptible canola was 70% at the high spore concentration site, 65% at the low spore
47 concentration site and 19% at the site with spore concentration below detectable limits (Table
2.6). Resistant canola in 2015 at all three sites developed low levels of clubroot symptoms (Table
2.6). There were no clubbing symptoms in any napa cabbage cultivars in 2015. This indicates that
in these conditions there was no susceptible control for napa cabbage. There were no clubroot
symptoms observed in cultivar Suzuko B-2961 following inoculation with clubroot pathotype 6
in a follow-up assessment under controlled conditions, although the seed company did not state
that this cultivar was clubroot-resistant.
Table 2.6. Clubroot incidence (CI) and severity (disease severity index, DSI) in susceptible and
resistant canola cultivars, grown in field soil at sites with high, lower and no measurable
concentration of resting spores at the Muck Crops Research Station, 2015.
Susceptible Resistant
Site ID Spores
(g-1 soil)
CI DSI CI DSI
2015-High 1 x 107 100 a1 70 a 6 ns2 3 ns
2015-Low 3 x 106 100 a 65 a 3 1
2015-BDL < 1000 45 b 19 b 3 1 1 Columns with the same letter do not differ at P = 0.05 based on Tukey’s multiple means
comparison test. 2 ns = not significant.
BDL = below detection limit.
The plant height of the resistant canola cultivars in 2014 was 30% (±1.3) shorter at the
high spore concentration site compared with the low spore concentration site at 8 weeks after
seeding (WAS) (P < 0.0001, Fig. 2.4). The biomass of resistant canola was 43% (±14.3) lower at
sites with high concentrations of resting spores (P = 0.006, Fig. 2.4). The proportion of resistant
plants at the pod development stage of plant maturity was reduced by 75% (±12.5) at the high
48 spore concentration site compared with the low spore concentration site at 8 WAS (P < 0.0001,
Fig. 2.4). There was no effect of resting spore concentration on plant height, biomass or maturity
in resistant canola in 2015 (Fig. 2.5).
49
Figure 2.3 Plant height of three resistant canola cultivars at the Muck Crops Research Station,
Ontario, at sites with high, lower and undetectable resting spore concentrations, 2014 and 2015.
0 10 20 30 40 50 60 70 80 90 100 110
4 5 6 7 8
0 10 20 30 40 50 60 70 80 90 100 110
4 5 6 7 8
Plant height (cm
)
Weeks after seeding
0 10 20 30 40 50 60 70 80 90 100 110
4 5 6 7 8 Weeks after seeding
0 10 20 30 40 50 60 70 80 90 100 110
4 5 6 7 8
Plant height (cm
)
Weeks after seeding
0 10 20 30 40 50 60 70 80 90 100 110
4 5 6 7 8
Plant height (cm
) 45H29 73-‐67 73-‐77
2014-H
2015-L 2015-BDL
2015-H
2015-H
2014-L
50
Figure 2.4 Plant height at 8 weeks after seeding (WAS), biomass at harvest and maturity at 8
WAS for resistant canola cultivars grown in field soil with 7 x 105 spores g-1 soil (Low) and
7 x 106 spores g-1 soil (High) at Muck Crops Research Station, 2014. Bars with the same letter do
not differ at P = 0.05 based on Tukey’s multiple means comparison test.
Figure 2.5 Plant height and maturity at 8 weeks after seeding (WAS), biomass at harvest for
resistant canola cultivars grown in field soil with <1000 spores g-1 soil (BDL), 3 x 106 spores g-1
soil (Low) and 1 x 107 spores g-1 soil (High) at Muck Crops Research Station, 2015. Bars with the
same letter do not differ at P = 0.05 based on Tukey’s multiple means comparison test.
a
b
0
10
20
30
40
50
60
70
Plan
t hei
ght (
cm)
Plant Height
Low High
a
b
0
100
200
300
400
500
600
Shoot w
eight of 10 plants (g)
Biomass
Low High
a
b
0
10
20
30
40
50
60
70
Plants at pod stage (%
)
Maturity
Low High
a a a
0
20
40
60
80
100
120
Plant height (cm
)
Plant Height
BDL Low High
a a
a
0
100
200
300
400
500
600
Shoot w
eight 10 plants (g)
Biomass
BDL Low High
a a a
0 10 20 30 40 50 60 70 80 90 100
Plants at pod stage (%
)
Maturity
BDL Low High
51 Leaf length of resistant napa cabbage in was 31% (±15.5) shorter in 2014 and 22% (±1.0)
shorter in 2015 at the high spore concentration site relative to the low spore concentration site at
8 WAS (P < 0.0001). There was no reduction in leaf length between the low spore concentration
stie and site with spore concentration below detectable limits in 2015 (Fig. 2.6). Biomass of napa
cabbage was 34% (±15.9) lower at the high spore concentration site compared with the low spore
concentration site at harvest in 2014 (P = 0.007). There were no differences in biomass in 2015,
but the trend in biomass was similar to that in leaf length (Fig. 2.7).
Figure 2.6 Leaf length and biomass of napa cabbage at sites with 7 x 105 spores g-1 soil (Low)
and 7 x 106 spores g-1 soil (High) in field trials at Muck Crops Research Station, Ontario, 2014.
Bars with the same letter do not differ at P = 0.05 based on Tukey’s multiple means comparison
test.
a
b
0
5
10
15
20
Leaf length (cm)
Leaf length
Low High
a
b
0 100 200 300 400 500 600 700 800 900 1000
Dry shoot weight (g)
Biomass
Low High
52
Figure 2.7 Leaf length and biomass of napa cabbage at sites with <1000 spores g-1 soil (BDL), 3
x 106 spores g-1 soil (Low) and 1 x 107 spores g-1 soil (High) in field trials at the Muck Crops
Research Station, Holland Marsh, Ontario, 2015. Bars with the same letter do not differ at
P = 0.05 based on Tukey’s multiple means comparison test.
In the 2 weeks following seeding of canola in 2014, the total rainfall was 6 mm with no
more than 4 mm in any single day. Mean air temperature was 29°C maximum and 14°C
minimum. In contrast, in the 2 weeks following napa cabbage seeding in 2014, the total rainfall
was 37 mm, of which 35 mm occurred over 2 days. In 2015, both canola and napa cabbage were
seeded at the same date. In the 2 weeks following seeding, the total rainfall was 8 mm with no
more than 4 mm in any single day. Mean air temperature was 27°C maximum and 14°C
minimum.
2.3.5 pH study
At 8 days after inoculation (DAI), there were no clubbing symptoms on any plant. At
15 DAI, clubroot severity increased as the concentration of resting spores increased (P < 0.0001),
but there was no interaction or main effect of pH. At 22 DAI, there was an interaction effect
a a
b
0
5
10
15
20
25
30
Leaf length (cm)
Leaf length
BDL Low High
a a
a
0
200
400
600
800
1000
1200
1400
Dry shoot weight (g)
Biomass
BDL Low High
53 (P = 0.03) for CI response to spore concentration and pH, but no interaction for DSI. There was
an increase in clubroot severity with increasing spore concentration (P < 0.0001) and an effect of
pH on clubroot severity (P = 0.01). Inoculation with 1 x 103 spores mL-1 at pH 6.0-7.0 resulted in
increased clubroot incidence relative to 1 x 103 spores mL-1 at pH 5.5 and 7.5, at 22 DAI. In
contrast, there was no effect of pH on clubroot incidence at 1 x 105 and 1 x 107 spores mL-1 (Fig.
2.8). At harvest (35 DAI), increasing concentration of resting spores increased CI and DSI
(P < 0.0001). There was no interaction between pH and inoculum concentration and no effect of
pH on CI or DSI (Fig. 2.8). There was no interaction for DSI at any time-point (data not shown).
Figure 2.8 Clubroot incidence (CI) in susceptible canola grown under controlled conditions at a
range of pH (5.5 to 7.5) and concentrations of resting spores of P. brassicae at 22 days after
inoculation (DAI).
0 10 20 30 40 50 60 70 80 90 100
0.0 1x10^3 1x10^5 1x10^7
Clubroot incidence (%
)
Spores mL-‐1
5.5 6 6.5 7 7.5 pH
54 2.4 Discussion
This study examined the hypothesis that there was a cost of resistance to P. brassicae.
The hypothesis was assessed in field trials, in large pots outdoors and under controlled
conditions. Overall, there was a metabolic cost of resistance at high spore concentrations in
canola and napa cabbage, but the results were inconsistent in pots, whether outdoor or under
controlled conditions. In the field trials, plant growth of canola and napa cabbage was generally
reduced and maturity was delayed at high spore concentrations.
The biomass of resistant canola was 43% lower when spore concentration was increased
by 10-fold (from 7 x 105 to 7 x 106 spores g-1) in the 2014 field trial at the Muck Crops Research
Station. A similar trend was observed for plant height, although variance was heterogeneous
across sites, indicating that there may have been an influence on plant height in addition to
P. brassicae. Crop development was also delayed, with the proportion of plants at the pod stage
reduced by 75% at 8 weeks after seeding. However, there was no effect of resting spore
concentration on plant height, biomass or maturity in 2015.
There were similar concentrations of resting spores in the sites with high spore
concentrations in 2014 (7 x 106 spores g-1) and 2015 (1 x 107 spores g-1). However, clubroot
severity was much lower in 2015 (DSI 70) relative to 2014 (DSI 100). Rainfall and air
temperature in the 3 weeks after seeding were similar in both years. Soil moisture may have had a
greater effect than rainfall on clubroot development in muck soil (Thuma et al., 1983; Cranmer,
2015). However, soil moisture was not recorded in this trial. Another difference between years
was that damage from Swede midge (Contarinia nasturtii) was higher in 2014 than in 2015. It is
possible that the metabolic cost of resistance to clubroot is larger when other biotic stresses are
present.
55 Plant height of canola cv. 45H29 (resistant) decreased by 12-14% at very high resting
spore concentrations (1 x 107 and 1 x 108), but not at 1 x 106 spores ml-1 under controlled
conditions. Similarly, biomass declined and maturity was delayed at 6 WAI in one of two
repetitions. However, there was no effect of resting spore concentration on plant height, biomass
or maturity in outdoor pot studies in 2014 or 2015. Clubroot incidence and severity were low in
the susceptible control in 2014. This was likely related to late transplanting of canola, so the
mean nighttime temperatures (12°C) were not warm enough for the development of clubroot
symptoms (McDonald and Westerveld, 2008; Gossen et al., 2012; Cranmer, 2015). In 2015,
canola seedlings were transplanted earlier in the season and were slightly younger when
transplanted (2- rather than 3-wks-old). Warmer nighttime air temperature (14-15°C) followed
inoculation, and clubroot incidence and severity were high. Previous research has demonstrated
that cool temperatures can slow pathogen colonization and reduce clubroot symptom
development (Gossen et al., 2012). Also, inoculation of older seedlings can result in slightly
lower levels of infection (Hwang et al., 2011b). However, low temperature was likely the most
important factor in the low levels of clubroot in 2014.
In general, reductions in plant height of resistant canola were apparent at 5 to 6 weeks
after inoculation (WAI) in indoor studies and 8 weeks after seeding (WAS) in field trials. In a
previous study under ideal conditions, restriction of pathogen development occurred from 2-4
weeks after inoculation (Deora et al., 2013). In the current study, that time-point corresponded
with initiation of flowering. From flowering through pod development, canola is highly sensitive
to abiotic stresses such as drought (Champolivier and Merrien, 1996). Abiotic and biotic stresses
may activate similar pathways for plant response. For example, it has been proposed that
synthesis of jasmonic acid (JA) from linolenic acid in a lipoxygenase-dependent process can be
initiated by abiotic and biotic stresses in soybean (Glycine max (L.) Merr.) (Creelman and Mullet,
56 1995). Thus, information about the timing of drought response could be helpful for understanding
response to P. brassicae during the flowering stage of development in canola.
More than one crop species was included in the current study to examine the effect of
resting spore concentration on plant growth of different Brassica species. Growth of napa
cabbage declined in sites with a high spore concentration in field trials and under controlled
conditions. In field conditions with a high concentration of resting spores, leaf length of resistant
napa cabbage grown was 31% shorter in 2014 and 22% shorter in 2015 relative to a site with a
lower spore concentration. This was consistent with one of two studies under controlled
conditions, where leaf length was 11% shorter in resistant napa cabbage inoculated with 1 x 106
spores mL-1. Growth of canola declined in sites with a high spore concentration in field trials.
However, under controlled conditions, canola did not exhibit reduction in plant height or biomass
at 1 x 106 spores mL-1 and cabbage did not exhibit reduction in leaf length or biomass (data not
shown).
Brassica vegetables have a variety of different sources of, and mechanisms of, resistance
to clubroot (Rahman et al., 2014; Chu et al., 2015; Gludovacz et al., 2015). It is possible that
there are differences in the metabolic cost of resistance to P. brassicae among species, which
may contribute to more consistent observation of cost of resistance in napa cabbage compared
with canola. It is also possible that leaf length may be more strongly correlated with cost of
resistance than plant height, because leaf area affects photosynthetic capacity, whereas plant
height does not (Freyman et al., 1973; Krogman and Hobbs, 1975; Campbell and Kondra, 1978).
Seed yield of canola was not assessed in the current study because canola was seeded late in the
season to optimize environmental conditions for pathogen development, such as warmer air and
soil temperature (Kasinathan et al., 2011a,b; Gossen et al., 2013; Cranmer, 2015). In addition,
57 there were other challenges to yield assessment at the Muck Crops Research Station, incuding
small plot size, damage from insect pests (Swede midge) and birds feeding on canola seed.
Field trials were arranged to allow for comparisons between resistant cultivars and the
susceptible line within one site. In 2015, one site with a spore concentration below the detection
limit was added as a clubroot-free control. Even though low levels of clubroot developed at that
site, it had a much lower (>1000-fold) concentration of resting spores relative to other sites.
The studies involving inoculated field soil generally did not exhibit a cost of resistance
response. In some indoor studies, plant height was slightly shorter in inoculated plants than the
non-inoculated control, but the differences were not significant. The effect of resting spore
concentration on resistant plants may have been influenced by displacement of the field soil into
pots. In a previous study, compaction and saturation of soil in pots resulted in increased clubroot
symptoms compared with drained soil (Kasinathan, 2012). However, the effect of soil saturation
on clubroot development is complex. Results from other studies have indicated that excessive soil
moisture may suppress pathogen development (Thuma et al., 1983; Cranmer, 2015), which may
be related to over-dilution of the spore concentration in soil or downward movement of inoculum
to the bottom of the pot. Conversely, much of the inoculum may have been trapped close to the
soil surface and never reached plant roots.
In the current study, clubroot symptoms were higher in the second repetition of the indoor
study, when 25% soil-less media was incorporated into field soil and a lower pH was maintained
using acetic acid. Soil at pH 6.0-6.5, which is favourable for infection and clubroot development,
may lead to increased fitness costs in resistant cultivars compared with soil pH >7.0, at a specific
concentration of resting spores. Furthermore, plant-to-plant variability within the containers may
have reduced the ability to separate out the cost of resistance from other variance in plant growth.
One possible reason that the field trials provide a much stronger and more consistent response to
58 clubroot is that only a relatively small number of resting spores were applied to the pots in the
inoculated trials. These may be highly concentrated in small areas of the root system, and it is
possible that large parts of the root system in older plants were not subject to continued infection
(Hwang et al., 2011b). However, infection can occur on new roots that develop throughout the
season, although young plants are more susceptible to infection and the rate of infection
decreases over time (Hwang et al., 2012, Fei et al., 2016).
There were several unexpected challenges in field trials. In the 2015 field trial, the napa
cabbage cultivar Suzuko B-2961 was seeded in place of the susceptible control Mirako, which
was seeded in 2014 but had been discontinued by the seed company. However, this new cultivar
turned out to be resistant to pathotype 6. Thus, susceptible canola ACS-N39 was used as a
substitute for the susceptible control for napa cabbage in 2015.
There was an unexpected increase from no clubroot symptoms in resistant canola in 2014
to the low levels of clubroot incidence and severity observed in 2015, but not in resistant napa
cabbage. High concentrations of resting spores in soil can result in selection of more aggressive
isolates of the pathogen (Kiyosawa, 1982; LeBoldus et al., 2012). It is possible that a shift in the
pathotype is occurring at Muck Crops Research Station that affects the resistance in the canola
cultivars but not napa cabbage. Alternatively, there may be have been susceptible off-types or
susceptible volunteer canola from a previous crop.
Susceptible canola grown in 2015 with 65-70 DSI was much shorter than susceptible
canola grown in 2015 with 100 DSI. These results support a previous report that plant height of
susceptible canola decreased with increasing clubbing symptoms, from 74 to 100 DSI (Hwang et
al., 2011). However, damage from swede midge (Bardner et al., 1971) may also have contributed
to the shorter plant height of susceptible canola in 2014.
59 The metabolic basis for a cost of resistance may be explained based on previous research
investigating the stages of pathogen development during which the resistance response takes
place and the metabolic pathways involved in host resistance to clubroot. Recognition and
induction of resistance takes place during root hair infection (McDonald et al., 2014). Plant host
recognition takes place when pathogen recognition receptors (PRRs) recognize conserved
pathogen-associated molecular patterns (PAMPs). Defense mechanisms may be activated by
endogenous signals that result from effectors released by the pathogen, or by microbial activity
that trigger the appropriate responses (Pal and McSpadden-Gardener, 2006). Pathogen
development is then halted or greatly restricted during cortical infection, which occurs 22-28
days after germination of resting spores under optimal conditions (Deora et al., 2012). In the
current study, this was 1-2 weeks before reductions in plant growth were observed under
controlled conditions. Although the cost of resistance is observed later, there may be an effect on
resource allocation occurring within the plant at an earlier stage that is not observable using the
indicators of growth and development in the current study. Also, there may be new infections
occurring later in the season on new root growth.
A recent study demonstrated up-regulation of signaling and metabolism of jasmonic acid
and ethylene, callose deposition and indole-containing compounds in inoculated plants of
clubroot-resistant B. rapa carrying the Rcr1 gene (Chu et al., 2015). Salicylic acid was not
elevated, and auxin and a chitinase-like protein were down-regulated. Clubroot-resistant B. napus
canola responded to secondary infection with P. brassicae by forming a concentrated reactive
oxygen species (ROS) ring in the inner cortex (Deora et al., 2013), which may contribute to
structural reinforcement of the host cell wall. ROS generation takes place in cellular
compartments such as mitochondria or chloroplasts, and involves several enzymes (Abel and
60 Hirt, 2004). ROS mediates ABA-induced stomatal closure and can react with proteins to decrease
enzyme activity (Moller, 2001). The mitochondrial electron transport chain (ETC) is a major site
of ROS production, which under normal conditions is responsible for oxidative phosphorylation,
the metabolic pathway that converts NADH into energy that is used for plant growth (Moller,
2001). Stress-induced co-expression of specific AOX (alternative oxidase) and NDH (NADH
dehydrogenase) genes in Arabidopsis (Clifton et al., 2005) may play a role in decreasing the
respiration function and leaf growth of plants in response to abiotic stress (Sevilla et al., 2015).
However, a decrease in classical pathway activity may have a greater effect than the alternative
pathway on decreasing respiration and plant growth (Sevilla et al., 2015). The effect of ROS
production on plant respiration and growth is not well understood, but provides one possible
explanation for the reduction in plant growth that occurred in the current study in resistance to
clubroot. Investigating the effect of P. brassicae on gene expression in a model plant host such as
Arabidopsis using a mutant construct with loss of function for an identified resistance gene could
improve understanding of metabolic cost of resistance to clubroot.
The data generally support the hypothesis that increasing spore concentration would
reduce plant growth and delay development in Brassica crops. However, the differences were not
quantifiable under low pathogen pressure or conditions of high variability in plant growth for
other reasons, such as soil compaction or saturation.
In the study of the effect of spore concentration and pH on clubroot incidence and
severity, susceptible canola inoculated with 1 x 103 spores mL-1 at pH 6.0-7.0 resulted in higher
clubroot incidence compared with 1 x 103 spores mL-1 at pH 5.5 and 7.5, at 22 days after
inoculation. In contrast, there was no effect of pH on clubroot incidence at 1 x 105 and 1 x 107
spores mL-1. This interaction was observed at 22 DAI but not at any other sample dates. These
results support previous reports by Colhoun (1953) that there is a strong influence of pH on
61 clubroot severity at low concentrations of resting spores, but not at high spore concentrations.
Previous studies have reported that 1000 spores g-1 soil is the infection threshold for susceptible
canola (Donald and Porter, 2009; Faggian and Strelkov, 2009). These results support the
hypothesis that pH 5.5 and >7.0 reduced clubroot incidence at low concentrations of resting
spores but not at concentrations ≥ 1 x 105 spores mL-1, although the differences only became
apparent at 22 days after inoculation. However, these results are based on only one repetition, and
therefore should be treated as preliminary until the study can be repeated. Study repetition should
include 10 plants per experimental unit instead of three plants to provide a more robust
assessment of clubroot incidence and disease severity. Plant root tissues collected at 8, 15 and 22
DAI have been stored for future examination of pathogen colonization of root tissue.
Quantification of pathogen DNA using quantitative polymerase chain reaction (qPCR) would
provide an assessment of pathogen colonization in host plant root tissue.
Increased understanding of the effect of resting spore concentration at various pH levels
will make it easier to predict clubroot severity and yield loss at specific field sites by providing a
more precise assessment of the risk of clubroot.
In summary, field studies indicated that there is a cost of resistance in canola and napa
cabbage. Reductions in plant height of resistant canola ranged from 12-14% indoors to 30% in
one field trial. Biomass was also reduced and maturity was delayed in some but not all studies.
Napa cabbage leaf length was reduced by 22-31% in field trials and biomass was reduced by 34%
in one field trial. Growers need to be aware that growing a resistant canola cultivar in a field with
a high concentration of resting spores results in a reduction in yield and delayed maturity, even if
no clubroot symptoms develop.
62
CHAPTER THREE
DECLINE IN RESTING SPORES AND EFFECT OF CROP ROTATION FOLLOWING A
SUSCEPTIBLE CROP
3.1 Introduction
Inoculum concentrations of soil-borne pathogens can increase rapidly in the presence of a
susceptible host (Gilligan et al., 1996; Hwang et al., 2013). High inoculum concentrations pose
an important threat to disease management strategies that rely heavily on genetic resistance. High
inoculum concentrations in soil can accelerate the breakdown of genetic resistance via selection
of more aggressive or virulent isolates of the pathogen (Kiyosawa, 1982; LeBoldus et al., 2012).
Furthermore, metabolic costs associated with disease resistance can lead to delayed development
and reduced plant growth and yield in canola (Tian et al., 2003; Hwang et al., 2011a).
Soil-borne pathogens such as Fusarium oxysporum f. sp. lactuca (Scott et al., 2012) and
Verticillium dahliae (Vos et al., 2012) can survive in soil in the absence of a host for 15 years or
more. The longevity of a soil-borne pathogen depends on many factors, including environmental
conditions, presence or absence of a host, and the morphological and chemical characteristics of
survival structures (McKeen and Wensley, 1961; Macfarlane, 1970; Coley-Smith et al., 1990).
Persistence in soil, and subsequent disease development following a long rotation, often lead
growers to conclude that crop rotation is not an effective method for reducing pathogen
populations.
The reduction in inoculum concentration of some soil-borne plant pathogens has been
studied in detail. Inoculum of V. dahliae in soil declined by 96% from the initial concentration
63 after 7 years of rotation with non-host crops. However, the inoculum concentration that remained
after 7 years was still three times the infection threshold (Ben-Yephet and Szmulewich, 1985).
Inoculum concentration of F. oxysporum f. sp. lactuca declined by 86% after a 12-month break
following a susceptible crop. In a resistant cultivar, however, F. oxysporum colonizes the root
cortex and produces inoculum, even though disease symptoms are not present (Scott et al., 2014).
Resistant plants contribute even more inoculum to the soil than susceptible plants, due to their
larger size.
For P. brassicae, a field survey was conducted on 190 field sites that were not uniform for
soil composition, structure, tillage practices, weed management, or cropping system during the
period of break from canola (Wallenhammar, 1996). Also, Brassica weeds were present in all of
the fields, some fields had adjacent areas highly infested with P. brassicae, and treatments were
not replicated to account for the variation in the initial concentration of resting spores. It took
about 17.3 years for the concentration of resting spores in a highly infested soil to decline below
the detection limit in Sweden. This study was conducted to a high standard using the technologies
available, but factors such as potential inflow of inoculum and increase on weeds complicate
interpretation of the data.
In Canada, resting spore concentration of P. brassicae declined by 98% from the initial
spore concentration of 3 x 106 spores g-1 soil following a 2-year break from susceptible canola, in
a recent study at the AAFC Research Farm at Normandin, Quebec (Peng et al., 2015). Thus, the
half-life of individual resting spores may be much shorter than the half-life of the infective
capacity of the soil (Wallenhammar, 1996; Hwang et al., 2013). Despite the decline in a large
number of resting spores, the resulting spore concentration (6 x 104 spores g-1 soil) after a 2-year
break from susceptible canola remained above the infection threshold for susceptible cultivars
64 (Peng et al., 2015), which is estimated to be 1000 spores g-1 soil (Donald and Porter, 2009;
Faggian and Strelkov, 2009).
Resting spores of P. brassicae are well-adapted to harsh environmental conditions
(Mafarlane, 1970). Spines covering the surface of the spore are enmeshed in fibres that are
approximately 5 nm in diameter (Buczacki et al., 1979). The protein that makes up the outermost
layers of the resting spore wall protects the inner chitin layer from natural chitinolytic processes
in the soil (Buczacki and Moxham, 1983). Environmental factors that affect germination include
pH, soil moisture, soil temperature, and the presence of calcium ions (Kageyama and Asano,
2009). Germination can take place with or without a host plant present (Ingram and Tommerup,
1972; Friberg, 2005), but the germination rate increases with the age and the level of decay of
spore-containing clubs (Macfarlane, 1970). The likelihood of germination also increases in the
presence of a compatible host (Macfarlane, 1970), such as susceptible canola volunteers. These
can persist in fields for many years and dramatically reduce the effectiveness of crop rotation
(Harker et al., 2014). Also, many of the common weeds in western Canada are susceptible to
clubroot, such as wild mustard (B. kaber L.), white mustard (B. hirta L.) shepherd’s purse
(Capsella bura-pastoris L.) and stinkweed (Thlaspi arvense L.).
From 1980 to 2000, more diverse crop rotations and reduced tillage were introduced in
western Canada as a strategy to reduce pest problems and decrease production risks. However,
from 2000 to the present, shorter rotations have become increasingly common, such as canola-on-
canola (no break from canola) and cereal-and-canola (1-year break from canola) (Kutcher et al.,
2013). Despite recommendations of canola 1 in 4 years (Rimmer et al., 2003) or canola 1 in 3
years (Cathcart et al., 2006), a shorter rotation is popular because of the high commodity price
and economic returns of canola relative to cereals and pulses (Strelkov et al., 2011). The
beneficial effect of crop rotation is associated with decreased allelopathy from plant residues
65 from previous canola crops, decreased presence of other canola pathogens, decreased weed
pressure, and improved soil structure and nutrition due to diverse rooting depths and nutrient
uptake of non-Brassica crops (Harker et al., 2014).
The effect of cropping rotation, especially of resistant canola cultivars, on resting spore
concentration in soil is not well understood. Resistant canola contributes fewer spores to the soil
than susceptible canola (Wallenhammar et al., 2000). Two studies of resistant cultivars of
Brassica vegetables observed a reduction in the concentration of resting spores of P. brassicae
relative to a non-host crop or fallow treatment (Yamagishi et al., 1986; Murakami et al., 2001).
In a more recent study, conducted in tubs placed outdoors, a resistant canola cultivar had either
no effect or produced a small increase in spore concentration in soil compared to an unplanted
control (Hwang et al., 2013). These studies used counts of spores extracted from soil, but recent
studies indicate that such estimates can be highly variable (Cranmer, 2015; F. Al-Daoud, personal
communication). Yield of resistant canola following 2- to 4-year break from susceptible canola
can be up to 25% higher than yield of canola grown in shorter rotations (Peng et al., 2015).
Accurate quantification of the concentration of P. brassicae resting spores in soil after
various lengths of break following susceptible and resistant canola can improve our
understanding of the long-term effect of crop rotation on canola production and have important
implications for clubroot management. Real-time quantitative polymerase chain reaction (qPCR)
assessments are effective for quantification of pathogen spore levels in the soil (Faggian and
Strelkov, 2009). However, physical or chemical substances that occur naturally in the soil, such
as humic acids, phenolic compounds, clay particles, or heavy metals, can inhibit amplification of
the target DNA during the qPCR process (Volossiouk et al. 1995; Matheson et al. 2010). Sample
dilution is a common approach to dealing with inhibition during qPCR amplification (Hoshino
and Inagaki, 2012). This is important because levels of inhibitors in soil cannot be assumed to be
66 consistent across treatments of crop rotation, which may have varying degrees of organic matter
breakdown. The use of an internal control allows for estimation of the level of inhibition that is
present in each sample, and so more accurate estimates of spore concentration (Deora et al.,
2015).
The first objective of this research was to examine the effect of crop rotation on the
decline of resting spores over time and the concentration of resting spores in soil. It was
hypothesized that the concentration of resting spores in soil would decline exponetially with
increasing length of break interval following a susceptible canola crop. The second objective was
to examine the interaction between crop rotation and inoculation with P. brassicae on growth of
clubroot-resistant canola. The main effect of crop rotation was expected to increase plant height
as length of break from canola increases. The main effect of inoculation with P. brassicae was
expected to decrease plant height compared with a non-inoculated control. It was hypothesized
that the interaction of crop rotation and inoculation with P. brassicae would affect the metabolic
cost of resistance to clubroot and subsequent reductions in plant height of resistant canola. It was
therefore expected that plant height would be shortest in the treatment of soil with no crop
rotation and inoculation with P. brassicae.
67
3.2 Materials and Methods
3.2.1 Decline in resting spores over time following susceptible canola
Soil samples were collected in the spring of 2014 from the Agriculture and Agri-Food Canada
(AAFC) research farm at Normandin, Québec (latitude 48°51’N, longitude 72°32’W). The soil is
a Labarre silt loam (Humic Gleysol) that consists of 8% sand, 70% silt and 22% clay and
approximately 4% organic matter (assessed separately). Soil composition and texture were
determined at the Agrifood Laboratories, Guelph, ON. The soil is naturally infested with
P. brassicae, identified as pathotype 2 (Strelkov et al., 2006) according to the Williams systems
of classifications (Williams, 1966). Small blocks (8 x 30 m) were selected based on cropping
history as part of a larger, long-term crop rotation study, where the rotation crops were canola,
barley, field pea and fallow. In fallow treatments weeds were controlled once a year (by
harrowing). However, fallow treatments were not weed-free throughout the entire season and
weeds may have included susceptible host species. Break intervals following susceptible canola
were 0, 1, 2, 3, 5, and 6 years (Table 3.1).
Resting spore concentration in soil was quantified after break intervals of 0, 1, 2, 3, 5, and
6 years following a susceptible canola crop. For each length of break, two plots were selected and
each plot was divided into two parts for subsampling, by collecting five soil cores (surface to 15-
cm depth) in a W pattern, with a core from each point in the W. The samples were dried at room
temperature and bulked. Each subsample was pulverized using a ceramic pestle and mortar
(CoorsTek Inc., Golden, CO), which were cleaned and autoclaved prior to each use to avoid
contamination. DNA was extracted using the PowerSoil DNA isolation kit (MO BIO
Laboratories Inc., Carlsbad, CA). Three biological replicates were conducted for each soil bulk,
with three technical replicates per biological replicate. For each biological replicate, 250 mg
68
Table 3.1 Cropping rotation following susceptible canola in selected plots sampled from a long-term rotation study at AAFC research farm in Normandin, Quebec.
Years break
from canola Block Cropping history
0 1
2
n/a
n/a
1 1
2
Fallow (F)
F
2 1
2
F-F
F-F
3 1
2
F-F-F
F-F-F
5 1
2
Pea-Barley-F-F-F
Barley-Pea-F-F-F
6 1
2
Barley-Barley-Pea-F-F-F
Barley-Barley-Pea-F-F-F
of soil was added directly into microbead tubes provided with the kit. DNA extraction and
purification were performed according to the manufacturer’s instructions. The spore
concentration in each sample was determined using a multi-plexed TaqMan qPCR assay, with a
competitive internal positive control (CIPC) to minimize differential inhibition among the
samples (Deora et al., 2015). Amplification of a known quantity of the CIPC is used to estimate
the level of PCR inhibition during the DNA amplification process (Wang et al., 2007). The CIPC
selected for these assessments was the DNA of Green Fluorescent Protein (GFP) from Aequorea
victoria, a jellyfish (Deora et al., 2014). Forward (DC1F) and reverse (DC1mR) primers
amplified a 90-bp fragment of the internal transcribed spacer (ITS1) region of P. brassicae. A
69 TaqMan probe (PB1) with a 5’ end reporter dye of FAM and 3’ quencher of NFQ-MGB was
used. The CIPC probe, GFP1, was derived from a plasmid (pDSK-GFPuv1) that contains a
variant of gene coding for GPF (GFPuv1) (Wang et al., 2007). Both primers and probe were
designed for conventional and quantitative PCR amplification of P. brassicae, using a partial
P. brassicae 18S ribosomal RNA (rRNA) gene sequence (Hwang et al., 2011b).
Assays were conducted in a 96-well Step-One Real-Time PCR system (Applied
Biosystems, CA), according to the manufacturer’s instructions. Each sample was homogenized
using a mechanical bead rupter (OMNI International, Inc., Kennesaw, GA) for 1 min. at high
speed. A 10x dilution with molecular grade water (Hyclone Laboratories Inc., Logan, UT) was
made for each genomic DNA (gDNA) sample. A standard series was produced using gDNA
derived from 1 x 108 P. brassicae resting spores mL-1 diluted in 10-fold increments down to 102
resting spores mL-1 (Deora et al., 2015). The standard series was used to generate a standard
curve for qPCR assays (as described by Deora et al., 2015, and previously conducted for
Entrococcus bacteria by Lievens and Thomma, 2005). Two technical replicates were included in
each concentration of the standard series. A negative control sample (Hypure™Molecular
Biology Grade Water, Hyclone Laboratories Inc.) with two technical replicates was also included
in each assay. An additional standard series without GFP was included as a negative control for
GFP in each assay. Each 20 µl reaction consisted of 10 µl Taqman® Universal PCR Master Mix
(P/N 4304437, Applied Biosystems), 1.8 µl of 250 nM of P. brassicae primer pairs, 0.5 µl of 250
nM of P. brassicae probe, 0.5 µl of 250 nM CIPC probe (GFP1), 1.4 µl of molecular grade water,
2 µl internal control and 2 µl of target DNA. Thermal cycling conditions were: 50°C for 2 min.,
95°C for 10 min., 40 cycles of 95°C for 15 sec., followed by 62°C for 1 min. Estimates of spore
concentrations were generated as the output for Cq values. The detection limit was Cq > 35 and
70 the lowest value on the standard curve was 103 spores mL-1 (Cq = 35.3). Therefore, a Cq value of
35.1 was selected as a cut-off point, and values lower than the cut-off were considered to be not
accurately quantifiable data.
Historical weather data for the Normandin Research Farm was accessed from
Environment Canada at http://climate.weather.gc.ca. Mean monthly air temperature was
calculated from historical weather data over a 10-year period, as well as total monthly rainfall
during the growing period of canola at Normandin, Quebec, from 2007–2014.
3.2.2 Effect of resistant canola on the concentration of resting spores in soil
In 2014, soil samples were collected from a follow-up study for the study described in
Section 3.2.1 at AAFC Normandin Research Farm. The clubroot-resistant canola cultivar 45H29
(Pioneer Hi-Bred Ltd, Chatham, ON) was seeded in 20 plots in 2014. Each plot is one
experimental unit. There were three treatments. One treatment was seeded with canola cv. 45H29
where susceptible canola had been cultivated the previous year, with eight field replicates.
Another treatment was seeded with canola cv. 45H29 where susceptible canola had been grown
in 2012, followed by a 1-year break (fallow), also with eight field replicates. Another treatment
was seeded canola cv. 45H29 where susceptible canola had been grown in 2011, followed by a 2-
year break (fallow), with four field replicates. Each plot was sampled as described above in early
spring of 2014 (before seeding), again in fall 2014 (after harvest), and in spring 2015
(approximately 8 months after harvest). Resting spore concentration was estimated using the
extraction and qPCR protocol described above.
71 3.3.3 Controlled environment study – crop rotation and spore concentration
Experiments examining the relationship between the presence of P. brassicae resting
spores, soil type and crop rotation were conducted in a controlled environment study using field
soil with no history of clubroot. The study was conducted as a 2×2×2 factorial arranged as a
randomized complete block design with four replicates. The factors were: crop rotation (0- vs. 2-
year break from canola), inoculation (P. brassicae spore suspension vs. water-only control), and
soil type, represented by two locations (Scott, SK and Elora, ON) (Table 3.2). A Dark Brown
Chernozemic mixed Elstow and shallow Elstow loam soil (Brandt and Zentner, 1995) was
collected in June of 2014 from two treatments in a rotation study at the Agriculture and Agri-
Food Canada Research Farm near Scott, SK. A Guelph series loam (Orthic Gray Brown Luvisol)
(Knezevic et al., 1994) was collected in June of 2014 from two adjacent sites at Elora Research
Station of the University of Guelph near Elora, ON. At each location, one sampling site had no
break from canola (planted to canola in 2013) and the other site had a 2-year break from canola
(planted to canola in 2011). During the 2-year break from canola, the site at Scott was cropped
with wheat the first year and field pea the second year. The site at Elora was cropped with
soybean followed by maize.
72 Table 3.2 Comparison of soil locations used in crop rotation and inoculum trials, under
controlled environment in Guelph, Ontario, 2014.
Location Latitude,
Longitude
Years break
from canola
pH Nutrient analysis (ppm) OM1
(%) P K Mg Ca
Elora,
ON
43°64' N,
80°40' W
0 7.2 37 135 346 3186 4.0
2 7.3 23 147 426 3038 3.9
Scott,
SK
52°36' N,
108°84' W
0 5.1 46 391 265 1145 3.5
2 6.5 31 338 344 2151 4.5 1 OM = Organic matter content (%)
The clubroot-resistant canola cultivar 45H29 was sown in tall plastic ‘conetainer’ pots
(Stuewe Sons Inc., Corvallis, OR) filled with soil. Two seeds were planted per pot and thinned to
one seedling per pot. There were 10 plants per experimental unit and four replicates. The
clubroot-susceptible canola line ACS-N39 (AAFC, Saskatoon Research Centre, Saskatoon, SK)
was included as a susceptible control for each treatment. The plants were maintained at 25°/20°C
day/night, with 16-hr photoperiod and 65% relative humidity, and fertilized with N-P-K solution
(20-20-20) at 2–3 day intervals. The plants were watered weekly with deionized water adjusted to
pH 6.0 using commercial white vinegar to maintain a pH level between 6.0 and 6.5. Each 10-day-
old seedling was inoculated with 5 mL of resting spore suspension of 1 x 106 resting spores mL-1
of P. brassicae pathotype 6. Inoculum was prepared following the protocol outlined by Sharma et
al. (2011). Briefly, mature clubs were washed and soaked in deionoized water; 10 g of clubbed
root was homogenized in 300 mL deionized water for 2 min. in an electronic grinder. The
mixture was filtered through eight layers of cheesecloth and resting spore concentration was
estimated using a haemocytometer. The spore suspension was diluted to 1 x 106 resting spores
mL-1.
73 Plant height, measured from the soil surface to the shoot apex, was assessed for each plant
in each experimental unit at 7-day intervals from 2−6 weeks after inoculation (WAI). At 10
weeks after planting, plants were harvested and weighed, and roots were assessed for clubroot
incidence (%) and severity using the standard 0-3 rating scale, where 0 = no clubbing, 1 < 1/3 of
root clubbed, 2 = 1/3–2/3 of roots clubbed and 3 > 2/3 of roots clubbed (Strelkov et al., 2006). A
disease severity index (DSI) was calculated according to Crete et al. (1963):
DSI = ∑ [(c [class no.)(no. of plants in each class)]
x100 (total no. plants per sample)(no. classes - 1)
3.3.4 Statistical analysis
All of the statistical analyses were performed with SAS software (version 9.3 SAS
Institute, Cary, NC) with a type I error set at P = 0.05. Data were tested for normality using the
Shapiro-Wilk test of residuals and tested for outliers using Lund’s test. Regression was tested
using PROC REG using data-points from two parts per block and two blocks per treatment.
The concentration of resting spores was not normally distributed, based on the Shapiro-
Wilk statistic (W = 0.431, P < 0.0001). Data were log transformed to reduce the heterogeneity of
the variance. Tests of normality on log-transformed data confirmed that distribution was normal
(W = 0.946, P = 0.22). Lund’s test of outliers identified only one extreme observation in the
treatment for 0-year break from canola. Variability in the concentration of resting spores
increased in treatments with higher concentration of resting spores in soil, in particular for the 0-
break from canola treatment. The data point was retained in the analysis, even though the
Studentized residual was 2.90, which is higher than the critical value for n=24, q=1 (Lund, 1975).
The concentration of resting spores in the 2014 resistant canola trial was also not
normally distributed, based on the Shapiro-Wilk statistic (W = 0.904, P = 0.008). Data were not
74 normally distributed when log-transformed (W = 0.943, P = 0.001). However, parametric
analyses were continued to remain consistent with analysis of all other data in the study.
Height measurements were collected for comparison at individual time-points and also to
examine overall growth over time. Growth over time, using plant height assessments over time as
a measurement for growth, was summarized using ‘area under the growth stairs’ (AUGS) as
previously described (Richards, 1959; Simko and Piepho, 2012). The area under the curve is
divided into a series of trapezoids using a simple calculation called the trapezoidal rule (Simko
and Piepho, 2012).
3.3 Results
3.3.1 Decline in resting spores over time following susceptible canola
Resting spores of P. brassicae were present in all plots and treatments sampled. The
concentration of resting spores following a susceptible canola cultivar declined over time in a
quadratic relationship, y = (1E + 07)e-0.759x, R2 = 0.65, as the length of break from the susceptible
canola crop increased (Fig. 3.1). Regressional analysis resulted in a linear relationship on log-
transformed data (P = 0.008). Resting spore concentration declined by 96.5% after a 1-yr break,
and 99.3% after a 2-yr break from canola, but then declined slowly compared to continuous
canola (1.3 x 108 spores g-1 soil).
75
Figure 3.1 Decline in resting spore concentrations of Plasmodiophora brassicae over time
following susceptible canola at Normandin, Quebec, assessed in 2014.
3.3.2 Effect of resistant canola on the concentration of resting spores in soil
Following continuous susceptible canola, one crop of resistant canola increased the
concentration of resting spores in soil by 12-fold, from 2 x 108 to 3 x 109 spores g-1 soil (Table
3.3). However, an unexpectedly high level of clubroot symptoms developed on canola cultivar
45H29, which has previously been resistant to clubroot at the Normandin site (D. Pageau,
personal communication). Following a 1-year break from susceptible canola, one crop of resistant
canola had no significant effect on the concentration of resting spores in soil. Following a 2-year
break from susceptible canola, one crop of resistant canola increased the concentration of resting
spores in soil by nearly 8-fold from 2 x 106 to 1 x 107 spores g-1 soil (Table 3.4).
Large numbers of resting spores were lost between the fall of 2014 and the spring of
2015. The concentration of resting spores was reduced by 97% (±0.3) from 3 x 109 spores g-1 soil
in the fall of 2014 to 8 x 107 in the spring of 2015, when one crop of resistant canola followed
y = 1E+07e-‐0.759x R² = 0.65, P = 0.008
1.0E+04
1.0E+05
1.0E+06
1.0E+07
1.0E+08
0 1 2 3 4 5 6
Log scale
Resting spores g-‐1 soil
Years break from canola
Decline in Resting Spores
76 continuous susceptible canola (Table 3.4). Similarly, resting spores were reduced by 99% (±0.2)
when resistant canola followed a 1-year break from susceptible canola and 90% (±1.0) when
resistant canola followed a 2-year break from susceptible canola (Table 3.4).
Prior to planting with canola cv. 45H29, the concentration of resting spores in soil
declined by 92% after a 1-year break and 99% after a 2-year break from canola, compared to no
break from susceptible canola (2 x 108 spores g-1 soil, Table 3.4). The concentration of resting
spores declined by >99% in treatments Fallow(F)-Resistant(R) and F-F-R, compared with
Susceptible(S)-R (3 x 109 spores g-1 soil) in soil samples collected in the fall of 2014. The
concentration of resting spores in soil declined by >98% in treatments F-R and F-F-R, compared
with S-R (8 x 107 spores g-1 soil) in soil samples collected in the spring of 2014.
After one crop of canola cv. 45H29, there was no difference between the F-R treatment
(1-yr break between susceptible and resistant canola) and the F-F-R treatment (2-yr break
between susceptible and resistant canola). Clubroot symptoms were observed in these fields,
indicating that spore concentration may have been influenced by susceptible weeds or off-types,
or a pathotype of P. brassicae that has overcome resistance of the host cultivar.
77 Table 3.3 Quantification of resting spore concentration using qPCR after crop rotation
treatments including fallow (F), resistant (R) and susceptible canola (S), sampled in spring and
fall of 2014 and spring of 2015 at Normandin, Quebec.
Treatment Inital1
(g-1 soil) SE2 Final3
(g-1 soil) SE
Spring 2014 S 5 x 106 2.0 x 106 2 x 108 1.8 x 108
F 1 x 107 5.7 x 106 2 x 107 3.6 x 106
F-F 8 x 105 1.8 x 105 2 x 106 4.3 x 105
Fall 2014 S-R 3 x 107 6.8 x 106 3 x 109 2.3 x 109
F-R 1 x 107 5.3 x 106 1 x 107 3.3 x 106
F-F-R 1 x 107 2.9 x 106 1 x 107 2.9 x 106
Spring 2015 S-R 2 x 107 1.2 x 107 8 x 107 5.6 x 107
F-R 5 x 104 3.7 x 104 2 x 105 1.2 x 105
F-F-R 5 x 104 2.3 x 104 1 x 106 7.8 x 105
1Initial= Initial spore concentration prior to adjustment of estimates for inhibition based on
amplification of the internal control. 2SE = Standard error. 3Final = Final spore concentrations (g-1 soil) following adjustment for the internal control
are indicated in bold.
3.3.3 Weather
At the site of the crop rotation study at Normandin, soil moisture and temperature were
generally conducive for clubroot development during the first 2 to 3 weeks after seeding (Table
A2.7). During the course of the portion of the long-term study examined in the current report
(2007-2014), weather conditions were generally near normal. Rainfall was lower than the 10-year
average by 12-36 mm from May to July 2007, higher by 17-57 mm from June to July 2008, lower
78 by 46 mm in July 2010, higher by 41 mm in May 2011, higher by 17-33 mm in May and June
2012, and higher by 16-39 mm in May and June 2013 (Table A2.7). Throughout the period of the
study, mean monthly air temperature varied up to 2°C cooler and 2°C warmer than the 10-year
average during the growing season (May to September, Fig. 3.2). Mean air temperature in May
was lower than 9°C in 2008 and 2009. Mean air temperature in May was 10°C in 2011 and 11°C
in 2010, 2012 and 2013.
Figure 3.2 Monthly rainfall (bars) and mean monthly air temperature (line) at Normandin,
Quebec, for May to September, 2007-2013.
3.3.4 Controlled environment – crop rotation and spore concentration
Clubroot incidence and severity in the susceptible control were higher in soil from Scott
than in soil from Elora (Table 3.5), likely associated with higher pH in the Elora soil.
0 2 4 6 8 10 12 14 16 18 20
0
50
100
150
200
250
M J J A S M J J A S M J J A S M J J A S M J J A S M J J A S M J J A S Monthly air temperature (m
ean °C)
Monthly rainfall (mm)
2007 2008 2009 2010 2011 2012 2013
79 Table 3.4 Clubroot incidence (CI) and severity (disease severity index, DSI) in canola breeding
line ACS-N39 (susceptible control) and the resistant cultivar 45H29, inoculated with P. brassicae
and grown under controlled conditions.
ACS-N39 45H29
Assessment CI DSI CI DSI
Scott soil, inoculated 90 a1 66 a 0 ns2 0 ns
Scott soil, not inoculated 0 c 0 c 0 0
Elora soil, inoculated 65 b 30 b 0 0
Elora soil, not inoculated 0 c 0 c 0 0 1 Means in a column followed by the same letter do not differ, based on Tukey’s multiple means
comparison test at P = 0.05. 2 ns = not significant.
Inoculation of the clubroot-resistant canola cultivar with resting spores of P. brassicae
reduced plant height by 7% (±3.3) at 4 WAI (P = 0.047), and 5 WAI (P = 0.038). Data from the
two repetitions of these dates were pooled because there were no significant repetition effects in
plant height (Tables A2.8-A2.9). There was a significant soil location effect at four out of five
time-points, as expected based on the broad differences in soil type, texture, pH and cropping
history.
Area Under the Growth Stairs (AUGS), calculated from plant height measured each week
from 2-6 WAI, indicated that overall growth of clubroot-resistant canola cultivars inoculated with
P. brassicae was reduced by 8% (±4.8) (P = 0.04, Table A2.11) across repetitions in soil from
both sites. In contrast, there was no effect of cropping rotation or interaction between inoculation
and crop rotation at either site (Fig. 3.3).
80
Figure 3.3 Effect of crop rotation and inoculation with Plasmodiophora brassicae on plant
height (area under growth stairs, AUGS) of clubroot-resistant canola, under controlled conditions
in field soil from Elora, ON, and Scott, SK in 2014.
The results for biomass (dry shoot weight) of resistant canola cultivars were more complex. The
biomass of resistant plants inoculated with P. brassicae was consistently lower than the non-
inoculated control (P = 0.004). For crop rotation, each site and repetition was analyzed separately
because there was a significant interaction between site, rotation and repetition (P = 0.001). In
Scott soil, a 2-year break from canola increased plant biomass by 41% compared with no break in
the first repetition (P = 0.002), but there was no effect in the second repetition (Table 3.6). The
trends were similar in repetition 2, but there was more variability in the treatment with 2-year
break from canola and therefore increased standard error. The opposite trend was observed in
Elora soil, where the 2-year break reduced biomass by 25% compared with no break (P = 0.003,
Table 3.6).
A A A A
0
30
60
90
120
150
180
Elora, ON
Plan
t hei
ght (
AU
GS)
2-yr Break- Inoculated 2-yr Break- Water Control No Break- Inoculated No Break- Water Control
AB A
B
AB
0
30
60
90
120
150
180
Scott, SK Pl
ant h
eigh
t (A
UG
S)
2-yr Break- Inoculated 2-yr Break- Water Control No Break- Inoculated No Break- Water Control
81 Table 3.5. Effect of crop rotation and inoculation with P. brassicae on biomass (dry shoot
weight, g) of clubroot-resistant canola, under controlled conditions in field soil from Elora, ON,
and Scott, SK, 2014.
Elora, ON Scott, SK
Repetition 1 Repetition 2 Repetition 1 Repetition 2
Spores mL-1
Biomass SE1 Biomass SE Biomass SE Biomass SE
No break
1x106 2.7 ab2 0.09 4.3 ns3 0.34 2.4 b 0.23 2.3 ns3 0.12
0 2.9 a 0.33 5.1 0.70 2.7 b 0.34 2.7 0.15
2-year break
1x106 2.0 b 0.24 3.0 0.50 3.3 ab 0.23 2.1 0.26
0 2.5 ab 0.26 3.7 0.64 4.0 a 0.27 2.8 0.70 1Means in a column followed by the same letter do not differ based on Tukey’s multiple means
comparison test at P = 0.05. 2ns = not significant. 3SE = standard error.
3.4 Discussion
In the current study, the concentration of resting spores in soil following a susceptible
canola cultivar declined over the first 2 years following a susceptible canola crop. These results
support a recent study that observed a decline in resting spores at the AAFC Research Farm at
Normandin, QC, conducted at the same site as the current study but in two different years (Peng
et al., 2015). Similarly, the concentration of resting spores in soil declined in mini-plots
consisting of plastic tubs containing field soil placed outdoors near Edmonton, AB, where both
continuous resistant canola and continuous fallow reduced the concentration in resting spores in
soil relative to continuous susceptible canola (Hwang et al., 2013).
The current study indicated that the concentration of resting spores in soil declined by
97% after a 1-year break and 99% after a 2-year break from susceptible canola, but then declined
82 very slowly (<1%) over the next four years. Following a 6-year break from canola, the
concentration of resting spores was 4.1 x 105 spores g-1 soil. This is more than 400-fold greater
than the estimated threshold for uniform infection (1000 spores g-1 soil) in a susceptible cultivar
(Donald and Porter, 2009). Therefore, a viable management strategy for a location with a high
concentration of P. brassicae resting spores should include crop rotation in combination with a
clubroot-resistant cultivar. A management strategy that combines a crop rotation with ≥ 2-year
break from canola and use of a clubroot-resistant cultivar can reduce the concentration of resting
spores and fitness cost of resistance to clubroot. This would also contribute to a reduction in
selection pressure within the pathogen population, which could delay the erosion of resistance
(Kiyosawa, 1982; LeBoldus et al., 2012).
Estimates of the half-life of the infective capacity of resting spores have ranged from 3.6
years (Wallenhammar, 1996) to 4.4 years (Hwang et al., 2013). However, a decrease in ≥ 98% of
resting spores in the first 2 years following susceptible canola was observed in the current study
and in a recent study at the same site in two different years (Peng et al., 2015). It is possible that
within one population of resting spores, there may be differences in the half-life of individual
resting spores. These differences may be related to decreased microbial degradation of resting
spores that have increased protein structure in the outer spore wall (Moxham et al., 1983). Thus,
although ≥ 98% of resting spores die or disappear within the first 2 years, the half-life of the
remaining ≤ 2% of spores may be closer to 5 years or longer. Similarly, there could be large
differences in the pattern of decline in resting spores at different sites. For example, in a site
where clubroot has been present for decades and many susceptible hosts have been grown, it is
possible that the spore population in the soil is more adapted for protection and would have a
longer half-life of infective capacity than in an inoculated soil or a soil with a relatively new
83 infestation. However, it is also possible that there could be an increase over time in the
concentration of microbes that degrade P. brassicae resting spores, which would lead to a shorter
half-life of resting spores at sites where clubroot has been established for a longer period of time.
Clubbing symptoms were observed in a large number of canola plants in the field at
Normandin, Quebec, seeded with cultivar 45H29. This cultivar was selected because it had
previously demonstrated resistance against the clubroot pathotype at Normandin (D. Pageau,
personal communication). The trend of a decline in the concentration of resting spores as the
length of break interval increased from 0- to 2-year break from susceptible canola was also
observed in soil samples collected before and after one crop of 45H29 canola. However, there
was an 8- to 12-fold increase in the concentration of resting spores in the soil samples taken in
the fall of 2014 after harvest of 45H29, and the concentration of resting spores in soil increased
following 45H29 in two of three treatments in the study. In samples collected in the spring of
2015, resting spore concentration decreased by ≥ 90% overwinter. These results contradict
reports that resistant canola has either no effect or a small increase in spore concentration in soil
compared to an unplanted control (Wallenhammar et al., 2000; Hwang et al., 2013). In a field
trial near Edmonton, AB, a decline in resting spores was observed despite a 14% clubroot
incidence in the field planted with resistant canola (Hwang et al., 2013). Similarly, the
concentration of resting spores in soil was reduced by 36-45% following clubroot-resistant leafy
daikon cultivars (Raphanus sativus var. longipinnatus) in soil artificially inoculated with 106
spores mL-1 (Murakami et al., 2001). Clubroot severity on subsequent napa cabbage plants was
reduced compared to a fallow control. In another study, the concentration of resting spores was
reduced by 99.9% from the starting concentration of 5 x 106 spores mL-1 soil to 5 x 103 spores
mL-1 soil following four years of cultivation of kale (B. oleracea ssp. acephala) and turnip
84 (B. rapa ssp. rapa) (Yamagishi et al.,1986). In each of these previous trials, soil was artificially
inoculated with resting spores. As mentioned previously, it is possible that the viability of spores
applied by inoculation may decline at a different rate than spores under natural conditions. This
could explain why a much greater number of resting spores was contributed to the soil by a
resistant cultivar in an naturally infested field, compared with artificially inoculated soil.
However, the different trend observed in the current study may also be explained by the very high
level of clubroot symptoms on the resistant canola, which indicates that there was either a very
high percentage of susceptible canola volunteers or susceptible off-types, or that genetic
resistance was overcome by the pathogen. This latter possibility is being explored in another
study (B. Gossen, personal communication). Therefore, these results may not be indicative of
crop rotation with a resistant canola cultivar in a field where genetic resistance has not been
overcome. However, differences in experimental methods may have also played a role.
Significant reductions in resting spore concentration following a resistant cultivar were observed
in two studies in Japan where there is a different pathotype of P. brassicae than in Canada. In
addition, clubroot-resistant cultivars of B. oleracea and B. rapa may have a different mechanism
of resistance than clubroot-resistant B. napus. Finally, several of these studies estimated the
concentration of resting spores in soil using extraction from soil and direct counts. Estimates
using counts can be highly variable and often not correlated with results from qPCR (Cranmer,
2015).
In the current study, there was substantial variation in the amount of P. brassicae DNA
detected. Similar high variability has been reported in previous studies (Cranmer, 2015;
Wallenhammar et al., 2012), despite the steps taken to minimize variability. For example, each
subsample consisted of five cores rather than three, and cores were homogenized prior to DNA
extraction. Variability in resting spore counts increased at higher concentrations of resting spores
85 in soil, and was generally not normally distributed about the mean. Inhibition levels in the soil
samples from Normandin were low compared with other studies (Wallenhammar et al., 2012;
Cranmer, 2015), likely because there were comparatively low levels of organic matter in the soil
(~3.7%). One drawback to qPCR assessment of resting spore numbers is that viable spores
cannot be distinguished from non-viable spores (Faggian and Strelkov, 2009). As a result, qPCR
may overestimate the concentration of viable resting spores in soil relative to a bioassay.
Previous studies have reported that rainfall during the first 2 to 3 weeks after seeding is
correlated with clubroot incidence and severity (Thuma et al., 1983; Gludovacz, 2013). Canola is
normally seeded in late May or early June at Normandin, Quebec. In the current study, the most
recent susceptible canola crop was planted in the same year for each block within that treatment
of time break from canola. Therefore, it is conceivable that a year with higher than average
rainfall during the 2 weeks after seeding could lead to a higher initial spore concentration relative
to another year with lower rainfall. For example, monthly rainfall was lower than average by
12-36 mm from May to July 2007 and higher than average by 17-57 mm from June to July 2008.
However, the concentration of resting spores following a 6-year break from susceptible canola
(planted in 2007) was slightly higher than following a 5-year break from susceptible canola
(planted in 2008). In 2011, 2012 and 2013, rainfall was moderately higher than average in May
and June, which indicates that there was no substantial effect of rainfall on the inoculum level
prior to the break from susceptible canola. This supports the conclusions from recent studies
(Kasinathan 2012; Gossen et al., 2013; Cramner, 2015) that beyond certain minimal threshold
values of temperature and soil moisture, clubroot develops at high levels irrespective of weather
in temperate regions when inoculum concentration is high.
Quantification of resting spores of P. brassicae in soil can be valuable in evaluation of
clubroot management practices. Current crop rotation recommendations are canola one in four
86 years (3-year break from canola) on the Canadian prairies (Kutcher et al., 2013). A crop rotation
of 5 to 7 years (4- to 6-year break from canola) is recommended in Ontario for Brassica crops
including canola (OMAFRA, 2000).
The first objective of this research was assessed by quantifying the DNA of P. brassicae
in soil samples with break intervals of 0- to 6-years following susceptible canola, and in a
companion study with treatments of resistant canola following various break intervals of 0- to 2-
years following susceptible canola. The results provided strong support for the hypothesis that the
concentration of resting spores in soil declined exponentially with length of break interval
following a susceptible canola crop. The greatest reduction in concentration of resting spores was
observed after 1-year break from canola, with a further reduction after the second year of break.
One limitation of this study is that the treatments (different rotations) in Normandin are
not directly comparable due to different weather conditions in different years of the rotation. For
example, fallow for one treatment may be in a wet year, and fallow for another may be in a dry
year. The study design included three biological and three technical replications of soil samples
for qPCR amplification. However, replication of crop rotation treatments in the field was limited
to only two replicates (each with two sub-samples) and the trial in it entirety was not repeated.
The second objective, to examine the interaction between crop rotation and inoculation
with P. brassicae on growth of clubroot-resistant canola, was assessed in a controlled
environment study. Resistant canola grown in soil from Scott, SK, was shorter in height,
indicating a greater cost of resistance than resistant canola grown in soil from Elora, ON. This
supports observations from Chapter 2 that the cost of resistance to P. brassicae causes greater
reductions in plant height when environmental conditions are more conducive to pathogen
development, including high soil moisture and more acidic soils. Susceptible canola grown in soil
from Scott, SK, had higher clubroot incidence and severity than susceptible canola grown in soil
87 from Elora, ON. This is likely related to the higher acidity of the soil from Scott, which makes
the site more conducive for clubroot development (Donald and Porter, 2004, Gossen et al., 2013;
Webster and Dixon, 1991a, 1991b).
No interaction between crop rotation and inoculation was observed. In Scott soil, plant
biomass increased by 42% in canola grown in soil following a 2-year break from canola, in one
repetition and a similar trend was observed in the second repetition although the difference was
not significant. These results are consistent with a previous study that investigated the effect of
crop rotation on canola in the absence of clubroot, where canola yield increased by 22% in a 1 in
6 year rotation (Harker et al., 2014). However, the decrease in plant biomass in canola grown in
Elora soil following a 2-year break from canola contradicts previous research that plant growth
increases with increasing diversity of crop rotation (Harker et al., 2014). This could be explained
by other differences in management of the Elora soil in addition to rotational diversity that were
not known, such as fertilizer or treatment with fungicide or insecticide that may have affected
microbial populations, or presence of other pathogens that were not consistent across rotational
treatments. Unlike in the location in Scott, SK, the sites at Elora Research Station were not
previously part of a replicated long-term crop rotation trial.
In the current study, field soil was used in conetainers to examine the effect of actual
cropping rotation that took place in a field rather than a simulated cropping history, for example
cycles of crops planted in tubs. Conducting crop rotation trials in controlled environments is not
recommended for future research. Plant growth was highly variable, likely due to disturbance of
the soil structure, disruption of microbial communities during drying and transport of soil, and
compaction from watering. This may explain why the results of the current study were
inconsistent and did not replicate previous results in field trials. Similar to the possible
88 differences between Elora soils, the differences between the soils from Elora, ON and Scott, SK
may be related to other soil micro-organisms that are pathogenic or beneficial to plant growth,
other environmental conditions such as pH, or agronomic practices such as crop type planted
during the break interval from canola (Table 3.2). One opportunity to improve crop rotation trials
for future research could be to investigate the effect of continuous resistant canola crops on plant
growth and yield, relative to resistant canola in rotation with fallow or non-host crops.
It was hypothesized that an additive effect of no crop rotation and P. brassicae would
reduce plant growth and development more in inoculated plants grown in soil with no break from
canola, compared with inoculated plants grown in soil with a 2-year break from canola. There
were several reasons to expect an additive effect, including allelopathy, presence of other root
pathogens, decreased soil nutrition, changes in soil structure, or even plant residues from
previous canola crops increasing the germination of resting spores. Some or all of these factors
were expected to contribute to reduced plant growth in the presence of P. brassicae in soils
previously cropped with continuous canola, compared with soils that had a 2-year break from
canola. However, this was not demonstrated in the current study. The effect of these factors was
likely limited because of the disruption of the soil, as discussed previously. As a result, there was
no main effect of crop rotation observed, and no interaction observed between crop rotation and
inoculation.
Examination of the effect of crop rotation on plant growth requires long-term planning
and implementation of a controlled experiment over many years in field plots uniform for pH,
soil structure, soil nutrient levels, fertilization and agronomic practices. In addition to the
challenges associated with a crop rotation trial, assessing the interaction between resting spore
concentration and crop rotation requires artificial inoculation of treatments to ensure that there is
a negative control with no resting spores. In the current study, an approach involving examination
89 of soil from field locations with no history of clubroot in a controlled environment facility was
selected instead of a field trial to avoid contamination of a field with a pathogen that did not yet
occur there.
An improvement to the method for the decline in resting spores study would be to
increase the soil sampling depth to 0-30 cm below the soil surface to include a greater proportion
of the total number of resting spores in the soil. Depending on soil type and tillage practices, the
concentration of resting spores in soil decreases with increasing depth (Cranmer, 2015). In a silt
loam soil at Bassano, AB, 54-65% of spores were observed in the top 0-16 cm at one site
following a 6-year break from susceptible canola and a 3-year break from resistant canola
(Cranmer, 2015). Low levels of resting spores were observed at 0-8 cm, and higher levels were
observed at 8-16 cm, 16-23 cm and 23-30 cm. This may indicate that few or no new resting
spores had been contributed to the soil near the surface in the past three or more years. In the
current study of a silt loam soil at Normandin, QC, samples were taken from 0-15 cm depth of
soil near the cortical zone of young roots (Gan et al., 2009), where the majority of clubroot
infection occurs. It is possible that an increased break interval from canola may provide an
opportunity for downward movement of resting spores rather than their death and disappearance.
The results reported here support the results of previous trials on crop rotation and
clubroot (Hwang et al., 2013; Peng et al., 2015). Research by Peng et al. (2015) assessed resting
spore concentrations in soil samples from the same site at AAFC Normandin Research Farm that
was assessed in the current study, but collected in two different years (2012, 2013). A break
interval ≥ 2-year following susceptible canola reduces the concentration of resting spores >90%
relative to 0- and 1-year break from canola (Peng et al., 2015). However, resting spore
concentration is not significantly different in soil samples following a 0- and 1-year break from
90 susceptible canola. In contrast, in the current study, a 97% reduction in the concentration of
resting spores was observed following 1-year break from susceptible canola relative to no break
from susceptible canola. Both studies quantified the concentration of resting spores in soil
samples using a qPCR protocol. The main difference in protocol was that an internal control was
included in the current study to adjust the final estimate of spore concentration based on
inhibition of pathogen DNA amplification. Using a protocol with an adjustment for internal
control resulted in a greater reduction in resting spore concentration from 0- to 1-year break
(97%) relative to the reduction observed from 0- to 1-year break without adjustment for internal
control (75%). It is likely that this was observed because inhibition was much higher in soil
samples collected following no break from canola, probably due to increased humic acids and
other organic compounds associated with plant residues. However, the difference may also be
related to limitations of the current study such as replication and repetition discussed above.
In conclusion, the results of this study indicate that when the initial concentration of
resting spores is high, increasing the length of break from canola up to 2 years can decrease the
fitness cost associated with clubroot resistance and increase the opportunity for high yield in a
resistant cultivar. In any case, where clubroot symptoms have been identified, a management
strategy should be implemented that includes a rotation interval ≥ 2 years break from canola
followed by planting of a clubroot-resistant cultivar.
91 CHAPTER FOUR
GENERAL DISCUSSION
Integrated clubroot management is necessary for the long-term viability of canola
production in Alberta and across the Canadian prairies. High concentrations of P. brassicae
resting spores in soil can reduce vegetative growth and seed production, delay the maturity of
canola and result in crop failure in susceptible canola. The primary strategy for clubroot
management in Canadian canola production is planting a clubroot-resistant cultivar, because
there are currently no viable cultural, biological and chemical strategies available to growers.
However, recent studies indicate that exposure to resting spores can result in failure to achieve
the yield potential of resistant cultivars (Hwang et al., 2013; Peng et al., 2015).
The current study investigated the effect of resting spore concentration and crop rotation on
growth of clubroot-resistant crops. The influence of spore concentration on growth of clubroot-
resistant cultivars was assessed in canola, napa cabbage and cabbage. Also, the effect of crop
rotation on resting spore concentration over time was examined. Another study investigated the
interaction of spore concentration and crop rotation on growth of clubroot-resistant canola under
controlled conditions. Finally, the interaction of spore concentration and pH on clubroot
incidence and development in susceptible canola was examined under controlled conditions.
This is the first study to specifically examine the cost of resistance to P. brassicae in
resistant cultivars of canola, napa cabbage and cabbage under a range of resting spore
concentrations. Resistant canola was 30% shorter, biomass was 43% lower and maturity was
delayed in the field site with higher spore concentration relative to a similar site with lower spore
concentration, in one of two years. The decrease in growth is consistent with previous studies
(Hwang et al., 2011a; Deora et al., 2012b), which indicated that there was a metabolic cost of
92 resistance to clubroot in canola under controlled conditions. In resistant napa cabbage, the cost of
resistance, indicated by leaf length, was consistent over both years for. Leaf length was reduced
by 31% in 2014 and 22% in 2015 at a site with high spore concentration. There were no
significant differences for cabbage.
A controlled environment study demonstrated that plant height of clubroot resistant canola
cv. 45H29 declined in a quadratic relationship with increasing concentration of resting spores.
Plant height was reduced by 12-14% compared to the non-inoculated control at 1 x 107 and
1 x 108 spores g-1 soil. Biomass of canola was also lower and maturity was delayed with
increasing concentration of resting spores in one of two repetitions of the trial.
This is the first study to examine the interaction of a range of pH levels with a range of
spore concentrations on clubroot incidence and severity in susceptible canola. It is known that
clubroot incidence and severity in susceptible cultivars varies depending on environmental
factors, such as temperature, pH and soil moisture (Cohoun, 1953; Sharma et al., 2011a, 2011b;
Gossen et al., 2013). Although it is known that the infection threshold for susceptible cultivars is
approximately 1000 spores g-1 soil (Donald and Porter, 2009; Faggian and Strelkov, 2009), the
interaction between spore concentration and environment is not well understood. The current
study indicated that inoculation with 1 x 103 spores ml-1 resulted in abundant clubbing symptoms
at pH 6.0-7.0, but fewer symptoms develop at lower or higher pH (5.5 or 7.5). These results
support previous reports that high pH suppresses clubroot severity at low concentrations of
resting spores, but is overwhelmed at high spore concentrations (Colhoun, 1953). This study
should be repeated. Future research could include extraction of DNA to investigate pathogen
colonization of plant root tissue, as well as extraction of RNA to investigate gene expression at
various time-points across a range of pH level and spore concentration. Increased understanding
of the interaction of spore concentration and pH could lead to more accurate predictions of
93 clubroot severity.
The current study demonstrated that large numbers of resting spores are lost from the
surface soil layers, where they are most effective at causing disease, in the 2 years following
susceptible canola. The concentration of resting spores in soil declined by 97% after a 1-year
break and 99% after a 2-year break from susceptible canola, but then declined very slowly (<1%)
over the next four years. These reults are consistent with previous reports (Hwang et al., 2013;
Peng et al., 2015). Although the concentration of resting spores after a 6-year break was still
much higher than the infection threshold for susceptible cultivars, there are benefits to reducing
spore concentration before seeding a resistant cultivar. Specifically, a crop rotation with a
≥ 2-year break from canola can reduce the concentration of resting spores and thus the fitness
cost of resistance to clubroot that would reduce the yield of the resistant canola crop. This may
also prolong the effectiveness of resistant cultivars by alleviating selection pressure with the
population of resting spores (Kiyosawa et al., 1982).
In the second year of this study, one crop of canola cv. 45H29 (resistant) was seeded in
field plots with break intervals of 0- to 2-years from susceptible canola. A 12-fold increase in
spore concentration when susceptible canola was followed by 45H29 is different from previous
reports of small changes or no difference following resistant canola (Wallenhammar et al., 2000;
Hwang et al., 2011). In the current study, a sampling date 8 months after harvest showed that
≥ 90% of resting spores in soil were lost. They may have disintegrated or moved downward more
than 15 cm from the soil surface over winter.
The presence of susceptible canola volunteers, weeds, or even a resistant cultivar can
increase resting spore germination (Macfarlane, 1970). For this reason, there has been speculation
that planting a resistant host could lead to germination of a large number of resting spores that do
94 not complete the lifecycle and thus reduce the overall concentration of resting spores in soil
(Murakami et al., 2001). In contrast, susceptible weeds can persist in a field over many years, and
dramatically reduce the effectiveness of crop rotation (Harker et al., 2014). In the current study,
the influence of growing one crop of resistant canola on the concentration of resting spores could
not be properly assessed because a very high level of clubroot symptoms developed on the
previously resistant canola cultivar 45H29 (D. Pageau, personal communication). This indicates
either a very high proportion of susceptible weeds / off-types, or that resistance in the canola
cultivar had been eroded.
A separate study is investigating the possibility that the pathogen has overcome clubroot
resistance at this site (B. Gossen et al., personal communication). Recently, cleaved amplified
polymorphic sequence (CAPS) markers have been developed to differentiate P. brassicae
populations and single-spore isolates for at least some of the new genotypes of the virulent
phenotype in Alberta, known as 5X (Cao et al., 2015). These markers could be used to analyze
the genetic similarity between the new pathotypes in Alberta with those from Normandin. Future
studies should investigate the influence of a resistant crop on spore concentration in soil in a field
trial with lower risk of the pathogen overcoming clubroot resistance during that growing season.
For example, this could include a field with slightly lower initial spore concentration, or a field
with no previous history of resistant canola.
There are several ways that the experiments described in this thesis could be improved.
First, clubroot severity was low to moderate in the susceptible control in studies of canola, napa
cabbage and cabbage under controlled conditions. There appeared to be problems with the
inoculum used in some of the controlled environment studies. The concentration of viable resting
spores may have been much lower than the estimation of 1 x 106 spores mL-1. Spore
concentration was estimated using haemocytometer spore counts, which cannot differentiate
95 between viable and non-viable resting spores. Also, the protocol used for extraction and
estimation of spores from soil, which was adapted by Sharma et al. (2012) from Dhingra and
Sinclair (1985), may overestimate the total number of resting spores because empty resting spore
shells, or possibly other debris, can be mistaken for spores of P. brassicae. A protocol is
currently being developed with improved filtration of the resting spore suspension prior to the
haemocytometer spore count (F. Al-Daoud, personal communication). Also, research is currently
underway to develop and test a protocol for quantification of viable P. brassicae resting spores
using propidium monoazide (PMA)-PCR (F. Al-Daoud, personal communication). PMA is a
photoreactive dye that is cell membrane-impermeable but can bind with DNA. It can modify non-
viable cells so that they are not amplified by PCR, while leaving the DNA of viable cells intact.
Other recent developments in quantification of resting spores in soil, including qPCR, droplet
digital PCR (ddPCR) and the loop mediated isothermal amplification (LAMP) assay, could
increase access to more accurate, more affordable and faster assessment of spore concentration,
relative to plant bioassays. Additional research should compare the accuracy of these methods of
quantification of resting spores in soil. In combination with improved site-specific spore
quantification, improved understanding of the relationship between spore concentration and seed
yield of canola under field conditions could aid growers and agronomists in selection of cultivars
and cropping intervals.
There are many challenges to investigating the cost of resistance in field trials. Finding a
suitable location to conduct the trial can be challenging. Field plots should have uniform soil
characteristics and nutrient levels, and differ only in spore concentration. However, introducing
pathogen inoculum to a field that is not currently infested would be detrimental to the goal of
reducing clubroot levels in soil. There is not currently a reliable method of eradicating resting
spores in soil. One potential option for future research on the metabolic cost of resistance would
96 be in the clubroot research nursery near Edmonton, AB. In the current study, the field trial was
located at Muck Crops Research Station in Holland Marsh, ON, where differences in spore
concentrations were already present within a small area of the field. The site with highest
concentration of resting spores is near the entrance to the field, where clubroot trials have taken
place in succession over many years, and also increased deposition of contaminated soil from
equipment entering the field may have occurred (Cao et al., 2009). Although research sites
nearby do not contain clubroot, equipment entering and leaving that site may have increased
clubroot due to compaction of soil from higher equipment traffic (Gossen et al., 2016).
In summary, the results of this study indicate that when the initial spore concentration in
soil is high, increasing the length of break from canola to 2 years can decrease the concentration
of resting spores, which in turn reduces the cost of resistance and maximizes the yield potential
of a resistant cultivar. A negative relationship was found between plant height and concentration
of resting spores in one of two field trials and under controlled conditions. Some of the variability
among trials could be due to reduced viability of some inoculum, variability among plants within
a study and environmental influences such as soil type, soil moisture or temperature. For canola
growers, a ≥ 2-year break from canola, in combination with a clubroot-resistant cultivar, is
recommended wherever clubroot occurs.
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APPENDIX 1: SUPPLEMENTARY TABLES FOR CHAPTER TWO
Table A1.1. Controlled environment study – canola: CI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 0 - - Repetition (R) 1 0 - - Error 3 0 Table A1.2. Controlled environment study – canola: DSI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 141 1.80 0.32 Repetition (R) 1 1147 14.64 0.03 Error 3 78 Table A1.3. Controlled environment study – canola: plant height at 6 WAI Source df Mean Square F value Pr>F Block (B) 3 16 2.13 0.12 Inoculation (I) 5 24 3.21 0.02 Repetition (R) 1 29 3.80 0.06 I x R 5 12 1.60 0.19 I linear (1) 40 5.22 0.03 I quadratic (1) 69 9.07 0.005 Error 33 8 Table A1.4. Controlled environment study – canola: Biomass (dry shoot weight) Source df Mean Square F value Pr>F Block (B) 3 2 2.16 0.11 Inoculation (I) 5 2 2.07 0.09 Repetition (R) 1 120 105.42 <0.0001 I x R 5 0 0.41 0.84 I linear (1) 4 3.68 0.06 I quadratic (1) 1 0.62 0.43 Error 33 1 Table A1.5. Controlled environment study – canola, repetition 1: Biomass (dry shoot weight) Source df Mean Square F value Pr>F Block (B) 3 6 10.01 0.0007 Inoculation (I) 5 1 2.41 0.09 I linear (1) 3 5.60 0.03 I quadratic (1) 0 0.05 0.82 Error 15 1
117 Table A1.6. Controlled environment study – canola, repetition 2: Biomass (dry shoot weight) Source df Mean Square F value Pr>F Block (B) 3 3 3.84 0.03 Inoculation (I) 5 1 2.03 0.13 I linear (1) 1 1.67 0.22 I quadratic (1) 1 1.49 0.24 Error 15 1 Table A1.7. Controlled environment study – canola, repetition 1: Maturation at 8 WAI Source df Mean Square F value Pr>F Inoculation (I) 5 968 6.95 0.0009 I linear (1) 3177 22.80 0.0002 I quadratic (1) 1314 9.43 0.007 I cubic (1) 12 0.08 0.78 Error 18 139 Table A1.8. Large pot studies – outdoor: CI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 60 1.69 0.27 Repetition (R) 2 7086 200.72 <0.0001 Error 6 35 Table A1.9. Large pot studies – outdoor: DSI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 145 1.18 0.39 Repetition (R) 2 5378 43.77 0.0003 Error 6 123 Table A1.10. Large pot studies – outdoor: plant height 6 WAI Source df Mean Square F value Pr>F Block 3 97 3.48 0.03 Repetition (R) 1 5146 185.71 <0.0001 Inoculation (I) 5 331 1.19 0.33 I linear (1) 27 0.96 0.34 I quadratic (1) 60 2.15 0.15 Error 33 28
Table A1.11. Large pot studies – outdoor, 2015 repetition 1: plant height 6 WAI Source df Mean Square F value Pr>F Block 3 18 1.30 0.31 Inoculation (I) 5 10 0.74 0.60 I linear (1) 32 2.36 0.15 I quadratic (1) 0 0.00 0.95 Error 15 14
118 Table A1.12. Large pot studies – outdoor, 2015 repetition 2: plant height 6 WAI Source df Mean Square F value Pr>F Block 3 150 4.52 0.02 Inoculation (I) 5 49 1.49 0.25 I linear (1) 3 0.08 0.78 I quadratic (1) 114 3.43 0.08 Error 15 33
Table A1.13. Large pot studies – outdoor, 2015: plant height 7 WAI Source df Mean Square F value Pr>F Block 3 40 2.14 0.11 Repetition 1 3977 214.39 <0.0001 Inoculation (I) 5 38 2.03 0.10 I linear (1) 0 0.03 0.88 I quadratic (1) 13 6.07 0.02 Error 33 19 Table A1.14. Large pot studies – outdoor, 2015 repetition 1: plant height 7 WAI Source df Mean Square F value Pr>F Block 3 6 0.43 0.73 Inoculation (I) 5 6 0.38 0.85 I linear (1) 8 0.55 0.47 I quadratic (1) 1 0.05 0.83 Error 15 14
119 Table A1.15. Large pot studies – outdoor, 2015 repetition 2: plant height 7 WAI Source df Mean Square F value Pr>F Block 3 47 2.00 0.16 Inoculation (I) 5 58 2.45 0.08 I linear (1) 3 0.14 0.71 I quadratic (1) 202 8.50 0.01 Error 15 24 Table A1.16. Controlled environment study – canola, napa cabbage, cabbage: CI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 168 1.18 0.37 Repetition (R) 1 2030 14.25 0.004 Inoculation (I) 1 6392 44.87 <0.0001 R x I 1 2030 14.25 0.004 Error 9 142 Table A1.17. Controlled environment study – canola, napa cabbage, cabbage: DSI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 33 1.94 0.19 Repetition (R) 1 693 40.74 0.0001 Inoculation (I) 1 1442 84.77 <0.0001 R x I 1 693 40.74 0.0001 Error 9 17 Table A1.18. Controlled environment study – canola, napa cabbage, cabbage: CI in susceptible napa cabbage Source df Mean Square F value Pr>F Block (B) 3 196 0.95 0.46 Repetition (R) 1 79 0.39 0.55 Inoculation (I) 1 4747 23.08 0.00 R x I 1 79 0.39 0.55 Error 9 206 Table A1.19. Controlled environment study – canola, napa cabbage, cabbage: DSI in susceptible napa cabbage Source df Mean Square F value Pr>F Block (B) 3 46 0.67 0.59 Repetition (R) 1 1 0.01 0.91 Inoculation (I) 1 1652 24.13 0.0008 R x I 1 1 0.01 0.91 Error 9 68
120 Table A1.20. Controlled environment study – canola, napa cabbage, cabbage: CI in susceptible cabbage Source df Mean Square F value Pr>F Block (B) 3 9 0.07 0.97 Repetition (R) 1 768 5.67 0.05 Inoculation (I) 1 4400 32.46 0.0007 R x I 1 751 5.54 0.05 Error 7 136 Table A1.21. Controlled environment study – canola, napa cabbage, cabbage: DSI in susceptible cabbage Source df Mean Square F value Pr>F Block (B) 3 6 0.12 0.94 Repetition (R) 1 95 2.11 0.19 Inoculation (I) 1 729 16.23 0.005 R x I 1 88 1.97 0.20 Error 7 45 Table A1.22. Controlled environment study – canola, napa cabbage, cabbage, repetition 1: plant height of resistant canola at 2 WAI Random Effects Estimate Standard Error Z Pr > Z Block 0.0 0.02 0.19 0.42 Residual 0.2 0.05 3.24 0.0006 Fixed Effects Num df Den df F Pr > F Cultivar 3 21 1.37 0.28 Inoculation 1 21 3.24 0.09 Cultivar*Inoculation 3 21 1.64 0.21 Contrast 1 21 5.40 0.03
Table A1.23. Controlled environment study – canola, napa cabbage, cabbage, repetition 1: plant height of resistant canola at 3 WAI Random Effects Estimate Standard Error Z Pr > Z Block 0.0 - - - Residual 0.2 0.05 3.46 0.0003 Fixed Effects Num df Den df F Pr > F Cultivar 3 21 0.90 0.46 Inoculation 1 21 1.68 0.21 Cultivar*Inoculation 3 21 0.98 0.42 Contrast 1 21 3.05 0.10 Table A1.24. Controlled environment study – canola, napa cabbage, cabbage, repetition 1: plant height of resistant canola at 4 WAI Random Effects Estimate Standard Error Z Pr > Z Block 0 - - - Residual 0.2 0.05 3.46 0.0003
121 Fixed Effects Num df Den df F Pr > F Cultivar 3 21 4.76 0.01 Inoculation 1 21 1.09 0.31 Cultivar*Inoculation 3 21 2.12 0.13 Contrast 1 21 3.54 0.07
Table A1.25. Controlled environment study – canola, napa cabbage, cabbage, repetition 1: plant height of resistant canola at 5 WAI Random Effects Estimate Standard Error Z Pr > Z Block 0.0 - - - Residual 0.5 0.14 3.46 0.0003 Fixed Effects Num df Den df F Pr > F Cultivar 3 21 11.15 0.0001 Inoculation 1 21 1.46 0.24 Cultivar*Inoculation 3 21 0.81 0.50 Contrast 1 21 3.27 0.08
Table A1.26. Controlled environment study – canola, napa cabbage, cabbage repetition 1: plant height at 6 WAI of resistant canola Random Effects Estimate Standard Error Z Pr > Z Block 0.0 - - - Residual 0.1 0.33 3.46 0.0003 Fixed Effects Num df Den df F Pr > F Cultivar 3 21 13.09 <0.0001 Inoculation 1 21 2.11 0.16 Cultivar*Inoculation 3 21 0.61 0.61 Contrast 1 21 3.30 0.08
Table A1.27. Controlled environment study – canola, napa cabbage, cabbage, repetition 1: Area under growth stairs (AUGS) resistant canola 2-6 WAI Random Effects Estimate Standard Error Z Pr > Z Block 0.0 - - - Residual 375.9 108.52 3.46 0.0003 Fixed Effects Num df Den df F Pr > F Cultivar 3 21 7.81 0.001 Inoculation 1 21 3.50 0.08 Cultivar*Inoculation 3 21 0.25 0.86 Contrast 1 21 2.49 0.13
122 Table A1.28. Controlled environment study – canola, napa cabbage, cabbage, repetition 1: Maturity Random Effects Estimate Standard Error Z Pr > Z Block 0.0 - - - Residual 264.9 76.47 3.46 0.0003 Fixed Effects Num df Den df F Pr > F Inoculation 1 21 0.00 1.00 Maturity 3 21 19.86 <0.0001 Inoculation*Maturity 3 21 0.38 0.77 Contrast (veg) Contrast (bud) Contrast (flower) Contrast (pod)
1 1 1 1
21 21 21 21
30.38 0.10 4.64 9.25
<0.0001 0.75 0.04 0.006
Table A1.29. Controlled environment study – canola, napa cabbage, cabbage: leaf length at 5 WAI of resistant napa cabbage Random Effects Estimate Standard Error Z Pr > Z Repetition 6.9 9.81 0.71 0.24 Block 0.2 0.16 1.01 0.16 Residual 0.6 0.11 5.10 <0.0001 Fixed Effects Num df Den df F Pr > F Cultivar 3 52 3.57 0.02 Inoculation 1 52 6.92 0.01 Cultivar*Inoculation 3 52 0.99 0.41 Contrast 1 52 7.76 0.007
123 Table A1.30. Controlled environment study – canola, napa cabbage, cabbage: biomass (dry shoot weight) of resistant napa cabbage Random Effects Estimate Standard Error Z Pr > Z Repetition 6.9 9.81 0.71 0.24 Block 0.2 0.16 1.01 0.16 Residual 0.6 0.11 5.10 <0.0001 Fixed Effects Num df Den df F Pr > F Cultivar 3 52 3.57 0.02 Inoculation 1 52 6.92 0.01 Cultivar*Inoculation 3 52 0.99 0.41 Contrast 1 52 7.76 0.007
Table A1.31. Controlled environment study – canola, napa cabbage, cabbage: leaf length at 6 WAI of resistant cabbage Random Effects Estimate Standard Error Z Pr > Z Repetition 3.2 4.62 0.70 0.24 Block 0.0 0.04 0.17 0.43 Residual 0.5 0.10 4.67 <0.0001 Fixed Effects Num df Den df F Pr > F Cultivar 3 44 5.50 0.00 Inoculation 1 44 2.85 0.09 Cultivar*Inoculation 3 44 0.85 0.47 Contrast 1 44 1.49 0.23
Table A1.32. Controlled environment study – canola, napa cabbage, cabbage: biomass (dry shoot weight, g) of resistant cabbage Random Effects Estimate Standard Error Z Pr > Z Repetition 7.1 11.8 0.61 0.27 Block 51.0 44.6 1.14 0.13 Residual 28.6 6.1 4.70 <0.0001 Fixed Effects Num df Den df F Pr > F Cultivar 3 44 0.71 0.55 Inoculation 1 44 0.00 0.95 Cultivar*Inoculation 3 44 0.48 0.70 Contrast 1 44 0.32 0.57
124 Table A1.33. Field trials – CI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 273 1.00 0.43 Repetition (R) 1 0 0.00 <0.0001 Site (S) 2 4840 17.71 0.0003 R x S 1 0 0.00 <0.0001 Error 12 273 Table A1.34. Field trials – DSI in susceptible canola Source df Mean Square F value Pr>F Block (B) 3 126 1.46 0.28 Repetition (R) 1 4225 48.73 <0.0001 Site (S) 2 6684 77.10 <0.0001 R x S 1 25 0.29 0.60 Error 12 87 Table A1.35. Field trials – Levene’s test, homogeneity of variance: plant height of resistant canola Source df Mean Square F value Pr>F Site x Repetition 4 11206 1.74 0.16 Error 55 6454 Table A1.36. Field trial, 2014 – Levene’s test, homogeneity of variance: plant height of resistant canola Source df Mean Square F value Pr>F Site (S) 1 13640 1.92 0.18 Error 22 7093 Table A1.37. Field trial, 2015 – Levene’s test, homogeneity of variance: plant height of resistant canola Source df Mean Square F value Pr>F Site (S) 2 15743 2.54 0.09 Error 33 5431 Table A1.38. Field trials – plant height at 8 WAI of resistant canola Source df Mean Square F value Pr>F Site (S) 3 27225 12.15 <0.0001 Repetition (R) 1 876 377.41 <0.0001 S x R 0 - - - Error 55 72 Table A1.39. Field trial, 2014 – plant height at 8 WAI of resistant canola Source df Mean Square F value Pr>F Site (S) 1 2330 30.55 <0.0001 Error 22 76
125 Table A1.40. Field trial, 2015 – plant height at 8 WAI of resistant canola Source df Mean Square F value Pr>F Site (S) 2 149 2.15 0.13 Error 33 69 Table A1.41. Field trials – Levene’s test, homogeneity of variance: biomass of resistant canola Source df Mean Square F value Pr>F S x R 4 15307 1.69 0.17 Error 55 6028 Table A1.42. Field trial, 2014 – Levene’s test, homogeneity of variance: biomass of resistant canola Source df Mean Square F value Pr>F Site (S) 2 4 x 109 2.30 0.14 Error 22 2 x 109 Table A1.43. Field trial, 2015 – Levene’s test, homogeneity of variance: biomass of resistant canola Source df Mean Square F value Pr>F Site (S) 2 15307 2.54 0.09 Error 33 6028 Table A1.44. Field trials – biomass of resistant canola Source df Mean Square F value Pr>F Site (S) 2 49447 1.66 0.20 Repetition (R) 1 48186 1.62 0.20 S x R 1 166218 5.59 0.02 Error 75 29754 Table A1.45. Field trial, 2014 – biomass Source df Mean Square F value Pr>F Site (S) 2 297131 9.50 0.006 Error 22 31286 Table A1.46. Field trial, 2015 – biomass Source df Mean Square F value Pr>F Site (S) 2 990 0.05 0.95 Error 33 18023 Table A1.47. Field trials – Levene’s test, homogeneity of variance: maturity of resistant canola Source df Mean Square F value Pr>F S x R 4 289000 4.77 0.0022 Error 55 60606
126 Table A1.48. Field trial, 2014 – Levene’s test, homogeneity of variance: maturity of resistant canola Source df Mean Square F value Pr>F Site (S) 1 33750 0.22 0.64 Error 22 151515 Table A1.49. Field trial, 2015 – Levene’s test, homogeneity of variance: maturity of resistant canola Source df Mean Square F value Pr>F Site (S) 2 0 - - Error 33 0 Table A1.50. Field trials – maturity of resistant canola Source df Mean Square F value Pr>F Site (S) 2 7725 63.41 <0.0001 Repetition (R) 1 46875 384.79 <0.0001 S x R 1 6075 49.87 <0.0001 Error 55 122 Table A1.51. Field trial, 2014 – maturity Source df Mean Square F value Pr>F Site (S) 1 12150 39.90 <0.0001 Error 22 305 Table A1.52. Field trial, 2015 – maturity Source df Mean Square F value Pr>F Site (S) 2 0 - - Error 33 0 Table A1.53. Field trials – CI in ‘susceptible’ napa cabbage Source df Mean Square F value Pr>F Block (B) 3 72 0.28 0.84 Repetition (R) 1 15188 59.56 <0.0001 Site (S) 2 2278 8.93 0.004 R x S 1 4556 17.87 0.001 Error 12 255 Table A1.54. Field trials – DSI in ‘susceptible’ napa cabbage Source df Mean Square F value Pr>F Block (B) 3 12 0.12 0.95 Repetition (R) 1 2756 36.67 0.0001 Site (S) 2 479 4.68 0.03 R x S 1 958 9.35 0.001 Error 12 102
127 Table A1.55. Field trial, 2014 – CI in susceptible napa cabbage Source df Mean Square F value Pr>F Site (S) 1 6171 11.87 0.02 Error 5 520 Table A1.56. Field trial, 2014 – DSI in susceptible napa cabbage Source df Mean Square F value Pr>F Site (S) 1 1230 5.82 0.06 Error 5 211 Table A1.57. Field trials – leaf length of resistant napa cabbage Source df Mean Square F value Pr>F Site (S) 2 1882 52.96 <0.0001 Repetition (R) 1 272 36.35 <0.0001 S x R 1 9 1.74 0.19 Error 67 5 Table A1.58. Field trials – biomass of resistant napa cabbage Source df Mean Square F value Pr>F Site (S) 2 1543411 5.68 0.005 Repetition (R) 1 1004429 3.70 0.06 S x R 1 41007 0.15 0.70 Error 67 271514 Table A1.59. Raw data for controlled environment study – canola: CI & DSI
Repetition Spores ml-1 Cultivar Block CI DSI 1 1x107 ACSN39 1 100.0 76.9 1 1x107 ACSN39 2 100.0 50.0 1 1x107 ACSN39 3 100.0 77.8 1 1x107 ACSN39 4 100.0 81.0 1 0 45H29 1 0.0 0.0 1 0 45H29 2 0.0 0.0 1 0 45H29 3 0.0 0.0 1 0 45H29 4 0.0 0.0 1 1x104 45H29 1 0.0 0.0 1 1x104 45H29 2 0.0 0.0 1 1x104 45H29 3 0.0 0.0 1 1x104 45H29 4 0.0 0.0 1 1x105 45H29 1 0.0 0.0 1 1x105 45H29 2 0.0 0.0 1 1x105 45H29 3 0.0 0.0 1 1x105 45H29 4 0.0 0.0 1 1x106 45H29 1 0.0 0.0
128 1 1x106 45H29 2 0.0 0.0 1 1x106 45H29 3 0.0 0.0 1 1x106 45H29 4 0.0 0.0 1 1x107 45H29 1 0.0 0.0 1 1x107 45H29 2 0.0 0.0 1 1x107 45H29 3 0.0 0.0 1 1x107 45H29 4 0.0 0.0 1 1x108 45H29 1 0.0 0.0 1 1x108 45H29 2 0.0 0.0 1 1x108 45H29 3 0.0 0.0 1 1x108 45H29 4 0.0 0.0 2 1x107 ACSN39 1 100.0 92.3 2 1x107 ACSN39 2 100.0 92.6 2 1x107 ACSN39 3 100.0 88.9 2 1x107 ACSN39 4 100.0 87.5 2 0 45H29 1 0.0 0.0 2 0 45H29 2 0.0 0.0 2 0 45H29 3 0.0 0.0 2 0 45H29 4 0.0 0.0 2 1x104 45H29 1 0.0 0.0 2 1x104 45H29 2 0.0 0.0 2 1x104 45H29 3 0.0 0.0 2 1x104 45H29 4 0.0 0.0 2 1x105 45H29 1 0.0 0.0 2 1x105 45H29 2 0.0 0.0 2 1x105 45H29 3 0.0 0.0 2 1x105 45H29 4 0.0 0.0 2 1x106 45H29 1 0.0 0.0 2 1x106 45H29 2 0.0 0.0 2 1x106 45H29 3 0.0 0.0 2 1x106 45H29 4 0.0 0.0 2 1x107 45H29 1 0.0 0.0 2 1x107 45H29 2 0.0 0.0 2 1x107 45H29 3 0.0 0.0 2 1x107 45H29 4 0.0 0.0 2 1x108 45H29 1 0.0 0.0 2 1x108 45H29 2 0.0 0.0 2 1x108 45H29 3 0.0 0.0 2 1x108 45H29 4 0.0 0.0 3 1x108 ACSN39 1 100.0 92.6 3 1x108 ACSN39 2 100.0 92.6 3 1x108 ACSN39 3 100.0 96.3 3 1x108 ACSN39 4 100.0 100.0
129 3 0 45H29 1 0.0 0.0 3 0 45H29 2 0.0 0.0 3 0 45H29 3 0.0 0.0 3 0 45H29 4 0.0 0.0 3 1x104 45H29 1 0.0 0.0 3 1x104 45H29 2 0.0 0.0 3 1x104 45H29 3 0.0 0.0 3 1x104 45H29 4 0.0 0.0 3 1x105 45H29 1 0.0 0.0 3 1x105 45H29 2 0.0 0.0 3 1x105 45H29 3 0.0 0.0 3 1x105 45H29 4 0.0 0.0 3 1x106 45H29 1 0.0 0.0 3 1x106 45H29 2 0.0 0.0 3 1x106 45H29 3 0.0 0.0 3 1x106 45H29 4 0.0 0.0 3 1x107 45H29 1 0.0 0.0 3 1x107 45H29 2 0.0 0.0 3 1x107 45H29 3 0.0 0.0 3 1x107 45H29 4 0.0 0.0 3 1x108 45H29 1 0.0 0.0 3 1x108 45H29 2 0.0 0.0 3 1x108 45H29 3 0.0 0.0 3 1x108 45H29 4 0.0 0.0
Table A1.60. Raw data for controlled environment study – canola: plant height
Repetition Spores g-1 soil Cultivar Block 2 WAI 3 WAI 4 WAI 5 WAI 6 WAI
1 0 45H29 1 3.99 7.82 17.77 25.67 30.74 1 0 45H29 2 3.29 6.82 16.70 24.07 27.38 1 0 45H29 3 3.25 7.25 18.20 24.55 28.93 1 0 45H29 4 3.50 6.88 14.16 27.44 36.56 1 1x104 45H29 1 3.50 7.11 14.55 25.45 31.53 1 1x104 45H29 2 3.44 5.84 14.26 24.91 28.98 1 1x104 45H29 3 3.04 6.54 13.06 21.65 28.77 1 1x104 45H29 4 3.38 6.16 15.52 26.56 33.41 1 1x105 45H29 1 3.78 6.20 15.08 24.58 29.28 1 1x105 45H29 2 3.96 8.70 23.57 34.45 36.21 1 1x105 45H29 3 3.63 6.90 17.82 28.20 31.43 1 1x105 45H29 4 3.87 6.67 15.16 26.05 28.86 1 1x106 45H29 1 3.49 5.79 12.98 23.47 28.10
130 1 1x106 45H29 2 3.43 5.73 12.71 23.06 27.29 1 1x106 45H29 3 3.46 6.35 17.11 29.38 32.23 1 1x106 45H29 4 3.47 6.26 13.44 22.73 27.16 1 1x107 45H29 1 3.93 5.51 10.33 17.33 18.27 1 1x107 45H29 2 4.10 6.60 15.32 25.41 28.58 1 1x107 45H29 3 3.49 6.99 16.81 23.43 26.72 1 1x107 45H29 4 2.95 4.92 11.41 20.24 23.65 1 1x108 45H29 1 3.87 5.84 11.50 19.75 22.01 1 1x108 45H29 2 2.73 4.88 12.44 20.71 26.96 1 1x108 45H29 3 3.37 6.26 13.75 24.40 33.50 1 1x108 45H29 4 3.10 5.11 13.57 21.04 25.36 2 0 45H29 1 3.06 4.57 6.43 17.77 26.36 2 0 45H29 2 4.26 8.06 14.19 32.20 34.08 2 0 45H29 3 4.98 9.33 14.02 26.70 33.88 2 0 45H29 4 4.73 9.40 17.54 29.24 30.56 2 1x104 45H29 1 3.03 3.63 4.50 7.82 13.39 2 1x104 45H29 2 3.08 4.63 6.96 15.82 24.88 2 1x104 45H29 3 5.11 9.08 14.60 34.86 34.81 2 1x104 45H29 4 3.93 6.54 8.28 14.75 19.63 2 1x105 45H29 1 2.93 3.63 4.68 8.38 14.23 2 1x105 45H29 2 3.90 7.24 10.66 21.73 27.06 2 1x105 45H29 3 4.27 7.01 10.62 20.87 27.27 2 1x105 45H29 4 5.04 8.32 14.38 25.91 29.43 2 1x106 45H29 1 2.64 3.30 4.38 8.42 14.46 2 1x106 45H29 2 3.55 6.01 8.76 18.35 28.44 2 1x106 45H29 3 4.90 9.36 16.29 33.14 34.92 2 1x106 45H29 4 4.03 8.55 12.86 25.59 28.13 2 1x107 45H29 1 2.89 3.68 5.29 11.08 18.68 2 1x107 45H29 2 3.55 5.36 7.62 15.29 22.51 2 1x107 45H29 3 4.34 7.32 12.28 25.79 26.55 2 1x107 45H29 4 4.00 6.28 8.99 19.50 25.59 2 1x108 45H29 1 2.85 3.78 4.90 9.00 17.61 2 1x108 45H29 2 3.50 4.88 7.24 15.18 22.11 2 1x108 45H29 3 5.32 9.19 14.35 30.58 33.75 2 1x108 45H29 4 4.17 6.29 11.32 25.17 29.29 3 0 45H29 1 4.09 7.44 15.84 . 34.42 3 0 45H29 2 3.88 7.45 21.49 . 32.81 3 0 45H29 3 3.57 5.80 12.68 . 31.69 3 0 45H29 4 3.41 6.23 16.39 . 30.55 3 1x104 45H29 1 3.71 6.79 16.47 . 26.56 3 1x104 45H29 2 3.76 6.20 14.15 . 32.71 3 1x104 45H29 3 3.23 6.37 16.08 . 30.21 3 1x104 45H29 4 3.88 8.04 21.50 . 32.04
131 3 1x105 45H29 1 3.97 7.03 13.26 . 27.89 3 1x105 45H29 2 3.84 6.55 15.94 . 31.59 3 1x105 45H29 3 3.15 6.35 16.56 . 30.17 3 1x105 45H29 4 4.05 7.93 17.53 . 29.98 3 1x106 45H29 1 4.04 6.97 16.40 . 30.50 3 1x106 45H29 2 3.45 7.58 19.11 . 32.80 3 1x106 45H29 3 3.13 5.25 10.33 . 28.72 3 1x106 45H29 4 3.54 6.38 13.31 . 28.56 3 1x107 45H29 1 4.43 7.70 15.69 . 27.28 3 1x107 45H29 2 3.70 6.88 14.82 . 32.10 3 1x107 45H29 3 3.36 6.82 15.73 . 31.28 3 1x107 45H29 4 3.80 7.13 16.94 . 30.27 3 1x108 45H29 1 3.28 5.96 12.68 . 26.84 3 1x108 45H29 2 3.95 7.10 16.49 . 30.01 3 1x108 45H29 3 3.28 6.03 14.78 . 28.71 3 1x108 45H29 4 4.03 8.16 16.82 . 29.33
Table A1.61. Raw data for controlled environment study – canola: biomass (dry shoot weight) of resistant canola
Repetition Spores ml-1 Block Mean dry shoot weight of 10 plants (g)
1 0 1 6.13 1 0 2 5.78 1 0 3 5.07 1 0 4 8.71 1 1x104 1 5.08 1 1x104 2 5.69 1 1x104 3 5.75 1 1x104 4 6.56 1 1x105 1 4.78 1 1x105 2 6.98 1 1x105 3 7.96 1 1x105 4 7.83 1 1x106 1 4.30 1 1x106 2 4.91 1 1x106 3 6.78 1 1x106 4 6.74 1 1x107 1 5.64 1 1x107 2 4.86 1 1x107 3 6.16 1 1x107 4 7.55 1 1x108 1 4.24 1 1x108 2 4.48
132 1 1x108 3 5.55 1 1x108 4 6.43 2 0 1 . 2 0 2 5.40 2 0 3 4.30 2 0 4 2.20 2 1x104 1 3.10 2 1x104 2 3.10 2 1x104 3 2.90 2 1x104 4 3.40 2 1x105 1 2.20 2 1x105 2 4.10 2 1x105 3 4.20 2 1x105 4 3.20 2 1x106 1 2.40 2 1x106 2 3.50 2 1x106 3 5.10 2 1x106 4 2.80 2 1x107 1 2.50 2 1x107 2 3.30 2 1x107 3 2.30 2 1x107 4 3.40 2 1x108 1 2.60 2 1x108 2 2.90 2 1x108 3 2.60 2 1x108 4 3.40 3 0 1 11.33 3 0 2 11.56 3 0 3 9.56 3 0 4 8.56 3 1x104 1 7.89 3 1x104 2 10.67 3 1x104 3 8.89 3 1x104 4 8.70 3 1x105 1 9.13 3 1x105 2 10.90 3 1x105 3 8.89 3 1x105 4 8.73 3 1x106 1 7.90 3 1x106 2 10.20 3 1x106 3 8.50 3 1x106 4 8.40 3 1x107 1 8.20 3 1x107 2 9.20
133 3 1x107 3 8.30 3 1x107 4 9.60 3 1x108 1 9.00 3 1x108 2 8.40 3 1x108 3 9.10 3 1x108 4 8.40 Table A1.62. Raw data for controlled environment study – canola: maturity (percentage of plants at seedpod stage) of resistant canola
Repetition Resting spores g-1 soil Block Percentage of plants at pod development stage
1 0 1 72.7 1 0 2 90.9 1 0 3 60.0 1 0 4 91.7 1 1x104 1 75.0 1 1x104 2 91.7 1 1x104 3 81.8 1 1x104 4 83.3 1 1x105 1 83.3 1 1x105 2 100.0 1 1x105 3 90.9 1 1x105 4 90.0 1 1x106 1 75.0 1 1x106 2 81.8 1 1x106 3 100.0 1 1x106 4 58.3 1 1x107 1 54.5 1 1x107 2 58.3 1 1x107 3 72.7 1 1x107 4 54.5 1 1x108 1 54.5 1 1x108 2 40.0 1 1x108 3 63.6 1 1x108 4 40.0
134 Table A1.63. Raw data for large pot studies – outdoor, 2014 and 2015: CI & DSI
Year Repetition Spores g-1 soil Cultivar CI DI
2014 1 1x107 ACSN39 12.5 8.3 2014 1 1x107 ACSN39 42.9 19.0 2014 1 1x107 ACSN39 57.1 28.6 2014 1 1x107 ACSN39 28.6 9.5 2014 1 0 45H29 0.0 0.0 2014 1 0 45H29 0.0 0.0 2014 1 0 45H29 0.0 0.0 2014 1 0 45H29 0.0 0.0 2014 1 1x103 45H29 0.0 0.0 2014 1 1x103 45H29 0.0 0.0 2014 1 1x103 45H29 0.0 0.0 2014 1 1x103 45H29 0.0 0.0 2014 1 1x104 45H29 0.0 0.0 2014 1 1x104 45H29 0.0 0.0 2014 1 1x104 45H29 0.0 0.0 2014 1 1x104 45H29 0.0 0.0 2014 1 1x105 45H29 0.0 0.0 2014 1 1x105 45H29 0.0 0.0 2014 1 1x105 45H29 0.0 0.0 2014 1 1x105 45H29 0.0 0.0 2014 1 1x106 45H29 0.0 0.0 2014 1 1x106 45H29 0.0 0.0 2014 1 1x106 45H29 0.0 0.0 2014 1 1x106 45H29 0.0 0.0 2014 1 1x107 45H29 0.0 0.0 2014 1 1x107 45H29 0.0 0.0 2014 1 1x107 45H29 0.0 0.0 2014 1 1x107 45H29 0.0 0.0 2015 1 1x107 ACSN39 100.0 83.3 2015 1 1x107 ACSN39 100.0 80.0 2015 1 1x107 ACSN39 100.0 80.0 2015 1 1x107 ACSN39 90.0 70.0 2015 1 0 45H29 0.0 0.0 2015 1 0 45H29 0.0 0.0 2015 1 0 45H29 0.0 0.0 2015 1 0 45H29 0.0 0.0 2015 1 1x103 45H29 0.0 0.0 2015 1 1x103 45H29 0.0 0.0 2015 1 1x103 45H29 0.0 0.0 2015 1 1x103 45H29 0.0 0.0
135 2015 1 1x104 45H29 0.0 0.0 2015 1 1x104 45H29 0.0 0.0 2015 1 1x104 45H29 0.0 0.0 2015 1 1x104 45H29 0.0 0.0 2015 1 1x105 45H29 0.0 0.0 2015 1 1x105 45H29 0.0 0.0 2015 1 1x105 45H29 0.0 0.0 2015 1 1x105 45H29 0.0 0.0 2015 1 1x106 45H29 0.0 0.0 2015 1 1x106 45H29 0.0 0.0 2015 1 1x106 45H29 0.0 0.0 2015 1 1x106 45H29 0.0 0.0 2015 1 1x107 45H29 10.0 10.0 2015 1 1x107 45H29 20.0 13.3 2015 1 1x107 45H29 0.0 0.0 2015 1 1x107 45H29 0.0 0.0 2015 2 1x107 ACSN39 100.0 96.7 2015 2 1x107 ACSN39 100.0 93.3 2015 2 1x107 ACSN39 100.0 100.0 2015 2 1x107 ACSN39 100.0 96.7 2015 2 0 45H29 0.0 0.0 2015 2 0 45H29 0.0 0.0 2015 2 0 45H29 0.0 0.0 2015 2 0 45H29 0.0 0.0 2015 2 1x103 45H29 0.0 0.0 2015 2 1x103 45H29 0.0 0.0 2015 2 1x103 45H29 0.0 0.0 2015 2 1x103 45H29 10.0 3.3 2015 2 1x104 45H29 10.0 6.7 2015 2 1x104 45H29 0.0 0.0 2015 2 1x104 45H29 0.0 0.0 2015 2 1x104 45H29 0.0 0.0 2015 2 1x105 45H29 0.0 0.0 2015 2 1x105 45H29 10.0 6.7 2015 2 1x105 45H29 0.0 0.0 2015 2 1x105 45H29 0.0 0.0 2015 2 1x106 45H29 0.0 0.0 2015 2 1x106 45H29 0.0 0.0 2015 2 1x106 45H29 0.0 0.0 2015 2 1x106 45H29 20.0 10.0 2015 2 1x107 45H29 10.0 3.3 2015 2 1x107 45H29 10.0 10.0 2015 2 1x107 45H29 20.0 20.0
136 2015 2 1x107 45H29 20.0 13.3
Table A1.64. Raw data for large pot studies – outdoor, 2014 and 2015: plant height of resistant canola
Year Repetition Spores g-1 soil Cultivar
2 WAI
3 WAI
4 WAI
5 WAI
6 WAI
7 WAI
2014 1 0 45H29 1.65 2.53 4.09 7.24 10.08 . 2014 1 0 45H29 1.91 3.00 4.38 8.60 11.69 . 2014 1 0 45H29 1.72 2.35 3.05 5.77 7.40 . 2014 1 0 45H29 1.28 2.32 3.08 4.66 5.80 . 2014 1 1x103 45H29 1.45 2.22 3.61 6.81 8.45 . 2014 1 1x103 45H29 1.87 3.05 4.67 10.72 14.06 . 2014 1 1x103 45H29 1.88 2.79 3.74 6.80 9.12 . 2014 1 1x103 45H29 0.78 1.66 2.82 4.78 6.90 . 2014 1 1x104 45H29 1.32 2.03 2.84 6.36 8.10 . 2014 1 1x104 45H29 1.63 2.77 4.28 8.13 10.31 . 2014 1 1x104 45H29 1.89 2.58 3.47 7.03 9.73 . 2014 1 1x104 45H29 0.98 2.26 2.96 5.02 6.48 . 2014 1 1x105 45H29 2.10 3.15 5.69 12.74 14.76 . 2014 1 1x105 45H29 1.29 2.22 3.30 7.36 10.14 . 2014 1 1x105 45H29 1.50 2.31 3.59 8.09 11.58 . 2014 1 1x105 45H29 1.36 2.12 3.40 5.74 7.04 . 2014 1 1x106 45H29 1.39 2.30 4.01 9.93 13.76 . 2014 1 1x106 45H29 1.81 2.74 4.44 8.87 16.05 . 2014 1 1x106 45H29 1.40 2.10 2.74 4.99 6.36 . 2014 1 1x106 45H29 1.12 1.82 2.54 4.54 6.14 . 2014 1 1x107 45H29 1.48 2.16 3.38 7.08 9.25 . 2014 1 1x107 45H29 1.89 2.76 3.67 8.15 10.75 . 2014 1 1x107 45H29 1.09 1.61 2.00 3.61 4.55 . 2014 1 1x107 45H29 1.22 2.38 2.98 4.80 6.10 . 2015 1 0 45H29 5.40 11.36 25.37 49.89 57.06 60.60 2015 1 0 45H29 7.19 17.63 38.70 50.17 50.71 57.15 2015 1 0 45H29 8.75 23.41 37.71 50.77 50.22 53.85 2015 1 0 45H29 6.06 17.90 30.11 47.65 49.51 52.90 2015 1 1x103 45H29 10.86 15.81 33.97 48.86 50.96 55.90 2015 1 1x103 45H29 6.62 18.98 36.07 50.56 52.21 57.64 2015 1 1x103 45H29 8.87 23.14 33.15 38.38 46.55 47.87 2015 1 1x103 45H29 6.14 21.14 35.13 48.27 48.40 52.24 2015 1 1x104 45H29 6.31 10.40 32.01 43.70 47.00 53.86 2015 1 1x104 45H29 6.08 14.53 32.15 43.98 45.44 53.99 2015 1 1x104 45H29 5.49 14.38 38.00 55.59 55.91 59.83
137 2015 1 1x104 45H29 7.79 22.94 33.20 44.21 47.97 55.34 2015 1 1x105 45H29 5.97 17.92 38.33 44.08 51.67 55.22 2015 1 1x105 45H29 6.47 16.48 33.39 52.72 51.37 58.87 2015 1 1x105 45H29 8.95 24.56 39.15 48.44 48.00 52.14 2015 1 1x105 45H29 5.13 13.58 29.94 48.63 50.65 56.86 2015 1 1x106 45H29 5.98 14.74 29.20 42.21 48.17 50.47 2015 1 1x106 45H29 8.72 24.62 33.84 46.04 46.41 55.87 2015 1 1x106 45H29 6.13 13.30 34.80 49.46 56.83 57.52 2015 1 1x106 45H29 6.27 16.79 28.70 47.22 47.68 54.48 2015 1 1x107 45H29 4.98 9.34 29.44 38.90 45.39 48.30 2015 1 1x107 45H29 5.35 9.94 23.26 44.62 44.94 53.71 2015 1 1x107 45H29 5.73 16.77 32.50 51.84 53.65 59.70 2015 1 1x107 45H29 4.53 15.34 34.52 43.83 44.27 52.86 2015 2 0 45H29 2.85 7.89 18.40 34.78 66.24 69.74 2015 2 0 45H29 2.82 12.19 18.17 52.13 77.23 78.14 2015 2 0 45H29 2.39 12.91 21.71 57.88 80.43 78.42 2015 2 0 45H29 2.88 12.27 20.29 46.14 70.07 76.03 2015 2 1x103 45H29 3.42 11.68 22.81 44.44 67.69 74.02 2015 2 1x103 45H29 2.68 5.37 13.17 35.57 62.31 70.77 2015 2 1x103 45H29 3.13 14.20 22.05 52.95 72.11 71.93 2015 2 1x103 45H29 2.79 12.70 20.33 51.46 69.23 72.03 2015 2 1x104 45H29 3.48 11.49 20.81 50.59 72.27 77.91 2015 2 1x104 45H29 3.17 9.29 20.39 45.18 68.54 67.97 2015 2 1x104 45H29 3.30 11.83 23.64 63.42 81.54 83.79 2015 2 1x104 45H29 2.68 11.29 23.02 57.30 78.48 81.29 2015 2 1x105 45H29 3.20 11.77 21.21 39.72 63.12 65.57 2015 2 1x105 45H29 2.22 10.89 20.49 49.48 72.79 75.58 2015 2 1x105 45H29 2.29 11.30 23.31 54.26 76.51 76.87 2015 2 1x105 45H29 2.50 10.57 22.90 49.64 66.28 68.69 2015 2 1x106 45H29 2.71 9.67 20.02 36.21 54.20 59.94 2015 2 1x106 45H29 3.51 9.67 19.84 50.46 78.14 76.04 2015 2 1x106 45H29 2.13 7.59 13.70 39.53 65.97 66.14 2015 2 1x106 45H29 3.02 10.95 18.86 47.21 64.54 65.00 2015 2 1x107 45H29 3.57 11.68 20.49 41.05 59.60 69.34 2015 2 1x107 45H29 3.08 10.19 21.87 49.73 81.31 80.07 2015 2 1x107 45H29 3.38 17.68 26.52 58.59 73.63 77.27 2015 2 1x107 45H29 2.33 9.66 20.71 47.38 65.74 71.54
Table A1.65. Raw data for large pot studies – outdoor, 2014 and 2015: biomass (dry shoot weight) of resistant canola Year Repetition Resting spores g^1 soil Block Mean dry shoot
138 weight of 10 plants (g)
2014 1 0 1 275.71 2014 1 0 2 316.00 2014 1 0 3 175.50 2014 1 0 4 186.50 2014 1 1x103 1 152.89 2014 1 1x103 2 248.11 2014 1 1x103 3 146.50 2014 1 1x103 4 28.50 2014 1 1x104 1 250.67 2014 1 1x104 2 108.67 2014 1 1x104 3 119.50 2014 1 1x104 4 277.40 2014 1 1x105 1 210.20 2014 1 1x105 2 210.50 2014 1 1x105 3 162.71 2014 1 1x105 4 199.60 2014 1 1x106 1 88.60 2014 1 1x106 2 80.67 2014 1 1x106 3 251.40 2014 1 1x106 4 123.25 2014 1 1x107 1 104.75 2014 1 1x107 2 182.67 2014 1 1x107 3 101.67 2014 1 1x107 4 42.00 2015 1 0 1 654.00 2015 1 0 2 702.20 2015 1 0 3 672.20 2015 1 0 4 771.80 2015 1 1x103 1 768.20 2015 1 1x103 2 694.40 2015 1 1x103 3 816.80 2015 1 1x103 4 660.00 2015 1 1x104 1 683.20 2015 1 1x104 2 717.60 2015 1 1x104 3 715.40 2015 1 1x104 4 764.20 2015 1 1x105 1 734.60 2015 1 1x105 2 755.20 2015 1 1x105 3 622.60 2015 1 1x105 4 665.60 2015 1 1x106 1 644.20 2015 1 1x106 2 727.80
139 2015 1 1x106 3 687.20 2015 1 1x106 4 824.80 2015 1 1x107 1 1023.80 2015 1 1x107 2 738.60 2015 1 1x107 3 701.00 2015 1 1x107 4 619.40 2015 2 0 1 286.20 2015 2 0 2 193.60 2015 2 0 3 225.60 2015 2 0 4 215.60 2015 2 1x103 1 215.80 2015 2 1x103 2 128.00 2015 2 1x103 3 305.20 2015 2 1x103 4 262.20 2015 2 1x104 1 242.00 2015 2 1x104 2 293.20 2015 2 1x104 3 380.75 2015 2 1x104 4 338.80 2015 2 1x105 1 302.20 2015 2 1x105 2 261.00 2015 2 1x105 3 251.20 2015 2 1x105 4 354.40 2015 2 1x106 1 263.20 2015 2 1x106 2 200.00 2015 2 1x106 3 245.20 2015 2 1x106 4 257.40 2015 2 1x107 1 251.50 2015 2 1x107 2 229.50 2015 2 1x107 3 418.67 2015 2 1x107 4 218.20 Table A1.66. Raw data for large pot studies – outdoor, 2014 and 2015: maturity (% plants at flowering and seedpod development stages) of resistant canola
Year Repetition Resting spores g-1 soil Block % plants at flowering stage
% plants at pod stage
2014 1 0 1 50.0 0.0 2014 1 0 2 57.1 0.0 2014 1 0 3 0.0 0.0 2014 1 0 4 25.0 0.0
140 2014 1 1x103 1 30.0 0.0 2014 1 1x103 2 44.4 0.0 2014 1 1x103 3 25.0 0.0 2014 1 1x103 4 0.0 0.0 2014 1 1x104 1 14.3 0.0 2014 1 1x104 2 44.4 0.0 2014 1 1x104 3 25.0 0.0 2014 1 1x104 4 25.0 0.0 2014 1 1x105 1 88.9 0.0 2014 1 1x105 2 42.9 0.0 2014 1 1x105 3 42.9 0.0 2014 1 1x105 4 40.0 0.0 2014 1 1x106 1 60.0 0.0 2014 1 1x106 2 66.7 0.0 2014 1 1x106 3 20.0 0.0 2014 1 1x106 4 20.0 0.0 2014 1 1x107 1 25.0 0.0 2014 1 1x107 2 80.0 0.0 2014 1 1x107 3 16.7 0.0 2014 1 1x107 4 0.0 0.0 2015 1 0 1 0.0 100.0 2015 1 0 2 0.0 100.0 2015 1 0 3 0.0 100.0 2015 1 0 4 0.0 100.0 2015 1 1x103 1 0.0 100.0 2015 1 1x103 2 0.0 100.0 2015 1 1x103 3 0.0 100.0 2015 1 1x103 4 0.0 100.0 2015 1 1x104 1 0.0 100.0 2015 1 1x104 2 0.0 100.0 2015 1 1x104 3 0.0 100.0 2015 1 1x104 4 0.0 100.0 2015 1 1x105 1 0.0 100.0 2015 1 1x105 2 0.0 100.0 2015 1 1x105 3 0.0 100.0 2015 1 1x105 4 0.0 100.0 2015 1 1x106 1 0.0 100.0 2015 1 1x106 2 0.0 100.0 2015 1 1x106 3 0.0 100.0 2015 1 1x106 4 0.0 100.0 2015 1 1x107 1 0.0 100.0 2015 1 1x107 2 0.0 100.0 2015 1 1x107 3 0.0 100.0
141 2015 1 1x107 4 0.0 100.0 2015 2 0 1 0.0 100.0 2015 2 0 2 0.0 100.0 2015 2 0 3 0.0 100.0 2015 2 0 4 0.0 100.0 2015 2 1x103 1 0.0 100.0 2015 2 1x103 2 0.0 100.0 2015 2 1x103 3 0.0 100.0 2015 2 1x103 4 0.0 100.0 2015 2 1x104 1 0.0 100.0 2015 2 1x104 2 0.0 100.0 2015 2 1x104 3 0.0 100.0 2015 2 1x104 4 0.0 100.0 2015 2 1x105 1 0.0 100.0 2015 2 1x105 2 0.0 100.0 2015 2 1x105 3 0.0 100.0 2015 2 1x105 4 0.0 100.0 2015 2 1x106 1 0.0 100.0 2015 2 1x106 2 0.0 100.0 2015 2 1x106 3 0.0 100.0 2015 2 1x106 4 0.0 100.0 2015 2 1x107 1 0.0 100.0 2015 2 1x107 2 0.0 100.0 2015 2 1x107 3 0.0 100.0 2015 2 1x107 4 0.0 100.0
Table A1.67. Raw data for controlled environment study – canola: CI & DSI
Repetition Spores ml-1 Cultivar Block CI DSI
1 1x106 46A76 1 33.3 11.1 1 1x106 46A76 2 0.0 0.0 1 1x106 46A76 3 22.2 7.4 1 1x106 46A76 4 14.3 4.8 1 0 46A76 1 0.0 0.0 1 0 46A76 2 0.0 0.0 1 0 46A76 3 0.0 0.0 1 0 46A76 4 0.0 0.0 1 1x106 45H29 1 0.0 0.0 1 1x106 45H29 2 0.0 0.0 1 1x106 45H29 3 0.0 0.0 1 1x106 45H29 4 0.0 0.0 1 0 45H29 1 0.0 0.0 1 0 45H29 2 0.0 0.0
142 1 0 45H29 3 0.0 0.0 1 0 45H29 4 0.0 0.0 1 1x106 7367 1 0.0 0.0 1 1x106 7367 2 0.0 0.0 1 1x106 7367 3 0.0 0.0 1 1x106 7367 4 0.0 0.0 1 0 7367 1 0.0 0.0 1 0 7367 2 0.0 0.0 1 0 7367 3 0.0 0.0 1 0 7367 4 0.0 0.0 1 1x106 7377 1 0.0 0.0 1 1x106 7377 2 0.0 0.0 1 1x106 7377 3 0.0 0.0 1 1x106 7377 4 0.0 0.0 1 0 7377 1 0.0 0.0 1 0 7377 2 0.0 0.0 1 0 7377 3 0.0 0.0 1 0 7377 4 0.0 0.0 2 1x106 ACSN39 1 66.7 40.7 2 1x106 ACSN39 2 50.0 25.0 2 1x106 ACSN39 3 44.4 25.9 2 1x106 ACSN39 4 88.9 37.0 2 0 ACSN39 1 0.0 0.0 2 0 ACSN39 2 0.0 0.0 2 0 ACSN39 3 0.0 0.0 2 0 ACSN39 4 0.0 0.0 2 1x106 45H29 1 0.0 0.0 2 1x106 45H29 2 0.0 0.0 2 1x106 45H29 3 0.0 0.0 2 1x106 45H29 4 0.0 0.0 2 0 45H29 1 0.0 0.0 2 0 45H29 2 0.0 0.0 2 0 45H29 3 0.0 0.0 2 0 45H29 4 0.0 0.0 2 1x106 7367 1 0.0 0.0 2 1x106 7367 2 0.0 0.0 2 1x106 7367 3 0.0 0.0 2 1x106 7367 4 0.0 0.0 2 0 7367 1 0.0 0.0 2 0 7367 2 0.0 0.0 2 0 7367 3 0.0 0.0 2 0 7367 4 0.0 0.0 2 1x106 7377 1 0.0 0.0
143 2 1x106 7377 2 0.0 0.0 2 1x106 7377 3 0.0 0.0 2 1x106 7377 4 0.0 0.0 2 0 7377 1 0.0 0.0 2 0 7377 2 0.0 0.0 2 0 7377 3 0.0 0.0 2 0 7377 4 0.0 0.0
144 Table A1.68 Raw data for controlled environment study – canola: plant height Repetition Spores ml-1 Cultivar Block 2 WAI 3 WAI 4 WAI 5 WAI 6 WAI 1 1x106 46A76 1 2.80 3.54 3.97 4.72 5.45 1 1x106 46A76 2 3.33 . 3.52 4.29 4.59 1 1x106 46A76 3 2.79 3.53 3.85 4.98 6.28 1 1x106 46A76 4 3.12 . 3.70 4.14 4.50 1 1x106 45H29 1 2.38 3.18 3.58 4.63 5.93 1 1x106 45H29 2 2.77 3.19 3.93 4.93 6.16 1 1x106 45H29 3 3.57 3.87 5.17 6.66 9.04 1 1x106 45H29 4 2.76 3.03 3.70 5.00 6.16 1 1x106 7377 1 2.35 2.99 3.05 3.61 4.25 1 1x106 7377 2 2.88 3.29 3.41 3.99 4.92 1 1x106 7377 3 1.56 1.78 2.60 3.20 3.60 1 1x106 7377 4 2.44 3.45 3.83 4.55 4.80 1 1x106 7367 1 2.75 3.32 3.47 4.29 5.26 1 1x106 7367 2 2.37 2.59 3.06 3.23 3.58 1 1x106 7367 3 2.41 3.07 3.04 3.53 4.08 1 1x106 7367 4 2.31 2.76 2.71 3.01 4.05 1 0 46A76 1 3.00 3.30 3.40 3.60 4.10 1 0 46A76 2 2.93 3.47 3.45 3.90 4.42 1 0 46A76 3 3.34 3.69 3.77 4.68 5.53 1 0 46A76 4 2.30 2.88 3.05 4.55 6.05 1 0 45H29 1 2.68 3.28 3.89 5.33 7.16 1 0 45H29 2 3.43 4.00 4.34 7.20 9.89 1 0 45H29 3 2.67 2.71 4.03 6.38 9.55 1 0 45H29 4 2.92 3.36 3.86 4.56 5.68 1 0 7377 1 3.26 3.74 3.90 5.04 5.63 1 0 7377 2 3.28 3.36 3.68 4.16 4.62 1 0 7377 3 3.08 3.38 3.99 4.44 5.20 1 0 7377 4 2.34 2.96 3.45 4.00 4.83 1 0 7367 1 2.48 2.96 3.10 3.40 4.00 1 0 7367 2 3.02 3.41 3.80 3.92 4.80 1 0 7367 3 2.75 3.30 3.20 3.61 4.19 1 0 7367 4 3.08 3.64 4.13 4.65 5.37 2 1x106 ACSN39 1 2.68 3.12 4.10 9.42 14.58 2 1x106 ACSN39 2 2.74 3.14 4.54 13.28 16.81 2 1x106 ACSN39 3 2.70 3.34 4.71 12.61 19.20 2 1x106 ACSN39 4 3.29 3.73 5.46 15.26 21.41 2 1x106 45H29 1 3.12 4.57 8.55 18.78 22.17 2 1x106 45H29 2 2.82 3.64 5.74 15.13 21.84 2 1x106 45H29 3 3.17 4.59 8.90 23.63 26.72 2 1x106 45H29 4 3.89 5.21 10.68 20.15 28.96
145 2 1x106 7377 1 2.86 3.49 4.96 13.21 20.33 2 1x106 7377 2 3.04 3.39 5.15 14.57 21.82 2 1x106 7377 3 2.74 3.16 4.00 10.13 14.53 2 1x106 7377 4 3.73 4.26 5.73 11.77 16.89 2 1x106 7367 1 3.03 3.38 4.21 8.06 11.03 2 1x106 7367 2 3.13 3.69 5.02 14.21 22.05 2 1x106 7367 3 3.35 3.68 4.90 11.73 15.94 2 1x106 7367 4 4.18 5.61 8.81 17.78 21.44 2 0 ACSN39 1 2.64 3.51 5.44 14.14 21.79 2 0 ACSN39 2 2.96 3.26 4.10 9.69 15.41 2 0 ACSN39 3 2.90 3.18 4.76 14.20 19.23 2 0 ACSN39 4 3.60 4.10 5.63 13.45 21.48 2 0 45H29 1 3.13 4.58 7.59 20.62 25.22 2 0 45H29 2 2.80 4.01 9.60 25.02 27.96 2 0 45H29 3 3.24 4.26 7.01 15.51 20.94 2 0 45H29 4 3.91 5.51 10.26 27.09 30.57 2 0 7377 1 2.84 3.54 4.92 12.02 17.41 2 0 7377 2 2.84 3.42 5.28 13.76 19.09 2 0 7377 3 2.98 3.56 5.34 13.83 17.19 2 0 7377 4 2.91 3.53 4.28 9.03 12.39 2 0 7367 1 3.54 4.10 6.26 16.47 21.75 2 0 7367 2 3.09 3.79 5.05 12.05 18.31 2 0 7367 3 3.60 4.07 5.77 14.08 20.37 2 0 7367 4 4.25 5.01 6.78 14.33 18.24
Table A1.69. Raw data for controlled environment study – canola: biomass (dry shoot weight)
Repetition Cultivar Resting spores ml-1 Block Mean dry shoot weight of 10 plants (g)
1 46A76 1x106 1 3.17 1 46A76 1x106 2 3.53 1 46A76 1x106 3 3.36 1 46A76 1x106 4 2.91 1 46A76 0 1 2.52 1 46A76 0 2 3.28 1 46A76 0 3 3.13 1 46A76 0 4 3.15 1 45H29 1x106 1 2.69 1 45H29 1x106 2 3.11 1 45H29 1x106 3 2.76 1 45H29 1x106 4 0.75 1 45H29 0 1 2.50 1 45H29 0 2 2.80
146 1 45H29 0 3 2.94 1 45H29 0 4 3.61 1 7367 1x106 1 3.07 1 7367 1x106 2 2.83 1 7367 1x106 3 2.10 1 7367 1x106 4 2.55 1 7367 0 1 3.58 1 7367 0 2 2.21 1 7367 0 3 2.47 1 7367 0 4 2.53 1 7377 1x106 1 2.61 1 7377 1x106 2 3.21 1 7377 1x106 3 2.75 1 7377 1x106 4 2.78 1 7377 0 1 2.40 1 7377 0 2 2.08 1 7377 0 3 2.31 1 7377 0 4 2.55 2 ACSN39 1x106 1 15.54 2 ACSN39 1x106 2 10.89 2 ACSN39 1x106 3 11.91 2 ACSN39 1x106 4 12.78 2 ACSN39 0 1 15.18 2 ACSN39 0 2 13.26 2 ACSN39 0 3 13.39 2 ACSN39 0 4 12.95 2 45H29 1x106 1 14.82 2 45H29 1x106 2 14.40 2 45H29 1x106 3 13.97 2 45H29 1x106 4 13.88 2 45H29 0 1 15.70 2 45H29 0 2 13.46 2 45H29 0 3 10.36 2 45H29 0 4 15.16 2 7367 1x106 1 14.16 2 7367 1x106 2 12.28 2 7367 1x106 3 11.13 2 7367 1x106 4 14.39 2 7367 0 4 9.70 2 7367 0 1 11.36 2 7367 0 2 12.80 2 7367 0 3 11.81 2 7377 1x106 1 15.56
147 2 7377 1x106 2 13.15 2 7377 1x106 3 11.88 2 7377 1x106 4 11.83 2 7377 0 1 16.12 2 7377 0 2 14.27 2 7377 0 3 11.23 2 7377 0 4 10.98
Table A1.70. Raw data for controlled environment study – canola: maturity: % plants at flowering and seedpod development stages
Repetition Cultivar Resting spores g^1 soil Block
% plants at flowering stage
% plants at pod stage
1 45H29 1x106 1 25.0 0.0 1 45H29 1x106 2 0.0 0.0 1 45H29 1x106 3 14.3 14.3 1 45H29 1x106 4 0.0 0.0 1 45H29 0 1 0.0 0.0 1 45H29 0 2 12.5 25.0 1 45H29 0 3 0.0 25.0 1 45H29 0 4 0.0 0.0 2 ACSN39 1x106 1 100.0 0.0 2 ACSN39 1x106 2 50.0 37.5 2 ACSN39 1x106 3 90.0 0.0 2 ACSN39 1x106 4 77.8 22.2 2 ACSN39 0 1 70.0 30.0 2 ACSN39 0 2 80.0 10.0 2 ACSN39 0 3 70.0 30.0 2 ACSN39 0 4 37.5 62.5 2 45H29 1x106 1 50.0 50.0 2 45H29 1x106 2 50.0 50.0 2 45H29 1x106 3 30.0 70.0 2 45H29 1x106 4 22.2 77.8 2 45H29 0 1 30.0 70.0 2 45H29 0 2 30.0 70.0 2 45H29 0 3 77.8 22.2 2 45H29 0 4 40.0 60.0 2 7367 1x106 1 90.0 0.0 2 7367 1x106 2 90.0 10.0 2 7367 1x106 3 80.0 10.0 2 7367 1x106 4 30.0 70.0 2 7367 0 1 80.0 20.0 2 7367 0 2 90.0 0.0 2 7367 0 3 90.0 0.0
148 2 7367 0 4 40.0 60.0 2 7377 1x106 1 90.0 10.0 2 7377 1x106 2 80.0 0.0 2 7377 1x106 3 80.0 0.0 2 7377 1x106 4 100.0 0.0 2 7377 0 1 100.0 0.0 2 7377 0 2 88.9 11.1 2 7377 0 3 90.0 0.0 2 7377 0 4 66.7 0.0
Table A1.71. Raw data for controlled environment study – napa cabbage: leaf length
Repetition Cultivar Spores ml-1 Block
2 WAI
3 WAI
4 WAI
5 WAI
6 WAI
1 ChinaGold 1x106 1 2.49 2.68 2.75 3.28 3.7 1 ChinaGold 1x106 2 2.30 2.47 2.83 3.10 3.78 1 ChinaGold 1x106 3 2.83 2.82 2.83 3.29 3.77 1 ChinaGold 1x106 4 2.51 2.89 2.76 3.00 3.43 1 ChinaGold 0 1 2.65 2.93 2.92 3.70 3.97 1 ChinaGold 0 2 2.67 2.76 3.02 3.20 3.84 1 ChinaGold 0 3 2.78 2.91 2.78 3.48 3.94 1 ChinaGold 0 4 2.43 2.54 2.95 3.27 3.50 1 Emiko 1x106 1 2.33 2.40 2.62 3.03 3.39 1 Emiko 1x106 2 2.53 2.56 2.91 2.73 3.02 1 Emiko 1x106 3 2.41 2.82 2.82 3.22 3.26 1 Emiko 1x106 4 2.36 2.63 2.80 2.90 3.13 1 Emiko 0 1 2.40 2.50 2.85 3.49 3.77 1 Emiko 0 2 2.39 2.51 2.68 2.51 2.83 1 Emiko 0 3 2.23 2.63 2.91 3.26 3.43 1 Emiko 0 4 2.44 3.54 2.70 2.99 3.05 1 Mirako 1x106 1 2.77 2.89 2.92 3.65 4.39 1 Mirako 1x106 2 2.80 2.91 3.16 3.38 3.77 1 Mirako 1x106 3 2.63 2.98 3.14 3.44 3.65 1 Mirako 1x106 4 2.53 2.94 2.97 3.27 3.51 1 Mirako 0 1 2.79 2.85 3.15 3.59 3.96 1 Mirako 0 2 2.72 2.93 3.21 3.41 3.81 1 Mirako 0 3 2.70 3.14 3.37 3.58 3.95 1 Mirako 0 4 2.71 3.17 3.13 3.71 3.78 1 Yuki 1x106 1 2.06 2.09 2.60 2.73 3.57 1 Yuki 1x106 2 2.08 2.41 2.68 3.03 3.53 1 Yuki 1x106 3 1.90 2.33 2.73 3.04 3.40 1 Yuki 1x106 4 2.30 2.67 2.79 2.99 3.27 1 Yuki 0 1 2.05 2.36 2.48 3.07 3.37 1 Yuki 0 2 2.36 2.65 2.97 3.32 3.71
149 1 Yuki 0 3 2.12 2.62 2.88 2.15 3.19 1 Yuki 0 4 2.13 2.70 2.83 3.18 3.42 2 ChinaGold 1x106 1 5.25 5.55 7.45 7.75 7.76 2 ChinaGold 1x106 2 5.16 5.30 6.01 6.92 6.01 2 ChinaGold 1x106 3 4.43 4.60 6.64 7.48 7.18 2 ChinaGold 1x106 4 4.53 5.26 6.06 5.96 5.75 2 ChinaGold 0 1 5.80 5.85 8.05 10.23 9.05 2 ChinaGold 0 2 4.94 5.74 6.82 7.71 7.37 2 ChinaGold 0 3 4.66 4.92 6.99 7.22 6.12 2 ChinaGold 0 4 4.17 4.26 6.43 7.52 6.55 2 Emiko 1x106 1 4.40 4.76 5.76 6.59 6.61 2 Emiko 1x106 2 4.03 4.31 5.73 6.09 6.08 2 Emiko 1x106 3 4.53 4.76 6.46 6.07 6.69 2 Emiko 1x106 4 3.61 3.87 4.64 5.07 5.18 2 Emiko 0 1 4.92 5.01 5.95 11.22 6.83 2 Emiko 0 2 4.22 4.66 5.73 6.97 6.49 2 Emiko 0 3 4.55 3.63 4.85 6.81 6.20 2 Emiko 0 4 3.34 3.63 5.08 5.78 5.81 2 Mirako 1x106 1 4.22 4.54 5.89 7.22 6.97 2 Mirako 1x106 2 4.77 4.84 5.93 6.58 5.70 2 Mirako 1x106 3 4.04 4.09 6.35 7.23 7.42 2 Mirako 1x106 4 4.10 4.09 5.35 6.23 5.70 2 Mirako 0 1 5.47 5.53 6.31 7.28 7.13 2 Mirako 0 2 4.94 5.20 7.17 7.69 7.13 2 Mirako 0 3 4.89 4.71 6.50 6.59 6.88 2 Mirako 0 4 5.02 4.71 5.56 6.46 6.37 2 Yuki 1x106 1 3.13 4.71 6.64 7.02 5.90 2 Yuki 1x106 2 3.80 4.03 10.31 6.23 5.90 2 Yuki 1x106 3 3.49 5.26 6.03 6.01 5.38 2 Yuki 1x106 4 3.45 4.22 5.16 5.19 5.05 2 Yuki 0 1 3.97 5.07 6.70 6.92 6.45 2 Yuki 0 2 4.55 5.21 7.00 7.06 6.45 2 Yuki 0 3 4.03 3.64 6.12 5.74 4.73 2 Yuki 0 4 3.93 4.33 6.30 6.37 5.70
Table A1.72. Raw data for controlled environment study – napa cabbage: biomass (dry shoot weight)
Repetition Cultivar Spores g-1 soil Block
Mean dry shoot weight (g)
1 ChinaGold 1x106 1 4.54 1 ChinaGold 1x106 2 2.88 1 ChinaGold 1x106 3 3.94 1 ChinaGold 1x106 4 2.49
150 1 ChinaGold 0 1 4.10 1 ChinaGold 0 2 2.90 1 ChinaGold 0 3 2.99 1 ChinaGold 0 4 3.10 1 Emiko 1x106 1 3.31 1 Emiko 1x106 2 4.11 1 Emiko 1x106 3 2.31 1 Emiko 1x106 4 1.99 1 Emiko 0 1 5.39 1 Emiko 0 2 3.32 1 Emiko 0 3 2.96 1 Emiko 0 4 2.16 1 Mirako 1x106 1 5.17 1 Mirako 1x106 2 4.46 1 Mirako 1x106 3 3.04 1 Mirako 1x106 4 2.67 1 Mirako 0 1 5.19 1 Mirako 0 2 5.42 1 Mirako 0 3 3.79 1 Mirako 0 4 3.08 1 Yuki 1x106 1 5.51 1 Yuki 1x106 2 5.05 1 Yuki 1x106 3 4.81 1 Yuki 1x106 4 4.47 1 Yuki 0 1 5.40 1 Yuki 0 2 4.08 1 Yuki 0 3 3.93 1 Yuki 0 4 3.96 2 ChinaGold 1x106 1 12.26 2 ChinaGold 1x106 2 10.13 2 ChinaGold 1x106 3 10.79 2 ChinaGold 1x106 4 9.17 2 ChinaGold 0 1 11.29 2 ChinaGold 0 2 11.53 2 ChinaGold 0 3 9.09 2 ChinaGold 0 4 10.83 2 Emiko 1x106 1 19.42 2 Emiko 1x106 2 17.18 2 Emiko 1x106 3 20.38 2 Emiko 1x106 4 11.35 2 Emiko 0 1 32.33 2 Emiko 0 2 17.38 2 Emiko 0 3 18.28
151 2 Emiko 0 4 17.57 2 Mirako 1x106 1 10.17 2 Mirako 1x106 2 9.35 2 Mirako 1x106 3 9.97 2 Mirako 1x106 4 9.64 2 Mirako 0 1 14.21 2 Mirako 0 2 18.79 2 Mirako 0 3 11.69 2 Mirako 0 4 13.36 2 Yuki 1x106 1 24.60 2 Yuki 1x106 2 18.72 2 Yuki 1x106 3 17.78 2 Yuki 1x106 4 14.93 2 Yuki 0 1 17.79 2 Yuki 0 2 17.70 2 Yuki 0 3 27.95 2 Yuki 0 4 27.45
152 Table A1.73. Raw data for field trials – cabbage: CI & DSI
Repetition Spores g-1 soil Cultivar Block CI DSI
1 1x106 Bronco 1 37.5 12.5 1 1x106 Bronco 2 62.5 29.2 1 1x106 Bronco 3 57.1 19.0 1 0 Bronco 1 0.0 0.0 1 0 Bronco 2 0.0 0.0 1 0 Bronco 3 0.0 0.0 1 1x106 Kilaxy 1 0.0 0.0 1 1x106 Kilaxy 2 0.0 0.0 1 1x106 Kilaxy 3 0.0 0.0 1 0 Kilaxy 1 0.0 0.0 1 0 Kilaxy 2 0.0 0.0 1 0 Kilaxy 3 0.0 0.0 1 1x106 Kilaton 1 0.0 0.0 1 1x106 Kilaton 2 0.0 0.0 1 1x106 Kilaton 3 0.0 0.0 1 0 Kilaton 1 0.0 0.0 1 0 Kilaton 2 0.0 0.0 1 0 Kilaton 3 0.0 0.0 1 1x106 Tekila 1 0.0 0.0 1 1x106 Tekila 2 0.0 0.0 1 1x106 Tekila 3 0.0 0.0 1 0 Tekila 1 0.0 0.0 1 0 Tekila 2 0.0 0.0 1 0 Tekila 3 0.0 0.0 2 1x106 Bronco 1 40.0 20.0 2 1x106 Bronco 2 10.0 3.3 2 1x106 Bronco 3 11.1 3.7 2 1x106 Bronco 4 30.0 13.3 2 0 Bronco 1 0.0 0.0 2 0 Bronco 2 0.0 0.0 2 0 Bronco 3 0.0 0.0 2 0 Bronco 4 0.0 0.0 2 1x106 Kilaherb 1 0.0 0.0 2 1x106 Kilaherb 2 0.0 0.0 2 1x106 Kilaherb 3 0.0 0.0 2 1x106 Kilaherb 4 0.0 0.0 2 0 Kilaherb 1 0.0 0.0 2 0 Kilaherb 2 0.0 0.0 2 0 Kilaherb 3 0.0 0.0 2 0 Kilaherb 4 0.0 0.0
153 2 1x106 Kilaton 1 0.0 0.0 2 1x106 Kilaton 2 0.0 0.0 2 1x106 Kilaton 3 0.0 0.0 2 1x106 Kilaton 4 0.0 0.0 2 0 Kilaton 1 0.0 0.0 2 0 Kilaton 2 0.0 0.0 2 0 Kilaton 3 0.0 0.0 2 0 Kilaton 4 0.0 0.0 2 1x106 Tekila 1 0.0 0.0 2 1x106 Tekila 2 0.0 0.0 2 1x106 Tekila 3 0.0 0.0 2 1x106 Tekila 4 0.0 0.0 2 0 Tekila 1 0.0 0.0 2 0 Tekila 2 0.0 0.0 2 0 Tekila 3 0.0 0.0 2 0 Tekila 4 0.0 0.0
Table A1.74. Raw data for controlled environment study – cabbage: leaf length
Run Cultivar Spores ml-1 Block 2 WAI 3 WAI 4 WAI 5 WAI 6 WAI
1 Bronco 1x106 1 2.82 3.09 3.30 3.43 3.49 1 Bronco 1x106 2 2.93 2.79 . 3.31 3.56 1 Bronco 1x106 3 2.98 2.82 3.23 3.31 3.34 1 Bronco 0 1 2.65 2.67 2.96 3.18 3.32 1 Bronco 0 2 2.98 2.97 3.21 3.47 3.63 1 Bronco 0 3 3.18 3.13 3.17 3.43 3.77 1 Kilaxy 1x106 1 2.47 2.65 3.13 3.65 3.04 1 Kilaxy 1x106 2 2.79 2.80 3.09 3.88 4.02 1 Kilaxy 1x106 3 2.94 2.93 2.91 3.10 3.36 1 Kilaxy 0 1 2.78 2.92 3.23 3.72 3.71 1 Kilaxy 0 2 2.76 3.00 . 3.50 3.57 1 Kilaxy 0 3 2.81 2.91 3.19 3.22 3.52 1 Kilaton 1x106 1 1.93 1.09 1.24 3.16 3.35 1 Kilaton 1x106 2 1.98 2.10 . 2.81 3.26 1 Kilaton 1x106 3 2.26 2.55 2.81 3.06 3.19 1 Kilaton 0 1 1.82 1.57 1.50 2.98 3.38 1 Kilaton 0 2 1.98 2.17 2.55 2.76 3.33 1 Kilaton 0 3 2.07 2.33 2.52 2.85 3.01 1 Tekila 1x106 1 2.50 2.57 2.84 3.11 3.13 1 Tekila 1x106 2 2.43 2.56 . 3.03 3.15 1 Tekila 1x106 3 2.62 2.90 3.09 3.62 3.70 1 Tekila 0 1 2.61 2.55 2.76 3.26 3.46
154 1 Tekila 0 2 2.63 2.84 3.84 3.13 3.17 1 Tekila 0 3 2.74 2.74 3.03 3.07 3.2 2 Bronco 1x106 1 4.37 4.55 5.06 5.49 5.68 2 Bronco 1x106 2 5.20 5.26 5.40 5.50 5.43 2 Bronco 1x106 3 4.72 4.37 4.35 4.16 4.26 2 Bronco 1x106 4 5.91 6.47 6.61 6.41 6.87 2 Bronco 0 1 6.48 6.35 6.74 7.04 7.24 2 Bronco 0 2 6.15 6.16 6.53 6.50 6.57 2 Bronco 0 3 6.58 6.82 7.19 7.24 6.92 2 Bronco 0 4 4.39 4.45 4.47 4.46 4.40 2 Kilaherb 1x106 1 4.93 5.76 6.55 7.07 7.03 2 Kilaherb 1x106 2 4.43 5.48 6.04 6.20 5.69 2 Kilaherb 1x106 3 4.54 5.13 5.49 5.42 5.06 2 Kilaherb 1x106 4 5.30 7.01 7.84 8.39 7.54 2 Kilaherb 0 1 5.28 6.69 7.02 7.54 7.77 2 Kilaherb 0 2 6.17 7.15 7.46 7.70 7.35 2 Kilaherb 0 3 5.00 6.48 7.02 7.62 7.48 2 Kilaherb 0 4 5.31 6.27 7.13 7.52 7.16 2 Kilaton 1x106 1 4.45 5.15 4.85 5.98 5.72 2 Kilaton 1x106 2 4.23 4.31 4.47 4.70 4.96 2 Kilaton 1x106 3 3.50 4.18 4.38 4.50 4.63 2 Kilaton 1x106 4 . 5.40 6.50 5.90 5.55 2 Kilaton 0 1 4.21 4.64 5.17 5.36 5.26 2 Kilaton 0 2 3.85 5.37 6.57 6.03 5.54 2 Kilaton 0 3 3.68 4.30 4.45 4.67 4.47 2 Kilaton 0 4 4.47 5.73 6.68 6.90 6.75 2 Tekila 1x106 1 4.84 5.37 5.67 6.04 5.86 2 Tekila 1x106 2 5.52 5.58 5.85 5.74 5.60 2 Tekila 1x106 3 5.55 5.74 6.11 6.16 5.85 2 Tekila 1x106 4 5.21 5.39 5.98 5.63 5.69 2 Tekila 0 1 4.34 4.89 5.43 5.21 5.41 2 Tekila 0 2 4.12 4.53 4.73 4.92 5.11 2 Tekila 0 3 5.43 5.74 6.16 6.14 5.92 2 Tekila 0 4 6.19 6.14 6.51 6.30 6.19
Table A1.75. Raw data for controlled environment study – cabbage: biomass (dry shoot weight of 10 plants, g)
Repetition Cultivar Spores ml-1 Block
Dry shoot weight (g)
1 Bronco 1x106 1 8.73 1 Bronco 1x106 2 8.16 1 Bronco 1x106 3 6.91 1 Bronco 0 1 6.88 1 Bronco 0 2 6.96 1 Bronco 0 3 6.74
155 1 Kilaxy 1x106 1 5.55 1 Kilaxy 1x106 2 4.80 1 Kilaxy 1x106 3 6.13 1 Kilaxy 0 1 5.80 1 Kilaxy 0 2 4.05 1 Kilaxy 0 3 5.12 1 Kilaton 1x106 1 7.70 1 Kilaton 1x106 2 7.03 1 Kilaton 1x106 3 6.20 1 Kilaton 0 1 7.90 1 Kilaton 0 2 8.04 1 Kilaton 0 3 7.57 1 Tekila 1x106 1 7.57 1 Tekila 1x106 2 8.14 1 Tekila 1x106 3 8.50 1 Tekila 0 1 8.09 1 Tekila 0 2 9.58 1 Tekila 0 3 8.37 2 Bronco 1x106 1 6.45 2 Bronco 1x106 2 10.39 2 Bronco 1x106 3 7.24 2 Bronco 1x106 4 27.94 2 Bronco 0 1 7.94 2 Bronco 0 2 9.59 2 Bronco 0 3 3.97 2 Bronco 0 4 16.78 2 Kilaherb 1x106 1 13.41 2 Kilaherb 1x106 2 13.96 2 Kilaherb 1x106 3 30.99 2 Kilaherb 1x106 4 9.18 2 Kilaherb 0 1 11.39 2 Kilaherb 0 2 16.48 2 Kilaherb 0 3 13.54 2 Kilaherb 0 4 28.68 2 Kilaton 1x106 1 7.10 2 Kilaton 1x106 2 10.93 2 Kilaton 1x106 3 5.71 2 Kilaton 1x106 4 40.00 2 Kilaton 0 1 9.25 2 Kilaton 0 2 10.79 2 Kilaton 0 3 16.42 2 Kilaton 0 4 26.53 2 Tekila 1x106 1 8.51 2 Tekila 1x106 2 9.48 2 Tekila 1x106 3 5.49 2 Tekila 1x106 4 25.10
156 2 Tekila 0 1 8.48 2 Tekila 0 2 6.29 2 Tekila 0 3 18.83 2 Tekila 0 4 30.01
Table A1.76. Raw data for field trials – canola: CI & DSI
Year Site ID Cultivar Block CI DSI
2014 Low ACSN39 1 100.0 100.0 2014 Low ACSN39 2 100.0 100.0 2014 Low ACSN39 3 100.0 100.0 2014 Low ACSN39 4 100.0 100.0 2014 Low 45H29 1 0.0 0.0 2014 Low 45H29 2 0.0 0.0 2014 Low 45H29 3 0.0 0.0 2014 Low 45H29 4 0.0 0.0 2014 Low 7377 1 0.0 0.0 2014 Low 7377 2 0.0 0.0 2014 Low 7377 3 0.0 0.0 2014 Low 7377 4 0.0 0.0 2014 Low 7367 1 0.0 0.0 2014 Low 7367 2 0.0 0.0 2014 Low 7367 3 0.0 0.0 2014 Low 7367 4 0.0 0.0 2014 High ACSN39 1 100.0 100.0 2014 High ACSN39 2 100.0 100.0 2014 High ACSN39 3 100.0 100.0 2014 High ACSN39 4 100.0 100.0 2014 High 45H29 1 0.0 0.0 2014 High 45H29 2 0.0 0.0 2014 High 45H29 3 0.0 0.0 2014 High 45H29 4 0.0 0.0 2014 High 7377 1 0.0 0.0 2014 High 7377 2 0.0 0.0 2014 High 7377 3 0.0 0.0 2014 High 7377 4 0.0 0.0 2014 High 7367 1 0.0 0.0 2014 High 7367 2 0.0 0.0 2014 High 7367 3 0.0 0.0 2014 High 7367 4 0.0 0.0 2015 BDL ACSN39 1 30.0 16.7
157 2015 BDL ACSN39 2 100.0 43.3 2015 BDL ACSN39 3 20.0 6.7 2015 BDL ACSN39 4 30.0 10.0 2015 BDL 45H29 1 0.0 0.0 2015 BDL 45H29 2 0.0 0.0 2015 BDL 45H29 3 10.0 3.3 2015 BDL 45H29 4 0.0 0.0 2015 BDL 7377 1 0.0 0.0 2015 BDL 7377 2 0.0 0.0 2015 BDL 7377 3 10.0 3.3 2015 BDL 7377 4 10.0 3.3 2015 BDL 7367 1 0.0 0.0 2015 BDL 7367 2 0.0 0.0 2015 BDL 7367 3 0.0 0.0 2015 BDL 7367 4 0.0 0.0 2015 Low ACSN39 1 100.0 63.3 2015 Low ACSN39 2 100.0 76.7 2015 Low ACSN39 3 100.0 56.7 2015 Low ACSN39 4 100.0 63.3 2015 Low 45H29 1 0.0 0.0 2015 Low 45H29 2 0.0 0.0 2015 Low 45H29 3 10.0 3.3 2015 Low 45H29 4 0.0 0.0 2015 Low 7377 1 0.0 0.0 2015 Low 7377 2 10.0 3.3 2015 Low 7377 3 0.0 0.0 2015 Low 7377 4 0.0 0.0 2015 Low 7367 1 0.0 0.0 2015 Low 7367 2 10.0 3.3 2015 Low 7367 3 10.0 3.3 2015 Low 7367 4 0.0 0.0 2015 High ACSN39 1 100.0 86.7 2015 High ACSN39 2 100.0 63.3 2015 High ACSN39 3 100.0 66.7 2015 High ACSN39 4 100.0 63.3 2015 High 45H29 1 0.0 0.0 2015 High 45H29 2 10.0 6.7 2015 High 45H29 3 0.0 0.0 2015 High 45H29 4 0.0 0.0 2015 High 7377 1 20.0 6.7 2015 High 7377 2 0.0 0.0 2015 High 7377 3 0.0 0.0 2015 High 7377 4 0.0 0.0
158 2015 High 7367 1 0.0 0.0 2015 High 7367 2 0.0 0.0 2015 High 7367 3 40.0 26.7 2015 High 7367 4 0.0 0.0
Table A1.77. Raw data for field trials – canola: plant height
Year Site ID Cultivar Block 4 WAS 5 WAS 6 WAS 7 WAS 8 WAS 9 WAS
2014 Low ACSN39 1 3.11 7.52 12.52 22.14 27.95 2014 Low ACSN39 2 2.98 6.80 13.97 22.63 23.06 2014 Low ACSN39 3 2.85 6.15 14.72 22.24 26.94 2014 Low ACSN39 4 3.52 11.00 24.33 32.20 37.93 2014 Low 45H29 1 2.54 7.14 26.07 78.63 73.04 2014 Low 45H29 2 2.72 8.11 33.96 65.73 76.30 2014 Low 45H29 3 2.58 5.18 15.83 46.38 68.96 2014 Low 45H29 4 2.79 6.25 27.78 63.99 72.73 2014 Low 7377 1 2.72 5.08 15.97 44.50 64.51 2014 Low 7377 2 2.76 5.00 12.90 29.88 45.38 2014 Low 7377 3 2.53 5.83 17.73 35.97 51.60 2014 Low 7377 4 2.58 6.70 22.03 31.22 51.50 2014 Low 7367 1 3.97 6.91 19.96 61.82 71.44 2014 Low 7367 2 3.82 8.48 32.25 55.22 72.77 2014 Low 7367 3 3.18 5.56 17.63 38.88 62.50 2014 Low 7367 4 2.95 6.97 19.55 49.74 66.60 2014 High ACSN39 1 2.99 4.25 8.66 21.71 39.13 2014 High ACSN39 2 1.92 4.27 9.81 26.85 34.49 2014 High ACSN39 3 3.11 5.17 10.45 19.55 18.89 2014 High ACSN39 4 2.82 6.05 11.37 23.16 30.83 2014 High 45H29 1 2.36 3.40 9.22 25.08 36.23 2014 High 45H29 2 2.36 3.98 7.91 20.09 39.65 2014 High 45H29 3 2.68 4.25 7.23 22.99 43.67 2014 High 45H29 4 2.69 4.42 10.89 32.50 47.41 2014 High 7377 1 3.10 9.72 8.25 20.16 41.45 2014 High 7377 2 2.73 4.71 9.78 27.20 46.03 2014 High 7377 3 2.83 4.19 8.63 29.71 54.82 2014 High 7377 4 3.19 5.06 12.63 33.16 54.49 2014 High 7367 1 3.08 4.77 9.03 19.62 34.06 2014 High 7367 2 3.98 6.15 9.64 29.60 44.82 2014 High 7367 3 4.23 5.21 10.07 26.98 42.57 2014 High 7367 4 3.83 5.11 7.89 27.65 55.67 2015 BDL ACSN39 1 15.22 29.70 55.22 83.58 103.02 112.51
159 2015 BDL ACSN39 2 10.69 27.07 63.55 87.98 110.60 112.20 2015 BDL ACSN39 3 13.91 25.51 59.49 91.96 101.82 101.77 2015 BDL ACSN39 4 9.90 19.42 43.60 75.18 98.74 106.47 2015 BDL 45H29 1 6.88 18.03 52.91 78.34 103.32 111.35 2015 BDL 45H29 2 7.33 18.20 63.59 97.28 110.69 113.65 2015 BDL 45H29 3 7.53 19.03 53.92 90.42 105.38 97.98 2015 BDL 45H29 4 5.50 12.14 55.67 80.91 100.50 107.53 2015 BDL 7377 1 4.48 9.74 44.12 68.50 104.28 96.44 2015 BDL 7377 2 4.09 8.04 35.03 64.44 110.95 110.99 2015 BDL 7377 3 5.15 9.40 40.16 72.52 99.40 109.24 2015 BDL 7377 4 4.43 8.69 45.23 82.48 100.34 131.24 2015 BDL 7367 1 7.61 14.30 40.21 59.98 95.65 94.59 2015 BDL 7367 2 8.00 15.53 40.45 62.48 98.47 106.73 2015 BDL 7367 3 7.07 12.84 53.59 64.98 103.84 105.68 2015 BDL 7367 4 7.25 11.13 41.78 72.00 95.82 113.83 2015 Low ACSN39 1 17.67 32.22 66.70 83.05 89.75 95.96 2015 Low ACSN39 2 13.85 28.29 56.85 99.12 101.01 110.28 2015 Low ACSN39 3 14.01 24.65 56.12 83.89 102.24 103.57 2015 Low ACSN39 4 13.14 22.68 49.73 83.23 103.24 98.96 2015 Low 45H29 1 7.70 16.67 52.90 75.41 94.12 95.99 2015 Low 45H29 2 6.43 15.32 47.75 65.91 97.96 105.84
160 2015 Low 45H29 3 6.49 18.14 65.70 92.81 111.51 103.20 2015 Low 45H29 4 4.81 13.06 50.89 77.69 99.00 107.44 2015 Low 7377 1 6.07 13.47 47.13 68.82 105.83 99.92 2015 Low 7377 2 5.65 12.98 54.49 86.51 104.06 105.15 2015 Low 7377 3 5.73 13.48 52.83 81.21 97.51 106.30 2015 Low 7377 4 5.39 10.81 44.31 67.40 88.25 100.54 2015 Low 7367 1 12.55 23.07 63.84 81.87 78.06 88.76 2015 Low 7367 2 8.03 14.00 49.36 84.90 102.67 99.37 2015 Low 7367 3 6.72 14.80 47.20 73.20 99.74 107.56 2015 Low 7367 4 7.25 14.79 43.76 67.00 87.29 107.90 2015 High ACSN39 1 9.48 21.05 66.74 86.51 96.05 99.19 2015 High ACSN39 2 13.20 25.44 61.17 81.22 99.41 104.79 2015 High ACSN39 3 16.29 28.90 63.53 71.43 81.10 105.76 2015 High ACSN39 4 14.21 26.98 72.82 92.97 100.07 102.55 2015 High 45H29 1 6.50 15.80 62.23 82.51 100.23 97.63 2015 High 45H29 2 5.34 14.88 57.67 83.06 103.83 102.11 2015 High 45H29 3 6.68 13.70 58.05 89.55 109.35 105.86 2015 High 45H29 4 6.55 14.16 57.32 83.52 105.01 104.62 2015 High 7377 1 6.31 8.44 31.40 51.35 82.05 84.43 2015 High 7377 2 5.61 11.64 39.66 59.14 92.21 97.31 2015 High 7377 3 5.40 10.37 33.18 63.41 100.50 97.83 2015 High 7377 4 9.31 15.29 58.80 64.63 105.74 85.39 2015 High 7367 1 10.47 13.72 44.50 60.68 88.93 83.80 2015 High 7367 2 10.80 16.73 61.58 71.67 95.69 91.59 2015 High 7367 3 12.04 21.86 69.82 75.78 82.06 98.41 2015 High 7367 4 6.94 8.34 30.48 53.96 82.39 75.87
WAS = Weeks after seeding 2014 Low = 7 x 105 spores g-1 soil
2014 High = 7 x 106 spores g-1 soil
2015 BDL = Below detection limit (<1000 spores g-1 soil) 2015 Low = 3 x 106 spores g-1 soil 2015 High = 1 x 107 spores g-1 soil
Table A1.78. Raw data for field trials – canola: biomass (dry shoot weight)
Year Site ID Cultivar Block Dry shoot weight (g)
2014 Low ACSN39 1 15.50 2014 Low ACSN39 2 22.60 2014 Low ACSN39 3 17.80 2014 Low ACSN39 4 374.57 2014 Low 7377 1 889.82 2014 Low 7377 2 581.00 2014 Low 7377 3 238.29
161 2014 Low 7377 4 482.20 2014 Low 7367 1 362.70 2014 Low 7367 2 680.40 2014 Low 7367 3 335.83 2014 Low 7367 4 451.20 2014 Low 45H29 1 304.60 2014 Low 45H29 2 501.80 2014 Low 45H29 3 885.43 2014 Low 45H29 4 443.80 2014 High ACSN39 1 126.20 2014 High ACSN39 2 117.40 2014 High ACSN39 3 31.40 2014 High ACSN39 4 112.50 2014 High 7377 1 372.60 2014 High 7377 2 601.50 2014 High 7377 3 178.70 2014 High 7377 4 122.75 2014 High 7367 1 302.80 2014 High 7367 2 281.70 2014 High 7367 3 330.20 2014 High 7367 4 183.70 2014 High 45H29 1 247.10 2014 High 45H29 2 161.30 2014 High 45H29 3 285.40 2014 High 45H29 4 418.90 2015 BDL ACSN39 1 422.40 2015 BDL ACSN39 2 733.20 2015 BDL ACSN39 3 291.40 2015 BDL ACSN39 4 647.20 2015 BDL 45H29 1 398.80 2015 BDL 45H29 2 529.40 2015 BDL 45H29 3 394.20 2015 BDL 45H29 4 665.20 2015 BDL 7377 1 265.20 2015 BDL 7377 2 326.00 2015 BDL 7377 3 535.60 2015 BDL 7377 4 260.00 2015 BDL 7367 1 375.60 2015 BDL 7367 2 324.20 2015 BDL 7367 3 453.40 2015 BDL 7367 4 271.60 2015 Low ACSN39 1 241.40 2015 Low ACSN39 2 430.80
162 2015 Low ACSN39 3 219.60 2015 Low ACSN39 4 351.00 2015 Low 45H29 1 433.20 2015 Low 45H29 2 572.00 2015 Low 45H29 3 106.80 2015 Low 45H29 4 609.20 2015 Low 7377 1 362.40 2015 Low 7377 2 397.20 2015 Low 7377 3 407.00 2015 Low 7377 4 235.20 2015 Low 7367 1 207.00 2015 Low 7367 2 443.00 2015 Low 7367 3 512.80 2015 Low 7367 4 306.20 2015 High ACSN39 1 353.80 2015 High ACSN39 2 494.00 2015 High ACSN39 3 299.20 2015 High ACSN39 4 481.80 2015 High 45H29 1 379.40 2015 High 45H29 2 327.00 2015 High 45H29 3 223.40 2015 High 45H29 4 544.60 2015 High 7377 1 481.20 2015 High 7377 2 384.00 2015 High 7377 3 398.40 2015 High 7377 4 337.60 2015 High 7367 1 175.20 2015 High 7367 2 505.20 2015 High 7367 3 610.00 2015 High 7367 4 388.20
2014 Low = 7 x 105 spores g-1 soil
2014 High = 7 x 106 spores g-1 soil
2015 BDL = Below detection limit (<1000 spores g-1 soil) 2015 Low = 3 x 106 spores g-1 soil 2015 High = 1 x 107 spores g-1 soil
Table A1.79. Raw data for field trials – canola: maturity (% plants at flowering and seepod
development stage)
Year Site ID Cultivar Block
% Flowering
% Seedpod
2014 Low ACSN39 1 0 80 2014 Low ACSN39 2 20 20
163 2014 Low ACSN39 3 50 20 2014 Low ACSN39 4 30 60 2014 Low 7377 1 60 40 2014 Low 7377 2 40 60 2014 Low 7377 3 40 60 2014 Low 7377 4 20 80 2014 Low 7367 1 30 70 2014 Low 7367 2 40 60 2014 Low 7367 3 80 20 2014 Low 7367 4 50 40 2014 Low 45H29 1 20 80 2014 Low 45H29 2 30 70 2014 Low 45H29 3 40 60 2014 Low 45H29 4 20 80 2014 High ACSN39 1 80 10 2014 High ACSN39 2 60 0 2014 High ACSN39 3 20 20 2014 High ACSN39 4 50 10 2014 High 7377 1 90 10 2014 High 7377 2 50 30 2014 High 7377 3 80 10 2014 High 7377 4 100 0 2014 High 7367 1 60 0 2014 High 7367 2 90 10 2014 High 7367 3 80 0 2014 High 7367 4 90 10 2014 High 45H29 1 40 50 2014 High 45H29 2 70 30 2014 High 45H29 3 90 0 2014 High 45H29 4 60 30 2015 BDL ACSN39 1 100 100 2015 BDL ACSN39 2 100 100 2015 BDL ACSN39 3 100 100 2015 BDL ACSN39 4 100 100 2015 BDL 7377 1 100 100 2015 BDL 7377 2 100 100 2015 BDL 7377 3 100 100 2015 BDL 7377 4 100 100 2015 BDL 7367 1 100 100 2015 BDL 7367 2 100 100 2015 BDL 7367 3 100 100 2015 BDL 7367 4 100 100 2015 BDL 45H29 1 100 100
164 2015 BDL 45H29 2 100 100 2015 BDL 45H29 3 100 100 2015 BDL 45H29 4 100 100 2015 Low ACSN39 1 100 100 2015 Low ACSN39 2 100 100 2015 Low ACSN39 3 100 100 2015 Low ACSN39 4 100 100 2015 Low 7377 1 100 100 2015 Low 7377 2 100 100 2015 Low 7377 3 100 100 2015 Low 7377 4 100 100 2015 Low 7367 1 100 100 2015 Low 7367 2 100 100 2015 Low 7367 3 100 100 2015 Low 7367 4 100 100 2015 Low 45H29 1 100 100 2015 Low 45H29 2 100 100 2015 Low 45H29 3 100 100 2015 Low 45H29 4 100 100 2015 High ACSN39 1 100 100 2015 High ACSN39 2 100 100 2015 High ACSN39 3 100 100 2015 High ACSN39 4 100 100 2015 High 7377 1 100 100 2015 High 7377 2 100 100 2015 High 7377 3 100 100 2015 High 7377 4 100 100 2015 High 7367 1 100 100 2015 High 7367 2 100 100 2015 High 7367 3 100 100 2015 High 7367 4 100 100 2015 High 45H29 1 100 100 2015 High 45H29 2 100 100 2015 High 45H29 3 100 100 2015 High 45H29 4 100 100
165 2014 Low = 7 x 105 spores g-1 soil
2014 High = 7 x 106 spores g-1 soil
2015 BDL = Below detection limit (<1000 spores g-1 soil) 2015 Low = 3 x 106 spores g-1 soil 2015 High = 1 x 107 spores g-1 soil
Table A1.80. Raw data for field trials – napa cabbage: CI & DSI
Year Site ID Cultivar Block CI DSI
2014 Low Chinagold 1 0.0 0.0 2014 Low Chinagold 2 0.0 0.0 2014 Low Chinagold 3 0.0 0.0 2014 Low Chinagold 4 0.0 0.0 2014 Low Emiko 1 0.0 0.0 2014 Low Emiko 2 0.0 0.0 2014 Low Emiko 3 0.0 0.0 2014 Low Emiko 4 0.0 0.0 2014 Low Mirako 1 0.0 0.0 2014 Low Mirako 2 50.0 33.3 2014 Low Mirako 3 40.0 16.7 2014 Low Mirako 4 0.0 0.0 2014 Low Yuki 1 0.0 0.0 2014 Low Yuki 2 0.0 0.0 2014 Low Yuki 3 0.0 0.0 2014 Low Yuki 4 0.0 0.0 2014 High Chinagold 1 0.0 0.0 2014 High Chinagold 2 0.0 0.0 2014 High Chinagold 3 0.0 0.0 2014 High Chinagold 4 0.0 0.0 2014 High Emiko 1 0.0 0.0 2014 High Emiko 2 0.0 0.0 2014 High Emiko 3 0.0 0.0 2014 High Emiko 4 0.0 0.0 2014 High Mirako 1 100.0 58.3 2014 High Mirako 2 60.0 26.7 2014 High Mirako 3 100.0 44.4 2014 High Mirako 4 100.0 44.4 2014 High Yuki 1 0.0 0.0 2014 High Yuki 2 0.0 0.0 2014 High Yuki 3 0.0 0.0 2014 High Yuki 4 0.0 0.0 2015 BDL Chinagold 1 0.0 0.0 2015 BDL Chinagold 2 0.0 0.0
166 2015 BDL Chinagold 3 0.0 0.0 2015 BDL Chinagold 4 0.0 0.0 2015 BDL Emiko 1 0.0 0.0 2015 BDL Emiko 2 0.0 0.0 2015 BDL Emiko 3 0.0 0.0 2015 BDL Emiko 4 0.0 0.0 2015 BDL Suzuko 1 0.0 0.0 2015 BDL Suzuko 2 0.0 0.0 2015 BDL Suzuko 3 0.0 0.0 2015 BDL Suzuko 4 0.0 0.0 2015 BDL Yuki 1 0.0 0.0 2015 BDL Yuki 2 0.0 0.0 2015 BDL Yuki 3 0.0 0.0 2015 BDL Yuki 4 0.0 0.0 2015 Low Chinagold 1 0.0 0.0 2015 Low Chinagold 2 0.0 0.0 2015 Low Chinagold 3 0.0 0.0 2015 Low Chinagold 4 0.0 0.0 2015 Low Emiko 1 0.0 0.0 2015 Low Emiko 2 0.0 0.0 2015 Low Emiko 3 0.0 0.0 2015 Low Emiko 4 0.0 0.0 2015 Low Suzuko 1 0.0 0.0 2015 Low Suzuko 2 0.0 0.0 2015 Low Suzuko 3 0.0 0.0 2015 Low Suzuko 4 0.0 0.0 2015 Low Yuki 1 0.0 0.0 2015 Low Yuki 2 0.0 0.0 2015 Low Yuki 3 0.0 0.0 2015 Low Yuki 4 0.0 0.0 2015 High Chinagold 1 0.0 0.0 2015 High Chinagold 2 0.0 0.0 2015 High Chinagold 3 0.0 0.0 2015 High Chinagold 4 0.0 0.0 2015 High Emiko 1 0.0 0.0 2015 High Emiko 2 0.0 0.0 2015 High Emiko 3 0.0 0.0 2015 High Emiko 4 0.0 0.0 2015 High Suzuko 1 0.0 0.0 2015 High Suzuko 2 0.0 0.0 2015 High Suzuko 3 0.0 0.0 2015 High Suzuko 4 0.0 0.0 2015 High Yuki 1 0.0 0.0
167 2015 High Yuki 2 0.0 0.0 2015 High Yuki 3 0.0 0.0 2015 High Yuki 4 0.0 0.0
Table A1.81. Raw data for field trials – napa cabbage: leaf length
Year Site ID Cultivar Block 4 WAS 5 WAS 6 WAS 7 WAS 8 WAS
2014 Low ChinaGold 1 7.52 10.82 13.46 15.46 14.65 2014 Low ChinaGold 2 7.45 11.69 14.47 15.18 16.91 2014 Low ChinaGold 3 6.41 10.75 13.70 15.30 17.10 2014 Low ChinaGold 4 9.51 12.26 14.53 18.94 19.41 2014 Low Emiko 1 9.65 12.75 14.22 16.96 18.23 2014 Low Emiko 2 9.05 14.10 15.17 19.05 18.82 2014 Low Emiko 3 10.04 12.30 14.25 16.80 17.17 2014 Low Emiko 4 11.51 14.05 16.17 20.74 18.71 2014 Low Yuki 1 4.64 9.06 12.39 13.51 12.78 2014 Low Yuki 2 8.13 9.40 12.24 12.64 13.70 2014 Low Yuki 3 8.08 9.80 11.78 11.97 12.13 2014 Low Yuki 4 5.65 10.56 12.87 12.04 10.53 2014 Low Mirako 1 9.09 12.98 13.45 14.44 14.18 2014 Low Mirako 2 9.22 13.25 14.43 16.15 15.36 2014 Low Mirako 3 8.92 12.80 14.40 17.91 15.83 2014 Low Mirako 4 9.16 11.65 14.69 15.72 15.34 2014 High ChinaGold 1 8.63 12.43 14.61 12.88 11.44 2014 High ChinaGold 2 7.67 11.60 11.80 12.74 12.34 2014 High ChinaGold 3 6.27 11.29 12.21 11.41 10.21 2014 High ChinaGold 4 6.57 9.39 9.73 11.67 11.73 2014 High Emiko 1 7.16 10.70 13.80 12.89 9.88 2014 High Emiko 2 7.48 10.90 12.54 12.88 12.10 2014 High Emiko 3 8.27 11.32 12.73 12.38 11.93 2014 High Emiko 4 7.95 10.75 11.52 11.82 11.18 2014 High Yuki 1 4.21 8.13 10.56 10.39 10.25 2014 High Yuki 2 5.79 10.12 11.24 10.66 10.35 2014 High Yuki 3 2.88 6.77 9.90 10.46 9.40 2014 High Yuki 4 2.92 5.31 8.99 10.06 9.59 2014 High Mirako 1 5.67 9.21 11.46 12.25 10.73 2014 High Mirako 2 7.48 10.65 12.82 11.04 11.84 2014 High Mirako 3 6.71 10.03 11.80 12.82 10.84 2014 High Mirako 4 7.42 11.70 11.80 12.38 12.07 2015 BDL Chinagold 1 9.45 7.18 29.55 32.95 28.15 2015 BDL Chinagold 2 10.26 9.84 24.60 30.40 27.55 2015 BDL Chinagold 3 10.57 9.26 23.40 27.57 27.40
168 2015 BDL Chinagold 4 9.05 9.48 21.56 22.92 21.83 2015 BDL Emiko 1 9.86 8.86 23.57 28.11 28.00 2015 BDL Emiko 2 8.73 7.23 25.06 30.59 30.45 2015 BDL Emiko 3 9.26 8.75 24.65 27.92 28.32 2015 BDL Emiko 4 8.39 8.42 23.70 24.37 26.45 2015 BDL Suzuko 1 9.92 8.77 30.91 33.61 25.47 2015 BDL Suzuko 2 8.54 8.96 22.06 24.61 26.33 2015 BDL Suzuko 3 8.44 7.20 22.75 22.04 28.69 2015 BDL Suzuko 4 7.87 7.54 18.13 17.63 22.43 2015 BDL Yuki 1 8.17 7.51 21.88 26.01 24.09 2015 BDL Yuki 2 9.25 7.69 23.51 24.80 22.22 2015 BDL Yuki 3 8.88 6.29 24.27 21.79 23.83 2015 BDL Yuki 4 9.48 6.63 23.01 19.43 20.61 2015 Low Chinagold 1 9.92 7.22 19.84 25.42 27.40 2015 Low Chinagold 2 10.02 7.21 17.44 25.55 25.89 2015 Low Chinagold 3 9.49 6.96 6.65 25.03 26.34 2015 Low Chinagold 4 8.28 6.91 19.01 23.07 27.37 2015 Low Emiko 1 8.41 7.60 20.31 28.91 29.57 2015 Low Emiko 2 8.96 6.73 18.92 25.49 29.98 2015 Low Emiko 3 8.13 6.48 19.09 23.54 25.08 2015 Low Emiko 4 8.66 6.73 15.66 20.52 24.14 2015 Low Suzuko 1 10.73 8.50 19.22 23.97 29.48 2015 Low Suzuko 2 9.47 8.49 14.75 25.37 25.75 2015 Low Suzuko 3 9.25 8.20 16.44 20.55 26.02 2015 Low Suzuko 4 8.75 8.26 13.10 20.11 26.75 2015 Low Yuki 1 9.30 6.03 14.49 25.31 26.32 2015 Low Yuki 2 8.39 6.05 16.31 23.67 26.64 2015 Low Yuki 3 8.28 5.89 6.59 20.65 24.86 2015 Low Yuki 4 7.20 6.58 16.21 19.85 26.08 2015 High Chinagold 1 9.54 9.58 19.08 20.05 19.56 2015 High Chinagold 2 9.30 10.14 18.40 24.06 19.95 2015 High Chinagold 3 10.87 9.11 18.38 21.02 20.65 2015 High Chinagold 4 8.37 8.84 16.16 22.49 22.65 2015 High Emiko 1 8.41 8.23 18.38 21.45 25.05 2015 High Emiko 2 9.46 8.16 18.02 21.61 21.58 2015 High Emiko 3 11.46 7.06 18.69 22.99 18.99 2015 High Emiko 4 10.35 7.02 19.86 22.93 20.00 2015 High Suzuko 1 10.25 10.15 17.59 16.82 19.84 2015 High Suzuko 2 10.69 8.89 14.57 19.00 22.26 2015 High Suzuko 3 9.89 9.26 17.96 20.58 20.01 2015 High Suzuko 4 10.48 8.86 16.86 20.52 20.63 2015 High Yuki 1 8.18 7.41 14.84 13.93 16.57 2015 High Yuki 2 9.61 6.82 17.14 19.08 18.39
169 2015 High Yuki 3 8.91 5.98 15.52 18.53 17.21 2015 High Yuki 4 8.81 6.17 17.05 19.74 18.85
2014 Low = 7 x 105 spores g-1 soil
2014 High = 7 x 106 spores g-1 soil
2015 BDL = Below detection limit (<1000 spores g-1 soil) 2015 Low = 3 x 106 spores g-1 soil 2015 High = 1 x 107 spores g-1 soil
Table A1.82. Raw data for field trials – napa cabbage: biomass (dry shoot weight)
Year Site ID Cultivar Block
Dry shoot weight (g)
2014 Low ChinaGold 1 664.11 2014 Low ChinaGold 2 857.90 2014 Low ChinaGold 3 900.70 2014 Low ChinaGold 4 861.50 2014 Low Emiko 1 826.30 2014 Low Emiko 2 750.40 2014 Low Emiko 3 917.33 2014 Low Emiko 4 1276.00 2014 Low Mirako 1 460.71 2014 Low Mirako 2 1045.63 2014 Low Mirako 3 1207.30 2014 Low Mirako 4 1175.67 2014 Low Yuki 1 191.63 2014 Low Yuki 2 771.00 2014 Low Yuki 3 614.56 2014 Low Yuki 4 979.86 2014 High ChinaGold 1 685.50 2014 High ChinaGold 2 912.00 2014 High ChinaGold 3 922.67 2014 High ChinaGold 4 609.63 2014 High Emiko 1 423.67 2014 High Emiko 2 437.50 2014 High Emiko 3 771.20 2014 High Emiko 4 247.50 2014 High Mirako 1 494.25 2014 High Mirako 2 995.75 2014 High Mirako 3 554.50 2014 High Mirako 4 1292.67 2014 High Yuki 1 124.56 2014 High Yuki 2 627.88 2014 High Yuki 3 278.00 2014 High Yuki 4 233.00
170 2015 BDL ChinaGold 1 1465.60 2015 BDL ChinaGold 2 2298.60 2015 BDL ChinaGold 3 885.60 2015 BDL ChinaGold 4 699.40 2015 BDL Emiko 1 720.00 2015 BDL Emiko 2 2117.80 2015 BDL Emiko 3 906.80 2015 BDL Emiko 4 2572.80 2015 BDL Suzuko 1 836.60 2015 BDL Suzuko 2 2066.40 2015 BDL Suzuko 3 652.20 2015 BDL Suzuko 4 1854.90 2015 BDL Yuki 1 650.50 2015 BDL Yuki 2 682.00 2015 BDL Yuki 3 1290.80 2015 BDL Yuki 4 442.20 2015 Low ChinaGold 1 741.60 2015 Low ChinaGold 2 1784.80 2015 Low ChinaGold 3 393.80 2015 Low ChinaGold 4 1022.80 2015 Low Emiko 1 208.40 2015 Low Emiko 2 541.60 2015 Low Emiko 3 1625.00 2015 Low Emiko 4 370.80 2015 Low Suzuko 1 499.00 2015 Low Suzuko 2 737.00 2015 Low Suzuko 3 1639.20 2015 Low Suzuko 4 1197.80 2015 Low Yuki 1 755.20 2015 Low Yuki 2 1132.40 2015 Low Yuki 3 2628.60 2015 Low Yuki 4 994.20 2015 High ChinaGold 1 356.00 2015 High ChinaGold 2 1900.80 2015 High ChinaGold 3 980.00 2015 High ChinaGold 4 976.60 2015 High Emiko 1 1280.90 2015 High Emiko 2 265.40 2015 High Emiko 3 867.80 2015 High Emiko 4 515.80 2015 High Suzuko 1 512.60 2015 High Suzuko 2 828.80 2015 High Suzuko 3 1703.80
171 2015 High Suzuko 4 658.60 2015 High Yuki 1 685.00 2015 High Yuki 2 761.40 2015 High Yuki 3 823.80 2015 High Yuki 4 441.80 2014 Low = 7 x 105 spores g-1 soil
2014 High = 7 x 106 spores g-1 soil
2015 BDL = Below detection limit (<1000 spores g-1 soil) 2015 Low = 3 x 106 spores g-1 soil 2015 High = 1 x 107 spores g-1 soil
172 APPENDIX 2: SUPPLEMENTARY TABLES FOR CHAPTER THREE
Table A2.1 Mean monthly air temperature and total monthly rainfall during the growing period of canola at Normandin, Quebec, from 2007-2014. Month Temperature (oC) Rainfall (mm)
LTA1 Actual LTA Actual 2007 May 9.7 10.0 49.5 12.6 June 15.9 14.8 71.2 52.4 July 17.1 16.3 102.0 90.4 Aug 15.9 14.0 63.2 97.8 Sept 13.1 11.2 82.5 95.9 2008 May 9.3 8.0 49.5 38.4 June 15.8 14.9 71.4 128.4 July 17.2 17.8 102.0 119.8 Aug 15.9 16.8 63.2 52.4 Sept 13.1 10.9 79.7 50.6 2009 May 8.7 7.0 52.3 70.6 June 14.7 15.0 68.4 39.0 July 17.2 16.3 97.3 105.2 Aug 16.0 16.2 63.6 106.8 Sept 12.8 11.5 74.1 49.2 2010 May 9.1 10.7 48.9 43.0 June 14.8 13.8 69.2 62.8 July 17.3 18.9 93.3 47.8 Aug 16.0 17.1 64.3 50.0 Sept 11.6 11.3 86.7 184.7 2011 May 8.8 9.3 55.2 96.3 June 14.8 15.1 68.7 64.9 July 17.4 18.1 90.8 94.8 Aug 16.0 17.0 81.1 202.4 Sept 11.7 12.4 79.4 53.7 2012 May 9.1 11.1 54.6 76.2 June 15.1 16.6 72.9 104.4 July 17.5 17.7 86.6 39.1 Aug 16.0 16.9 85.9 103.3 Sept 11.7 11.9 88.1 120.9 2013 May 9.2 10.6 62.0 99.3 June 14.9 13.7 78.2 94.9 July 17.6 18.0 85.2 50.7
173 Aug 16.0 16.6 83.9 20.9 Sept 11.5 11.6 93.0 132.0 2014 May 9.6 10.2 59.6 67.1 June 15.4 16.7 76.1 78.1 July 17.6 17.9 85.8 101.1 Aug 16.2 17.2 93.5 143.4 Sept 11.5 11.3 95.3 85.3
1Based on 10-year average
Table A2.2. Decline in resting spores, qPCR: Type I MS
Source df Sum of Squares Mean Square F value Pr>F Rotation 5 11.62 2.32 9.29 0.0008 Block(Rotation) 6 2.10 0.35 1.40 0.2913 Error 12 3.00 0.25 Total 23 16.72 2.92 Table A2.3. Decline in resting spores, qPCR: Type III MS block(rotation) as Error Term
Source df Sum of Squares Mean Square F value Pr>F Rotation 5 11.62 2.32 6.64 0.0196 Block(Rotation) 6 2.10 0.35 1.40 0.2913 Error 12 3.00 0.25 Total 23 16.72 2.92 Table A2.4. Decline in resting spores, log residuals: Type I MS
Source df Sum of Squares Mean Square F value Pr>F Rotation 5 11.62 2.32 9.29 0.0008 Block(Rotation) 6 2.10 0.35 1.40 0.2913 Error 12 3.00 0.25 Total 23 16.72 2.92 Table A2.5. Decline in resting spores, qPCR: Type III MS block(rotation) as Error Term
Source df Sum of Squares Mean Square F value Pr>F Rotation 5 11.62 2.32 6.64 0.0196 Block(Rotation) 6 2.10 0.35 1.40 0.2913 Error 12 3.00 0.25 Total 23 16.72 2.92
174 Table A2.6. Controlled environment study: crop rotation and spore concentration: plant height 2 WAI
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0 . . . Residual 0.2977 0.06077 4.90 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 0.50 0.4813 Soil (S) 1 45 15.42 0.0003 Rotation (R) 1 45 1.67 0.2029 Inoculation (I) 1 45 3.38 0.0728 S x R 1 45 2.98 0.0910 R x I 1 45 0.01 0.9093 I x S 1 45 0.68 0.4139 S x R x I 1 45 0.10 0.7568 Repetition S x R 1 45 0.02 0.8913 Repetition x R x I 1 45 0.01 0.9419 Repetition x I x S 1 45 1.01 0.3210 Repetition x S x R x I 1 45 2.47 0.1231 Table A2.7. Controlled environment study: crop rotation and spore concentration: plant height 3 WAI
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0 . . . Residual 0.3233 0.06599 4.90 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 0.56 0.4600 Soil (S) 1 45 5.50 0.0235 Rotation (R) 1 45 1.94 0.1709 Inoculation (I) 1 45 3.98 0.0522 S x R 1 45 1.92 0.1722 R x I 1 45 0.55 0.6607 I x S 1 45 0.62 0.4366 S x R x I 1 45 0.35 0.5543 Repetition S x R 1 45 0.12 0.7283 Repetition x R x I 1 45 0.26 0.6140 Repetition x I x S 1 45 0.37 0.5485 Repetition x S x R x I 1 45 1.59 0.2143
175 Table A2.8. Controlled environment study: crop rotation and spore concentration: plant height 4 WAI
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0 . . . Residual 0.4233 0.08641 4.90 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 2.97 0.0917 Soil (S) 1 45 0.23 0.6320 Rotation (R) 1 45 0.18 0.6704 Inoculation (I) 1 45 4.17 0.0470 S x R 1 45 2.05 0.1593 R x I 1 45 0.14 0.7069 I x S 1 45 1.35 0.2512 S x R x I 1 45 0.45 0.5035 Repetition S x R 1 45 0.64 0.4273 Repetition x R x I 1 45 1.31 0.2590 Repetition x I x S 1 45 0.05 0.8202 Repetition x S x R x I 1 45 0.25 0.6185 Table A2.9. Controlled environment study: crop rotation and spore concentration: plant height 5 WAI
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0.04533 0.06688 0.68 0.2490 Residual 0.5768 0.1216 4.74 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 0.27 0.6090 Soil (S) 1 45 7.64 0.0083 Rotation (R) 1 45 0.20 0.6577 Inoculation (I) 1 45 4.58 0.0377 S x R 1 45 1.63 0.2087 R x I 1 45 0.14 0.7080 I x S 1 45 0.44 0.5085 S x R x I 1 45 0.53 0.4717 Repetition S x R 1 45 0.59 0.4481 Repetition x R x I 1 45 0.91 0.3441 Repetition x I x S 1 45 0.37 0.5467 Repetition x S x R x I 1 45 0.13 0.7178
176 Table A2.10. Controlled environment study: crop rotation and spore concentration: plant height 6 WAI
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0.4210 0.4397 0.96 0.1692 Residual 1.8665 0.3935 4.74 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 6.22 0.0164 Soil (S) 1 45 34.24 <0.0001 Rotation (R) 1 45 2.22 0.1431 Inoculation (I) 1 45 4.35 0.0428 S x R 1 45 0.11 0.7468 R x I 1 45 0.05 0.8279 I x S 1 45 0.02 0.9008 S x R x I 1 45 0.07 0.7941 Repetition S x R 1 45 0.25 0.6217 Repetition x R x I 1 45 0.01 0.9427 Repetition x I x S 1 45 0.39 0.5340 Repetition x S x R x I 1 45 0.04 0.8350 Table A2.11. Controlled environment study: crop rotation and spore concentration: Area Under Growth Stairs
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0 . . . Residual 286.52 58.4860 4.90 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 0.93 0.3411 Soil (S) 1 45 0.19 0.6639 Rotation (R) 1 45 0.77 0.3862 Inoculation (I) 1 45 4.44 0.0407 S x R 1 45 13.36 0.0007 R x I 1 45 0.18 0.6761 I x S 1 45 0.80 0.3760 S x R x I 1 45 0.38 0.5383 Repetition S x R 1 45 1.26 0.2941 Repetition x R x I 1 45 0.37 0.6953 Repetition x I x S 1 45 0.59 0.5569 Repetition x S x R x I 1 45 0.60 0.5538
177 Table A2.12. Controlled environment study – crop rotation and spore concentration: biomass (dry shoot weight)
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0.06136 0.07768 0.79 0.2148 Residual 0.5340 0.1126 4.74 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 6.53 0.0141 Soil (S) 1 45 6.78 0.0125 Rotation (R) 1 45 1.31 0.2578 Inoculation (I) 1 45 9.21 0.0040 S x R 1 45 16.16 0.0002 R x I 1 45 0.48 0.4910 I x S 1 45 0.00 0.9864 S x R x I 1 45 0.12 0.7262 Repetition S x R 1 45 0.17 0.6859 Repetition x R x I 1 45 0.15 0.7009 Repetition x I x S 1 45 0.33 0.5660 Repetition x S x R x I 1 45 0.04 0.8410 Table A2.13. Controlled environment study – crop rotation and spore concentration: biomass (dry root weight)
Random Effects Estimate Standard Error Z value Pr>Z Block(Repetition) 0 . . . Residual 0.2304 0.04704 4.90 <0.0001 Fixed Effects Numerator df Denominator df F value Pr>F Repetition 1 45 0.31 0.5832 Soil (S) 1 45 8.52 0.0055 Rotation (R) 1 45 0.08 0.7843 Inoculation (I) 1 45 0.94 0.3371 S x R 1 45 4.29 0.0440 R x I 1 45 1.20 0.2797 I x S 1 45 0.11 0.7466 S x R x I 1 45 0.00 0.9707 Repetition S x R 1 45 10.05 0.0027 Repetition x R x I 1 45 0.00 0.9523 Repetition x I x S 1 45 0.60 0.4411 Repetition x S x R x I 1 45 1.63 0.2077
178 Table A2.14. Raw data for decline in resting spores (g -1 soil) following susceptible canola – Normandin, QC
Break interval (years) Block Part Initial Eff.% R2 Final 0 3 1 31842906.8 99.4 0.97 489754836.7 0 3 2 10736450.5 99.4 0.97 13566309.7 0 4 1 18367851.3 99.4 0.97 20425424.5 0 4 2 2127172.3 99.4 0.97 2127172.3 1 58 1 3660434.3 99.4 0.97 3660434.3 1 58 2 550095.4 99.4 0.97 577176.2 1 59 1 7147390.5 99.4 0.98 9865309.3 1 59 2 4469322.2 99.4 0.98 4486544.3 2 81 1 721554.4 99.4 0.98 1038424.3 2 81 2 217536.2 99.4 0.98 260023.4 2 85 1 902728.1 99.4 0.98 902728.1 2 85 2 1747163.9 99.4 0.98 1747163.9 3 98 1 651739.4 99.4 0.98 747297.1 3 98 2 898102.4 99.4 0.97 1260552.5 3 111 1 110304.8 91.8 1.00 474457.9 3 111 2 160636.5 91.8 1.00 397480.0 5 112 1 149810.9 91.8 1.00 367118.9 5 112 2 89076.2 91.8 1.00 234167.2 5 103 1 49711.5 91.8 1.00 163516.6 5 103 2 48105.1 91.8 1.00 129578.9 6 93 1 95385.8 96.7 0.97 95385.8 6 93 2 1134565.0 96.7 0.97 1134565.0 6 94 1 208450.0 96.7 0.97 208450.0 6 94 2 166969.3 96.7 0.97 218985.7
179 Table A2.15. Raw data for decline in resting spores (g -1 soil) before and after a resistant canola crop – Normandin, QC
Sample date
Break interval (years)
Block Part Initial Eff.% R2 Final
Spring 2014 0 62 1 1651544.5 95.0 0.99 2407924.0 Spring 2014 0 62 2 2432379.4 95.0 0.99 4085490.3 Spring 2014 0 64 1 751138.1 95.0 0.99 1390224.9 Spring 2014 0 64 2 778070.2 95.0 0.99 1008971.3 Spring 2014 0 57 1 24030.5 104.0 1.00 2433273.5 Spring 2014 0 57 2 74935.4 104.0 1.00 792413.2 Spring 2014 0 61 1 2878129.9 104.0 1.00 222577291.0 Spring 2014 0 61 2 4400837.8 104.0 1.00 38820982.7 Spring 2014 0 63 1 865120.5 104.0 1.00 6112557.0 Spring 2014 0 63 2 1044708 104.0 1.00 19444601.8 Spring 2014 0 66 1 4842109.8 102.7 0.99 4842109.8 Spring 2014 0 66 2 17842108.9 102.7 0.99 43852775.5 Spring 2014 0 67 1 30437267.3 102.7 0.99 3012874574.7 Spring 2014 0 67 2 1897319.5 102.7 0.99 3063775.6 Spring 2014 0 68 1 9490721.5 102.7 0.99 9490721.5 Spring 2014 0 68 2 5393416.2 102.7 0.99 6333519.5 Spring 2014 1 72 1 980652.4 90.9 0.97 2581562.3 Spring 2014 1 72 2 6438091 90.9 0.97 7502654.0 Spring 2014 1 74 1 4171803.9 90.9 0.97 7822848.2 Spring 2014 1 74 2 7893526.2 90.9 0.97 17115389.4 Spring 2014 1 76 1 3070363.6 95.0 0.99 11387450.6 Spring 2014 1 76 2 2948671.3 95.0 0.99 33757681.4 Spring 2014 1 75 1 11829802.9 102.7 1.00 11829802.8 Spring 2014 1 75 2 795743 102.7 1.00 2902677.6 Spring 2014 1 78 1 25239920.1 96.7 1.00 28063929.4 Spring 2014 1 78 2 81249517.2 96.7 1.00 81249517.2 Spring 2014 1 79 1 54963075.3 96.7 1.00 55828524.1 Spring 2014 1 79 2 5481857.7 96.7 1.00 6075074.2 Spring 2014 1 80 1 1120969.6 96.7 1.00 1469296.0 Spring 2014 1 80 2 2489989.4 96.7 1.00 4048575.5 Spring 2014 1 84 1 2560205.8 96.7 1.00 3377778.0 Spring 2014 1 84 2 1095075.2 102.7 0.99 1376149.4 Spring 2014 2 105 1 943795.7 90.9 0.97 2506020.5 Spring 2014 2 105 2 1142227.3 90.9 0.97 1909299.3 Spring 2014 2 106 1 616045.7 90.9 0.97 1142510.9 Spring 2014 2 106 2 61862.8 95.0 0.99 96755.0 Spring 2014 2 109 1 346908.8 98.2 0.99 636929.6
180 Spring 2014 2 109 2 638354.4 98.2 0.99 737502.0 Spring 2014 2 110 1 626307.4 98.2 0.99 1816699.4 Spring 2014 2 110 2 1722110 98.2 0.99 4532246.3 Fall 2014 0 57 1 4009275.2 100.4 0.99 12548967.5 Fall 2014 0 57 2 4389860.8 100.4 0.99 18374536.3 Fall 2014 0 62 1 33975334 100.4 0.99 286276162.8 Fall 2014 0 62 2 13121072.1 100.4 0.99 52611456.8 Fall 2014 0 64 1 54824620.6 100.4 0.99 125596197.0 Fall 2014 0 64 2 104555166.7 100.4 0.99 1028095440.1 Fall 2014 0 63 1 6236049.5 100.4 0.99 48642096.4 Fall 2014 0 63 2 36554772.8 93.6 0.98 36554772.8 Fall 2014 0 61 2 55606010.6 93.6 0.98 55606010.6 Fall 2014 0 61 1 20959422.5 93.6 0.98 20959422.5 Fall 2014 0 66 1 28964918.2 98.0 0.99 1010164290.6 Fall 2014 0 66 2 1381877.6 98.0 0.99 5589932.9 Fall 2014 0 67 1 3435511.9 98.0 0.99 34678241.9 Fall 2014 0 67 2 40368238.5 98.0 0.99 36748276082.1 Fall 2014 0 68 1 19896775 98.0 0.99 224088420.3 Fall 2014 0 68 2 8684497.9 98.0 0.99 448972582.9 Fall 2014 1 72 1 10363884.7 93.6 0.98 10363884.7 Fall 2014 1 72 2 3604292.6 93.6 0.98 3604292.6 Fall 2014 1 74 1 29772023.1 93.6 0.98 29772023.1 Fall 2014 1 74 2 87501751.1 93.6 0.98 87501751.1 Fall 2014 1 76 1 16723855.5 98.2 0.99 16723855.5 Fall 2014 1 76 2 4320259.9 98.2 0.99 4320259.9 Fall 2014 1 78 1 9794831.8 109.1 0.98 16315751.8 Fall 2014 1 78 2 1300472.6 109.1 0.98 3961106.4 Fall 2014 1 79 1 473213.3 109.1 0.98 608892.2 Fall 2014 1 79 2 4910536.7 109.1 0.98 5144730.0 Fall 2014 1 80 1 8499003.4 109.1 0.98 14202418.5 Fall 2014 1 80 2 5312898.9 109.1 0.98 5312898.9 Fall 2014 1 84 1 4804560.5 109.1 0.98 4842045.6 Fall 2014 1 84 2 3694633.3 98.0 0.99 13733426.3 Fall 2014 1 75 1 7444487.3 102.7 1.00 7925338.2 Fall 2014 1 75 2 4594269.9 102.7 1.00 5067467.5 Fall 2014 2 105 1 19272362.8 108.7 0.98 19272362.8 Fall 2014 2 105 2 3016737.3 108.7 0.98 10785712.6 Fall 2014 2 106 1 1231227.5 108.7 0.98 4519633.6 Fall 2014 2 106 2 798544.8 108.7 0.98 2578024.3 Fall 2014 2 109 1 8208424.4 108.7 0.98 23565906.8 Fall 2014 2 109 2 2646966 108.7 0.98 9507192.6 Fall 2014 2 110 1 6943413 108.7 0.98 22825209.2 Fall 2014 2 110 2 61862.8 98.2 0.99 9235407.1
181 Spring 2015 0 57 1 773872 103.2 1.00 15382540.8 Spring 2015 0 61 1 4252068.6 103.2 1.00 20787474.6 Spring 2015 0 62 1 33233427.1 103.2 1.00 464890407.6 Spring 2015 0 63 1 27519.6 93.2 1.00 65331202.4 Spring 2015 0 64 1 100906027.2 103.2 1.00 100906027.2 Spring 2015 0 66 1 3147604.5 93.2 1.00 4540324.0 Spring 2015 0 67 1 958731.2 93.2 1.00 958731.2 Spring 2015 0 68 1 660392.3 93.2 1.00 660392.3 Spring 2015 1 69 1 23561.9 93.2 1.00 101009.3 Spring 2015 1 70 1 123488.8 93.2 1.00 409156.9 Spring 2015 1 82 1 5249.8 93.2 1.00 8946.4 Spring 2015 2 95 1 17864 93.2 1.00 542752.6 Spring 2015 2 96 1 73407.1 93.2 1.00 2094928.3 Table A2.16. Raw data for controlled environmental study – crop rotation and spore concentration: CI & DSI
Repetition Soil Spores ml-1 Cultivar Block CI DSI 1 Scott 1x106 ACSN39 1 71.4 47.6 1 Scott 1x106 ACSN39 2 80.0 53.3 1 Scott 1x106 ACSN39 3 100.0 70.4 1 Scott 1x106 ACSN39 4 100.0 83.3 1 Scott 0 ACSN39 1 0.0 0.0 1 Scott 0 ACSN39 2 0.0 0.0 1 Scott 0 ACSN39 3 0.0 0.0 1 Scott 0 ACSN39 4 0.0 0.0 1 Scott 1x106 45H29 1 0.0 0.0 1 Scott 1x106 45H29 2 0.0 0.0 1 Scott 1x106 45H29 3 0.0 0.0 1 Scott 1x106 45H29 4 0.0 0.0 1 Scott 0 45H29 1 0.0 0.0 1 Scott 0 45H29 2 0.0 0.0 1 Scott 0 45H29 3 0.0 0.0 1 Scott 0 45H29 4 0.0 0.0 1 Scott 1x106 7367 1 0.0 0.0 1 Scott 1x106 7367 2 0.0 0.0 1 Scott 1x106 7367 3 0.0 0.0 1 Scott 1x106 7367 4 0.0 0.0 1 Scott 0 7367 1 0.0 0.0 1 Scott 0 7367 2 0.0 0.0 1 Scott 0 7367 3 0.0 0.0 1 Scott 0 7367 4 0.0 0.0 1 Scott 1x106 7377 1 0.0 0.0
182 1 Scott 1x106 7377 2 0.0 0.0 1 Scott 1x106 7377 3 0.0 0.0 1 Scott 1x106 7377 4 0.0 0.0 1 Scott 0 7377 1 0.0 0.0 1 Scott 0 7377 2 0.0 0.0 1 Scott 0 7377 3 0.0 0.0 1 Scott 0 7377 4 0.0 0.0 2 Scott 1x106 ACSN39 1 100.0 62.5 2 Scott 1x106 ACSN39 2 87.5 54.2 2 Scott 1x106 ACSN39 3 80.0 60.0 2 Scott 1x106 ACSN39 4 100.0 100.0 2 Scott 0 ACSN39 1 0.0 0.0 2 Scott 0 ACSN39 2 0.0 0.0 2 Scott 0 ACSN39 3 0.0 0.0 2 Scott 0 ACSN39 4 0.0 0.0 2 Scott 1x106 45H29 1 0.0 0.0 2 Scott 1x106 45H29 2 0.0 0.0 2 Scott 1x106 45H29 3 0.0 0.0 2 Scott 1x106 45H29 4 0.0 0.0 2 Scott 0 45H29 1 0.0 0.0 2 Scott 0 45H29 2 0.0 0.0 2 Scott 0 45H29 3 0.0 0.0 2 Scott 0 45H29 4 0.0 0.0 2 Scott 1x106 7367 1 0.0 0.0 2 Scott 1x106 7367 2 0.0 0.0 2 Scott 1x106 7367 3 0.0 0.0 2 Scott 1x106 7367 4 0.0 0.0 2 Scott 0 7367 1 0.0 0.0 2 Scott 0 7367 2 0.0 0.0 2 Scott 0 7367 3 0.0 0.0 2 Scott 0 7367 4 0.0 0.0 2 Scott 1x106 7377 1 0.0 0.0 2 Scott 1x106 7377 2 0.0 0.0 2 Scott 1x106 7377 3 0.0 0.0 2 Scott 1x106 7377 4 0.0 0.0 2 Scott 0 7377 1 0.0 0.0 2 Scott 0 7377 2 0.0 0.0 2 Scott 0 7377 3 0.0 0.0 2 Scott 0 7377 4 0.0 0.0 1 Elora 1x106 ACSN39 1 80.0 30.0 1 Elora 1x106 ACSN39 2 60.0 30.0 1 Elora 1x106 ACSN39 3 50.0 26.7 1 Elora 1x106 ACSN39 4 62.5 20.8
183 1 Elora 0 ACSN39 1 0.0 0.0 1 Elora 0 ACSN39 2 0.0 0.0 1 Elora 0 ACSN39 3 0.0 0.0 1 Elora 0 ACSN39 4 0.0 0.0 1 Elora 1x106 45H29 1 0.0 0.0 1 Elora 1x106 45H29 2 0.0 0.0 1 Elora 1x106 45H29 3 0.0 0.0 1 Elora 1x106 45H29 4 0.0 0.0 1 Elora 0 45H29 1 0.0 0.0 1 Elora 0 45H29 2 0.0 0.0 1 Elora 0 45H29 3 0.0 0.0 1 Elora 0 45H29 4 0.0 0.0 1 Elora 1x106 7367 1 0.0 0.0 1 Elora 1x106 7367 2 0.0 0.0 1 Elora 1x106 7367 3 0.0 0.0 1 Elora 1x106 7367 4 0.0 0.0 1 Elora 0 7367 1 0.0 0.0 1 Elora 0 7367 2 0.0 0.0 1 Elora 0 7367 3 0.0 0.0 1 Elora 0 7367 4 0.0 0.0 1 Elora 1x106 7377 1 0.0 0.0 1 Elora 1x106 7377 2 0.0 0.0 1 Elora 1x106 7377 3 0.0 0.0 1 Elora 1x106 7377 4 0.0 0.0 1 Elora 0 7377 1 0.0 0.0 1 Elora 0 7377 2 0.0 0.0 1 Elora 0 7377 3 0.0 0.0 1 Elora 0 7377 4 0.0 0.0 2 Elora 1x106 ACSN39 1 80.0 43.3 2 Elora 1x106 ACSN39 2 60.0 26.7 2 Elora 1x106 ACSN39 3 55.6 25.9 2 Elora 1x106 ACSN39 4 70.0 33.3 2 Elora 0 ACSN39 1 0.0 0.0 2 Elora 0 ACSN39 2 0.0 0.0 2 Elora 0 ACSN39 3 0.0 0.0 2 Elora 0 ACSN39 4 0.0 0.0 2 Elora 1x106 45H29 1 0.0 0.0 2 Elora 1x106 45H29 2 0.0 0.0 2 Elora 1x106 45H29 3 0.0 0.0 2 Elora 1x106 45H29 4 0.0 0.0 2 Elora 0 45H29 1 0.0 0.0 2 Elora 0 45H29 2 0.0 0.0 2 Elora 0 45H29 3 0.0 0.0
184 2 Elora 0 45H29 4 0.0 0.0 2 Elora 1x106 7367 1 0.0 0.0 2 Elora 1x106 7367 2 0.0 0.0 2 Elora 1x106 7367 3 0.0 0.0 2 Elora 1x106 7367 4 0.0 0.0 2 Elora 0 7367 1 0.0 0.0 2 Elora 0 7367 2 0.0 0.0 2 Elora 0 7367 3 0.0 0.0 2 Elora 0 7367 4 0.0 0.0 2 Elora 1x106 7377 1 0.0 0.0 2 Elora 1x106 7377 2 0.0 0.0 2 Elora 1x106 7377 3 0.0 0.0 2 Elora 1x106 7377 4 0.0 0.0 2 Elora 0 7377 1 0.0 0.0 2 Elora 0 7377 2 0.0 0.0 2 Elora 0 7377 3 0.0 0.0 2 Elora 0 7377 4 0.0 0.0
Table A2.17. Raw data for controlled environmental study – crop rotation and spore concentration: plant height
Repe-tition Soil
Rota-tion
Spores ml-1 Block 2 WAI 3 WAI 4 WAI 5 WAI 6 WAI
1 Scott 2 1x106 1 3.27 3.55 3.81 4.15 5.79 1 Scott 2 1x106 2 3.84 4.29 4.60 5.31 7.07 1 Scott 2 1x106 3 4.79 5.14 5.48 6.15 8.84 1 Scott 2 1x106 4 4.08 4.81 5.04 6.13 6.96 1 Scott 2 0 1 3.60 3.83 4.26 4.89 6.07 1 Scott 2 0 2 4.21 4.52 4.87 5.28 6.61 1 Scott 2 0 3 4.18 4.48 5.22 6.53 10.05 1 Scott 2 0 4 5.13 5.37 5.71 6.47 9.66 1 Scott 0 1x106 1 3.24 3.61 3.99 4.61 5.80 1 Scott 0 1x106 2 3.30 3.73 3.96 4.58 5.90 1 Scott 0 1x106 3 3.03 3.52 3.92 4.62 7.02 1 Scott 0 1x106 4 3.44 3.64 4.03 4.88 6.63 1 Scott 0 0 1 4.41 4.81 5.07 5.92 6.81 1 Scott 0 0 2 4.22 4.61 5.21 5.81 6.46 1 Scott 0 0 3 4.05 4.65 5.41 6.89 9.83 1 Scott 0 0 4 3.73 4.37 4.85 5.18 6.63 2 Scott 2 1x106 1 3.22 3.67 3.66 3.98 4.32 2 Scott 2 1x106 2 4.18 4.56 4.81 5.09 5.43 2 Scott 2 1x106 3 3.71 4.19 4.31 4.97 5.24 2 Scott 2 1x106 4 2.61 2.86 3.02 3.71 4.21 2 Scott 2 0 1 4.10 4.37 4.43 4.77 5.63
185 2 Scott 2 0 2 4.01 4.54 4.98 5.33 6.19 2 Scott 2 0 3 3.87 4.34 4.52 5.02 5.60 2 Scott 2 0 4 3.04 3.26 3.54 3.79 4.21 2 Scott 0 1x106 1 3.46 3.86 4.26 4.59 4.69 2 Scott 0 1x106 2 3.57 3.73 3.93 4.47 5.10 2 Scott 0 1x106 3 3.37 3.54 3.94 4.48 4.60 2 Scott 0 1x106 4 2.53 2.76 2.91 2.97 3.28 2 Scott 0 0 1 3.67 4.38 4.79 5.04 5.38 2 Scott 0 0 2 3.19 3.45 3.86 4.16 4.39 2 Scott 0 0 3 3.93 4.27 4.32 4.65 5.27 2 Scott 0 0 4 2.11 2.53 2.97 3.49 4.17 1 Elora 2 1x106 1 2.79 3.18 4.10 4.75 5.36 1 Elora 2 1x106 2 2.91 3.83 4.10 4.75 5.59 1 Elora 2 1x106 3 2.64 3.24 4.22 4.68 6.08 1 Elora 2 1x106 4 2.85 3.62 4.88 5.58 6.59 1 Elora 2 0 1 2.90 3.46 3.94 4.69 5.65 1 Elora 2 0 2 2.89 3.43 4.20 4.77 5.75 1 Elora 2 0 3 3.09 3.82 4.47 5.53 6.46 1 Elora 2 0 4 3.51 4.10 5.54 5.92 7.29 1 Elora 0 1x106 1 3.31 3.67 4.16 4.63 5.70 1 Elora 0 1x106 2 2.86 3.48 3.90 4.92 5.49 1 Elora 0 1x106 3 2.80 3.27 4.32 4.80 5.60 1 Elora 0 1x106 4 3.34 3.45 4.24 4.79 5.62 1 Elora 0 0 1 2.90 3.44 4.19 4.80 6.06 1 Elora 0 0 2 2.99 3.56 4.18 4.87 5.72 1 Elora 0 0 3 2.37 2.93 4.32 5.07 6.08 1 Elora 0 0 4 3.40 4.19 5.23 5.92 7.02 2 Elora 2 1x106 1 3.79 4.22 4.38 5.59 9.14 2 Elora 2 1x106 2 3.25 3.73 4.30 5.69 10.99 2 Elora 2 1x106 3 2.34 3.08 3.62 4.90 11.77 2 Elora 2 1x106 4 3.52 4.14 4.53 5.72 8.50 2 Elora 2 0 1 4.00 4.44 5.14 6.70 12.83 2 Elora 2 0 2 3.18 3.81 4.18 5.54 10.60 2 Elora 2 0 3 2.46 3.13 3.52 5.09 10.06 2 Elora 2 0 4 3.38 4.06 4.67 6.19 9.42 2 Elora 0 1x106 1 2.79 3.34 3.73 4.68 6.63 2 Elora 0 1x106 2 2.43 3.08 3.73 5.26 8.86 2 Elora 0 1x106 3 3.99 4.80 5.87 8.16 15.02 2 Elora 0 1x106 4 3.26 3.78 5.87 6.21 6.60 2 Elora 0 0 1 4.10 4.36 4.68 5.76 9.26 2 Elora 0 0 2 2.35 2.97 3.69 5.21 8.14 2 Elora 0 0 3 4.10 5.03 5.88 8.26 12.86 2 Elora 0 0 4 3.46 3.93 4.41 5.27 11.05
186 Table A2.18. Raw data for controlled environmental study – crop rotation and spore concentration: biomass (dry shoot weight)
Repetition Soil Rotation Spore ml-1 Block
Dry shoot weight (g)
1 Scott 2 1x106 1 3.30 1 Scott 2 1x106 2 3.83 1 Scott 2 1x106 3 3.13 1 Scott 2 1x106 4 2.75 1 Scott 2 0 1 3.50 1 Scott 2 0 2 4.00 1 Scott 2 0 3 5.00 1 Scott 2 0 4 3.67 1 Scott 0 1x106 1 2.11 1 Scott 0 1x106 2 2.00 1 Scott 0 1x106 3 3.00 1 Scott 0 1x106 4 2.50 1 Scott 0 0 1 3.11 1 Scott 0 0 2 2.11 1 Scott 0 0 3 3.25 1 Scott 0 0 4 2.50 2 Scott 2 1x106 1 1.90 2 Scott 2 1x106 2 2.44 2 Scott 2 1x106 3 2.11 2 Scott 2 1x106 4 2.00 2 Scott 2 0 1 2.89 2 Scott 2 0 2 3.22 2 Scott 2 0 3 2.60 2 Scott 2 0 4 2.57 2 Scott 0 1x106 1 2.00 2 Scott 0 1x106 2 1.71 2 Scott 0 1x106 3 2.67 2 Scott 0 1x106 4 2.80 2 Scott 0 0 1 2.25 2 Scott 0 0 2 1.75 2 Scott 0 0 3 2.00 2 Scott 0 0 4 4.75 1 Elora 2 1x106 1 2.00 1 Elora 2 1x106 2 1.80 1 Elora 2 1x106 3 1.83 1 Elora 2 1x106 4 2.20 1 Elora 2 0 1 2.00 1 Elora 2 0 2 2.50 1 Elora 2 0 3 2.00 1 Elora 2 0 4 3.40
187 1 Elora 0 1x106 1 2.80 1 Elora 0 1x106 2 3.10 1 Elora 0 1x106 3 2.00 1 Elora 0 1x106 4 2.78 1 Elora 0 0 1 2.90 1 Elora 0 0 2 2.60 1 Elora 0 0 3 2.33 1 Elora 0 0 4 3.56 2 Elora 2 1x106 1 2.10 2 Elora 2 1x106 2 3.20 2 Elora 2 1x106 3 3.70 2 Elora 2 1x106 4 3.10 2 Elora 2 0 1 3.80 2 Elora 2 0 2 2.00 2 Elora 2 0 3 3.70 2 Elora 2 0 4 5.44 2 Elora 0 1x106 1 3.20 2 Elora 0 1x106 2 4.70 2 Elora 0 1x106 3 5.50 2 Elora 0 1x106 4 3.90 2 Elora 0 0 1 4.40 2 Elora 0 0 2 4.00 2 Elora 0 0 3 5.20 2 Elora 0 0 4 6.90
