IDENTIFICATION OF FUNGI AND BACTERIA ASSOCIATED WITH
INTERNALLY DISCOLORED HORSERADISH ROOTS
BY
JUNMYOUNG YU
THESIS
Submitted in partial fulfillment of the requirements
for the degree of Master of Science in Crop Sciences
in the Graduate College of the
University of Illinois at Urbana-Champaign, 2010
Urbana, Illinois
Adviser:
Professor Mohammad Babadoost
ABSTRACT
This study was conducted to identify fungi and bacteria associated with the
internally discolored horseradish roots. During 2008-2009, 75, 306, 115, and 120,
horseradish roots from California, Illinois, Ontario (Canada), and Wisconsin, respectively,
were collected and tested for the presence of fungi and bacteria. The roots were surfacedisinfected and five segments (5 mm thick) were cut from each root and placed onto
acidified potato dextrose agar (A-PDA) and nutrient agar (NA) in Petri plates. The plates
were incubated at 22-24 C with 12 h light/12 h darkness.
Morphological characteristics of fungal isolates were examined and compared
with the references. Molecular identification of the fungal isolates was accomplished by
sequence analysis of the PCR products, using fungal universal primers (ITS5 and ITS4)
for all fungal isolates, mitochondrial DNA primers cox3A-cox3B and nad1A-nad1B for
Verticillium isolates, and elongation factor 1 alpha primer EF-1H-EF-2T and
mitochondrial small subunit rDNA primers NMS1-NMS2 for Fusarium isolates. A total
of 18 fungal genera were identified, which included Alternaria (3.8% roots),
Arthopyreniaceae (0.4%), Chaetomium (0.6%), Colletotrichum (2.3%), Cordyceps
(2.3%), Cylindrocarpon (0.9%), Fusarium (31.8%), Neonectria (1.3%), Penicillium
(4.4%), Phoma (0.1%), Phomopsis (0.4%), Pyrenochaeta (2.7%), Rhizoctonia (0.1%),
Rhizopus (1.0%), Rhizopycnis (0.4), Trichurus (0.6%), Ulocladium (0.4%), and
Verticillium (38.0%). Verticillium (38.0%) and Fusarium (31.8%) were the most
frequently isolated genera. Two species of Verticillium and six species of Fusarium were
identified which were two types of V. dahliae, V. longisporum, F. acuminatum, F.
ii
commune, F. equiseti-like species, F. oxysporum, F. prolfiertaum, and F. solani. In
greenhouse tests Verticillium dahliae, V. longisporum, Fusarium acuminatum, F.
commune, F. oxysporum, and F. solani caused internal root discoloration, and F.
commune caused root rot in horseradish plants.
Single-cell colonies of isolated bacteria were grown on NA. Characteristics of
each purified colony were recorded. Isolated bacteria were identified based on the
analysis of sequences of 16S rDNA, resulting from polymerase chain reaction (PCR) restriction fragment length polymorphism (RFLP) assay. A total of 52 bacterial species in
20 genera were identified, which were classified into five classes, Actinobacteria (50
roots), Alphaproteobacteria (10 roots), Bacilli (234 roots), Betaproteobacteria (4 roots),
and Gammaproteobacteria (287 roots). The majority of the isolates were identified as
Bacillus and Pseudomonas, which were isolated from 185 and 204 roots, respectively.
Results of in vitro tests showed that three Bacillus isolates and two Pseudomonas isolates
strongly inhibited the growth of Verticillium and Fusarium species.
iii
To my Family
iv
ACKNOWLEDGMENTS
I would like to acknowledge Dr. Mohammad Babadoost for his sincere advice and
support, constant encouragement, and giving me an opportunity to work under his
guidance. I also would like to convey my great appreciation to Drs. Darin Eastburn, Kris
Lambert, Andrew Miller, and Frank Zhao for serving as my committee members,
providing good instruction and deep encouragement. I also thank the faculty members of
Department of Crop Sciences for their excellent instruction and valuable lessons on my
coursework.
v
TABLE OF CONTENTS
CHAPTER 1 ........................................................................................................................1
LITERATURE REVIEW ...............................................................................................1
THESIS OBJECTIVES ................................................................................................10
LITERATURE CITED ................................................................................................11
CHAPTER 2 ......................................................................................................................15
FUNGAL SPECIES ASSOCIATED WITH INTERNALLY DISCOLORED
HORSERADISH ROOTS .................................................................................................15
MATERIALS AND METHODS ..................................................................................15
RESULTS .....................................................................................................................26
DISCUSSION ..............................................................................................................37
FIGURES AND TABLES ...........................................................................................42
LITERATURE CITED ................................................................................................65
CHAPTER 3 ......................................................................................................................68
BACTERIAL SPECIES ASSOCIATED WITH INTERNALLY DISCOLORED
HORSERADISH ROOTS .................................................................................................68
MATERIALS AND METHODS ..................................................................................68
RESULTS .....................................................................................................................72
DISCUSSION ..............................................................................................................76
FIGURES AND TABLES ...........................................................................................79
LITERATURE CITED .................................................................................................89
vi
CHAPTER 1
LITERATURE REVIEW
Origin of horseradish
Horseradish has been cultivated for at least 2000 years, and is thought to have
originated in temperate Eastern Europe and Western Asia; wild types of horseradish are
found growing from Finland and Poland to the regions around the Caspian Sea and the
deserts of Cumania (now Romania) and Turkey (De Candolle, 1959; Hedrick, 1919). The
most primitive name for horseradish is chren in Slavic languages of Eastern Europe.
Chren then spread to Germany and France and changed in various forms (e.g., kren,
kreen, cran and cranson). Meerretig, meer-radys and meridi, which means sea-radish,
are other names for horseradish that are common in western European languages. The
name of horseradish in English, raifort in French, pepperrot in Swedish are more recent
than chren. The first use of the term “horseradish” was made by John Gerard in his
famous English herbal (1633) that contains a lengthy entry with a woodcut and a clear
description of the plant (Shehata et al., 2009).
Nomenclature and classification of horseradish
Taxonomy of horseradish has changed over time. Since the 18th century,
horseradish has been placed in five genera: Armoracia, Cochlearia, Nasturtium, Rorippa
and Radicula (Robinson and Fernald, 1908). Ancient writers, Dioscorides of Greece and
Pliny of Rome, listed horseradish under Thlaspi or Persicon; early Renaissance
herbalists, such as Gerard (1633) classified it under Raphanus; Linnaeus considered it
1
under the genus Cochlearia; and today‟s taxonomists include it under the genus
Armoracia (Courter and Rhodes, 1969). “Armoracia” is derived from Celtic, which
means “by the sea” (Barton and Castle, 1877): ar = „near‟, mor = „the sea‟, rich =
„against‟, viz., a plant growing near the sea (Barton and Castle, 1877; Courter and
Rhodes, 1969).
Horseradish was first named Armoracia rusticana in the 1800s by Gaetner and his
coworkers (Courter and Rhodes, 1969; Fosberg, 1965). Since then, all modern literature
has been using this nomenclature for horseradish (Courter and Rhodes, 1969).
Horseradishes have been grouped into three types of cultivars based on their leaf
morphology (Rhodes et al., 1965). Type I plants (also called Bohemian types) includes
the plants with narrow, smooth leaves with tapered leaf bases. Type III plants (also called
Maliner Kren; common type) typically have broad, crinkled leaves with a heart-shaped
leaf base. Type II plants (also called Swiss cultivar) are intermediated between the other
two types; the leaves are smooth to slightly crinkled and rounded at the base (Courter and
Rhodes, 1969; Rhodes et al., 1965; Tucker and Debaggio, 2000). The common types are
considered to have superior root quality, and the Bohemian types have lower root quality.
Maliner Kren and Bohemian type are cultivars imported by the U.S. Department
of Agriculture (USDA) in 1899 (Anonymous, 1937). The common type has been
preferred for commercial cultivation in Illinois for many years (Horwitz, 1983). Other
commercially grown cultivars include Swiss cultivar (Type II), and Big Top Western, (a
2
Type I cultivar). „Big Top Western‟ gained popularity owing to its vigor and resistance to
white rust and mosaic diseases (Horwitz, 1983). Since horseradish is propagated
asexually (cut roots), which prevent cross-fertilization, the characteristics of the cultivars
have persisted and are present in horseradish cultivars grown today (Courter and Rhodes,
1969).
Cultivation
Horseradish was introduced into the United States (US) from Europe by early
settlers and became popular in gardens in the New England states in the early 1800s
(Shehata et al., 2009). Commercial cultivation of horseradish in the US began in the
1850s. It was brought to Chicago in 1856 by immigrants from Germany (Shehata et al.,
2009). By the late 1890s, a thriving horseradish industry had been developed in the
Mississippi river valley in Illinois and Eau Claire, Wisconsin (Horseradish information
council, 2001). In the Tule Lake region of northern California as well as Michigan, Ohio,
Pennsylvania, New Jersey, Connecticut, Massachusetts, and Washington, production of
horseradish began after World War II (Weber, 1949).
In most parts of the US, horseradish is cultivated commercially as an annual crop
and propagated asexually by planting lateral roots named as “sets”. Root pieces 25 to 35
cm (10 to 14 inches) long with a diameter of 1.0 to 3.75 cm (0.5 to 1.5 inches) are
considered ideal sets (Rhodes, 1977). Shorter root cuttings (15-20 cm; 6-8 inches) can
also be used (Dinkelmann, 2003). In order to produce a good horseradish yield, it is
necessary to remove all side roots, leaving only those at the bottom of the main root
3
(enlarged set). To produce a high value crop, the roots are “lifted” twice during the
season, the first lifting being done when the largest leaves are about 20 to 25 cm (8 to 10
inches) in length. In lifting the roots, the soil is first carefully removed, and then all
crowns except the main root crown are cut. About 6 weeks later, the roots are lifted again
and the newly grown side roots are removed (Anonymous, 1937). By practicing this
method, side roots are removed, which results in larger and smoother main roots (Rhodes,
1977).
Usage
Originally, horseradish was cultivated as a medicinal herb. Both the root and
leaves were used as a medicine during the Middle Ages and as a condiment in Denmark
and Germany (Grieve, 1967). Horseradish can reduce pain from sciatica, expell afterbirth, relieve colic, increase urination, and kill worms in children. Also, the fresh root of
horseradish is still considered natural medicine for an antiseptic, diaphoretic, diuretic,
rubefacient, stimulant, stomachic, and vermifuge (Simon et al., 1984). The material has
also been used as a remedy for asthma, cancer, coughs, colic, rheumatism, scurvy,
toothache, ulcers, and venereal diseases (Simon et al., 1984).
Use of horseradish spread to England and Scandinavia from central Europe during
the Renaissance, but the British would not eat it because they saw it as a low-grade root
(Dinkelmann, 2003). By the late 1600s, however, horseradish accompanied beef and
oysters on the tables of all British classes. It was also grown at inns and coach stations in
England to revive the weary travelers (Horseradish information council, 2001).
4
Early settlers brought horseradish to the North American colonies by the 1800s,
and its use has been prevalent in the northeastern part of the US since then (Dinkelmann,
2003). In the US, an estimated 10,886 metric ton (24 million pounds) of horseradish roots
are ground and processed annually to produce approximately 23 million liters (6 million
gallons) of prepared horseradish (Horseradish information council, 2001). In addition to
the most popular basic prepared horseradish, a number of other horseradish products are
available, including cream-style prepared horseradish, horseradish sauce, beet
horseradish sauce and dehydrated horseradish (Horseradish information council, 2001).
Cocktail sauces (specialty mustards), and many other sauces, dips, spreads, relishes, and
dressings may also contain horseradish.
Disease and insect pests
Horseradish is attacked by several pathogens and insects that are damaging where
effective control measures are not applied (Anonymous, 1937). Compared to other
vegetable crops, relatively fewer herbicides, insecticides, and fungicides are available for
use on horseradish.
Insect Pests. Several insects feed on the foliage, but foliage-feeding by insects is
less concerning, as horseradish can tolerate a significant amount of early damage before
yield is decreased. However, defoliation of plants by insects late in the season can be
damaging to the yield and the insects should be controlled. The major insects that cause
damage on horseradish plants are the beet leaf hopper (Circulifer tenellus) and the
horseradish flea beetle. The beet leaf hopper is the primary vector for brittle root disease
agent (Sprioplasma citri). Brittle root of horseradish is one of the destructive spiroplasma
5
diseases of plants (Bratsch, 2006; Shehata et al., 2009). Horseradish flea beetles (Epitrix
spp.) can damage emerging shoots after planting sets. In the northwest US, horseradish is
sometimes severely affected by the curly top disease, a viral disease transmitted by
leafhoppers (Anonymous, 1937).
Virus Diseases. Turnip mosaic virus (TuMV) is the most common virus disease
reported on horseradish (Bratsch, 2006; Dana and McWhorter, 1932; Herold, 1957; Li
and Cheo, 1964; Yoshii et al., 1963). TuMV occurs everywhere that horseradish is grown
and has a wide host range. It is transmitted by aphids and mechanical means. Symptoms
include foliar mosaic mottling, yellow rings on the leaf, and black streaking of the petiole
(Bratsch, 2006). TuMV and other viruses may play a secondary role in the horseradish
disease complex by weakening plants and making them more susceptible to other stresses,
such as drought, insects, and nematodes (Horwitz et al., 1985).
Fungal Diseases. Horseradish is susceptible to several foliar diseases (Bratsch,
2006). White rust, caused by the Albugo candida, a member of oomycota is one of the
most destructive foliar diseases of horseradish (Eastburn and Weinzierl, 1995). This
disease commonly occurs in Europe. In Illinois, white rust has been reported during the
spring and fall months following prolonged periods of cool, dewy nights and slightly
warmer days (Babadoost, 1990; Takashi et al., 2002). Shehata et al. (2009) reported that
white rust was not a significant problem in horseradish growing areas in the US since
1999. There are other fungal diseases of horseradish, which include Cercospora leaf spot
6
(Cercospora armoraciae) and Alternaria leaf spot (Alternaria brassicae), and internal
root discoloration (Verticillium and Fusarium species).
Internal discoloration of horseradish root
Internal discoloration of horseradish roots occurs throughout the world and is the
main limiting factor in horseradish production (Babadoost et al., 2004; Eastburn and
Chang, 1994; Gerber et al., 1983; Potschke, 1923; Stark, 1961). The disease has been
known in Europe since 1920 (Potschke, 1923). Internal discoloration of horseradish roots
in the US was first described in 1931 in Michigan, where 20% of the yield was lost due to
root infection (Boning, 1938). The discoloration of horseradish roots was reported in
western Washington in 1937 by Heald et al. (1937). The disease was diagnosed in
Wisconsin in 1973 (Mueller et al., 1982). Since the early 1980s, horseradish producers in
Illinois have experienced substantial reductions in marketable yield of horseradish due to
the internal discoloration of roots (Atibalentja and Eastburn, 1997; Babadoost, 2006;
Eastburn and Chang, 1994; Gerber et al., 1983). Yield losses of up to 100%, caused by
internal root discoloration, have frequently occurred in Illinois (Babadoost, 2006;
Babadoost et al., 2004).
Potschke (1923) was the first to report a Verticillium species as the causal agent of
the internal discoloration of horseradish. Heald et al. (1937) identified Verticillium
dahliae in the internally discolored roots of horseradish in Washington. Eastburn and
Chang (1994) reported V. dahliae as the primary causal agent of internal discoloration of
horseradish roots in Illinois. Percich & Johnson (1990) and Babadoost et al. (2004)
7
reported that internal discoloration of horseradish roots is a disease complex. Three
fungal species, V. dahliae, V. longisporum and Fusarium solani, were identified as the
causal agents of the internal root discoloration (Babadoost et al., 2004).
There has been no method available to provide an effective control of the internal
discoloration of horseradish root. Growers have tried to control the internal discoloration
of horseradish by soil fumigation and/or fungicide applications, but these practices have
failed to control the disease (Atibalentja and Eastburn, 1997). Intensive research has been
conducted at the University of Illinois to develop effective methods for management of
internal discoloration of horseradish roots. Management of the internal discoloration of
horseradish roots has been a challenging task because the causal pathogens are carried in
sets (set-borne inoculum) and survive in the soil (soil-borne inoculum) (Babadoost et al.,
2010). No chemical is available to control set-borne inoculum of the root discoloration
of horseradish. This was the main reason why soil-fumigation, crop rotations, and use of
fungicides have not been successful in controlling internal discoloration of horseradish
roots (Atibalentja and Eastburn, 1997; Babadoost, 2006).
Horseradish producers save their horseradish sets from their harvest to plant in the
following season. Most of the sets that the producers save are apparently healthy
(asymptomatic), but they are often infected with Verticillium and Fusarium spp.
(Babadoost, 2006). Set-transmission of these pathogens is important because infected sets
usually give rise to severely discolored roots, which are unmarketable (Babadoost, 2006;
8
Babadoost et al., 2004). Thus, starting horseradish production from pathogen-free sets is
essential for managing the internal discoloration of roots (Babadoost and Islam, 2002).
Pathogen-free sets of horseradish can be generated by tissue-culturing of
horseradish leaves (Meyer and Milbrath, 1977; Norton et al., 2001; Uchanski et al.,
2004). However, tissue-culture generation of horseradish sets with existing technologies
is not economically feasible (Aitken-Christie et al., 1995; Uchanski et al., 2007). In
addition, the sets can become infected before used by producers because the final stage of
tissue-culture set production involves increasing sets in the field for at least one full
growing season. Thus, alternative methods of producing pathogen-free horseradish sets
are needed.
Eranthodi and Babadoost (2007) developed a hot-water therapy to eradicate setborne inoculum of the internal discoloration of horseradish roots. They reported that
treatment of horseradish sets in 47 ºC for 20 minutes eradicated Verticillium and
Fusarium fungi carried in the sets without adversely affecting set germination or plant
vigor. Treatment of the sets in water with temperatures at < 46 ºC did not eradicate the
set-borne inoculum of the fungus, and treatment at temperature > 48 ºC reduced either set
germination and/or vigor of the plants grown from hot-water treated sets.
If pathogen-free sets are planted in a field with a history of the internal root
discoloration of horseradish, or in a field with a history of Verticillium and/or Fusarium
diseases in other crops, internal discoloration of roots could develop, which is caused by
9
soil-borne inoculum of these pathogens. Babadoost et al. (2010) evaluated the
effectiveness of several fungicides and bio-fungicides for control of horseradish plants
against soil-born inoculum of the internal discoloration of horseradish roots. They
reported that application of either the fungicide fludioxonil (Maxim 4FS or Maxim Potato
WP) or biofungicide Trichoderma virens (SoilGard 12G or G-41), onto the pathogen-free
sets prior to planting protected the plants against soil-borne Verticillium and Fusarium
species.
THESIS OBJECTIVES
The overall objective of this research was to identify all fungi and bacteria associated
with internally discolored horseradish roots. The specific objectives were:
1) To identify fungal and bacterial species associated with internally
discolored roots using classical identification methods;
2) To develop appropriated molecular methods for accurate identification of
fungal and bacterial species associated with internally discolored
horseradish roots;
3) To determine the pathogenecity of isolated fungal species on horseradish
roots; and
4) To determine interactions among isolated fungal and bacterial species
from horseradish roots.
10
LITERATURE CITED
Aitken-Christie, J., Kozai, T., and Smith, M.L.A. 1995. Automation and Environmental
Control in Plant Tissue Culture. Kluwer Academic Publishers. Dordrecht, The
Netherlands. 574 pp.
Anonymous. 1937. Production and preparation of horseradish. USDA Leaflet No. 129.
Atibalentja, N., and Eastburn, D.M. 1997. Evaluation of inoculation methods for
screening horseradish cultivars for resistance to Verticillium dahliae. Plant Dis. 81:356362.
Babadoost, M. 1990. White rust of vegetables. University of Illinois Extension. Report on
Plant Disease, No. 960.
Babadoost, M. 2006. Development of internal discoloration of horseradish roots in
commercial fields. pp 5-6. In 2006 Horseradish Res. Rev. & Proc. Horseradish Growers
School.
Babadoost, M., Chen, W., Bratsch, A.D., and Eastman, C.E. 2004. Verticillium
longisporum and Fusarium solani: two new species in the complex of internal
discoloration of horseradish roots. Plant Pathol. 53:669-676.
Babadoost, M. and Islam, S.Z. 2002. Effect of selected fungicides and biocontrol agents
on the incidence of internal discoloration of horseradish root. Phytopathology
(Supplement) 92:S5.
Babadoost, M., Ravanlou, A., and Yu, J. 2010. Hot-Water treatment and use of
fungicides and biofungicides for management of internal discoloration of horseradish
roots in Illinois in 2009. pp 7-12. In 2010 Horseradish Res. Rev. & Proc. Horseradish
Growers School.
Barton, B.H., and Castle, T. 1877. The British flora medica. Page 231. Chatto and
Windus, (ed). London.UK. 447 pp.
Boning, K. 1938. Die wichtigsten krankheiten und schadlinge des meerrettichs. [The
most important diseases and pests of horseradish]. Rev. Appl. Mycol. 18:365.
Bratsch, A. 2006. Specialty crop profile: horseradish. Virginia Cooperative Extension
Publication No. 438-104.
11
Courter, J.W., and Rhodes, A.M. 1969. Historical notes on horseradish. Econ. Bot.
23:156-164.
Dana, B.F., and McWhorter, F.P. 1932. Mosaic disease of horseradish. Phytopathology
22:1000-1001.
De Candolle, A. 1959. Origin of cultivated plants. Hafner Publishing Co., New York. NY.
468 pp.
Dinkelmann, N. 2003. Horseradish in southern Illinois. Belleville Township Highschool
West Belleville, IL. http://www.lib.niu.edu/ipu/ihy000234.html.
Eastburn, D.M., and Chang, R.J. 1994. Verticillium dahliae: A causal agent of root
discoloration of horseradish in Illinois. Plant Dis. 78:496-498.
Eastburn, D.M., and Weinzierl, R. 1995. Horseradish: a guide to major insect and disease
problems. University of Illinois Extension. Report on Plant Disease, No. 944.
Eranthodi, A., and Babadoost, M. 2007. Thermo-therapy for eradication of fungi
pathogens in propagative root-stocks (sets) of horseradish. Phytopathology (supplement)
97 (7): S33.
Fosberg, F. 1965. Nomenclature of the horseradish (Cruciferae). Baileya 13:1-4.
Gerard, J. 1633. The Herbal: or General History of Plants. J. Thomas (ed). Dover
Publications, New York, NY. 1630 pp.
Gerber, J.M., Doll, C.C., Simons, R.K., and Fillingim, K.E. 1983. Internal discoloration
of horseradish-development of symptoms. Univ. Ill. Veg. Res. Rep. & Hortic. Serv.
47:34-37.
Grieve, M. 1967. Horseradish. pp 417-419. In A Modern Herbal, Vol. I.C.F. Leyel (ed).
Hafner Publishing Co., New York, NY. 888 pp.
Heald, F.D., Jonesk, L.K., and Huber, G.A. 1937. Plant Disease Surveys. Washington
Agric. Exp. Sta. Bull. 354.
Hedrick, U.P. 1919. Sturtevant's Notes on Edible Plants. State of New York Dept. Agric.
J.B. Lyon, Albany, NY. 686 pp.
Herold, R. 1957. Zur symptomatik und schadwirkung des kohlschwarzringflecken virus.
J. Phytopathology 55:530-532.
Horseradish information council. 2001. Facts about America's favorite root.
http://www.horseradish.org/facts.html.
12
Horwitz, D.K. 1983. Studies on horseradish (Armoracia rusticana Gaertn. Mey. &
Scherb): Meiosis in fertile and sterile cultivars; seed storage; and methods of assaying for
turnip mosaic virus. M.S. Thesis in Horticulture, Graduate College, University of
Illlinois, Urbana, Illinois.
Horwitz, D., Fletcher, J., and D'Arcy, C. 1985. Turnip mosaic virus in the Illinois
horseradish germ plasm collection. Plant Dis. 69:46-248.
Li, T.B., and Cheo, C.C. 1964. Radish mosaic diseases and experiments on the mixed
infection. J. Plant Protection 3:155-164.
Meyer, M.M. and Milbrath, G.M. 1977. In Vitro propagation of horseradish with leaf
pieces. HortScience 12:544-45.
Mueller, J.P., Percich, J.A., and Mitchell, J.E. 1982. Root deterioration associated with
Verticillium wilt of horseradish. Plant Dis. 66:410-414.
Norton, M., Uchanski, M., Scoggins, K., and Skirvin, R.M. 2001. Tissue culture project
progress. pp 18-20. In 2001 Horseradish Res. Rev. & Proc. Horseradish Growers School.
Percich, J.A., and Johnson, D.R. 1990. A root rot complex of horseradish. Plant Dis.
74:391-393.
Potschke A. 1923. Uber das Scharzwerden des Meerrettichs [On the black discoloration
of horseradish]. Biologischen Reichsanstalt für Land und Forstwirtschaft 11:337-338.
Rhodes, A.M. 1977. Horseradish–problems and research in Illinois. pp 137-147. In Crop
Resources. D.S. Seigler (ed). Academic Press, Inc., New York, NY 233 pp.
Rhodes, A.M., Courter, J.W., and Shurtleff, M.C. 1965. Identification of horseradish
types. Trans. Ill. State Acad. Sci. 58:115-122
Robinson, B.L., and Fernald, M.L. 1908. Gray's New Manual of Botany: A Handbook of
the Flowering Plants and Ferns of the Central and Northeastern United States and
Adjacent Canada. American Book Co. New York, NY. 926 pp.
Shehata, A., Mulwa, R.M.S., Babadoost, M., Uchanski, M., Norton, M.A., Skirvin, R.,
and Walters, S.A. 2009. Horseradish: Botany, Horticulture, Breeding. pp 221-261. In
Horticultural Review. J. Janick (ed). Wiley-Blackwell, Hoboken, NJ. 529 pp.
Simon, J.E., Chadwick, A.F., and Craker, L.E. 1984. Herbs: An Indexed Bibliography,
1971-1980: the scientific literature on selected herbs, and aromatic and medicinal plants
of the temperate zone. Archon Books, Hamden, CT. 770 pp.
13
Stark, C. 1961. Das Auftreten der Verticillium-Tracheomykosen in Hamburger
Gartenbaukulturen. Gartenbauwissenchaft 26:493-528.
Takashi, H., Seiog, I., and Eiji, N. 2002. White rust of horseradish caused by Albugo sp.
Bulletin of Hokkaido Prefectural Ag. Exp. Sta. 82:83-88.
Tucker, A.O., and DeBaggio, D. 2000. The Big Book of Herbs: A Comprehensive
Illustrated Reference to Herbs of Flavor and Fragrance. Intererweave Press, Loveland ,
CO. 672 pp.
Uchanski, M., Skirvin, R.M., and Norton, M.A. 2004. The use of In Vitro thermo-therapy
to obtain Turnip mosaic virus-free horseradish plants. Acta Hortic. 631:175-179.
Uchanski, M., Norton, M.A., and Skrivin, R.M. 2007. The logistic and economics of
producing Pathogen-Free (PF), tissue culture derived horseradish sets. pp 14-19. In 2007
Horseradish Res. Rev. & Proc. Horseradish Growers School.
Weber, W.W. 1949. Seed production in horseradish. J. Hered. 40:223-227.
Yoshii, H., Sugiura, M., and Iwata, T. 1963. Studies on the daikon mosaic virus (DMV),
the Japanese strain of turnip mosaic virus. Proc. Assoc. Plant Protec. Kyushu 1. 26 pp.
14
CHAPTER 2
FUNGAL SPECIES ASSOCIATED WITH INTERNALLY DISCOLORED
HORSERADISH ROOTS
MATERIALS AND METHODS
Root samples
A total of 616 internally discolored horseradish roots (Figure 2-1) including 75,
306, 115, and 120 roots from California, Illinois, Ontario (Canada), and Wisconsin,
respectively, were collected in 2008 and 2009 and tested for isolating fungi (Table 2-1).
The roots from California, Illinois, and Ontario were collected from commercial fields,
while roots from Wisconsin were collected from research plots. The roots were either
tested immediately or were stored at 4 C until they were tested.
Fungal isolation
A segment of 5-cm was cut from the middle of each collected root. The outer
layers (ca 1 mm) of the segments were peeled to remove fungal and bacterial
contaminants on the root surface. The peeled-segments were surface sterilized by dipping
them in a 6% sodium hypochlorite (NaOCl) solution for 1 min, followed by dipping in a
95% ethanol concentration for 3 min, and then they were rinsed in sterile-distilled water
(SDW) three times (1 min each). Each surface-sterilized segment was blotted with
sterilized blotter paper, and five small segments (0.5-cm thick) from each root were cut
15
and placed onto acidified potato dextrose agar (A-PDA) in Petri plates. A-PDA was
prepared by adding 39 g of dehydrated PDA (Difco™, Sparks, MD) to one liter of
distilled water in a conical flask. After mixing it thoroughly, the medium was autoclaved
for 25 min, cooled to 50 C, and then 4 ml of 25% lactic acid was added to the medium
before being poured into Petri plates.
The plates with horseradish root segments were incubated at 22-24 C with 12 h
light/12 h darkness. The incubated plates were observed for growth of fungal colonies 5,
10, and 15 days after plating the segments. Plaques of culture medium (ca 2mm in
diameter) with hyphal tips of the developing fungal colonies were cut and transferred
onto PDA for purification.
Culture media
Purified fungal colonies were transferred onto four different culture media,
including: (1) V8 juice agar (V8A) containing 150 μg/ml rifampicin; (2) antibiotic-PDA
(AB-PDA), with 100 μg/ml of neomycin and 10 μg/ml of streptomycin sulfate; (3)
Verticillium selective medium (VSM); and (4) Synthetischer Nährstoffarmer agar (SNA).
V8A was used for spolurating fungi and studying their reproductive bodies; AB-PDA
was used for studying colony patterns; VSM was used for screening Verticillium spp.;
and SNA was used for studying morphological features of Fusarium spp.

V8 Juice agar (V8A)
16
V8A was prepared by adding 200 ml of V8-juice and 2 g of CaCO3 to 100 ml of
distilled water, passing the suspension through four-layer cheesecloth and facial
tissue into a conical flask, adding 20 g agar, and then adding distilled water to
bring the volume up to one liter. The mixture was heated to boil and then was
autoclaved at 15 psi for 25 min. The culture medium was cooled to approximately
50 C, and 150 μg/ml of rifampicin dissolved in 100 % ethanol was added to the
medium and poured into Petri plates.

Antibiotic potato dextrose agar (AB-PDA)
AB-PDA was prepared by adding 39 g of dehydrated PDA (Difco™, Sparks,
MD) to one liter of distilled water and dissolved by heating. Then, the mixture
was autoclaved at 15 psi for 25 minutes. After cooling the medium to
approximately 50 C, 100 μg/ml of neomycin and 10 μg/ml of streptomycin
sulfate were added and poured into the Petri plates.

Verticillium selective culture medium (VSM)
VSM was prepared by adding 7.6 g of sucrose, 2 g of NaNO3, 0.5 g of KCl, 0.5 g
of MgSO4*7 H2O, 1.0 g of K2HPO4, 0.01 g of FeSO4, 5 ml of 95% ethanol, 0.05 g
of PCNB, and 20 g of agar to distilled water in a conical flask and then bringing
the volume up to one liter. The mixture was first boiled to dissolve the
compounds and then was autoclaved at 15 psi for 25 min. After cooling the
culture medium to approximately 50 C, 100 mg of streptomycin sulfate and 250
mg chloramphenicol were added to the medium prior to pouring into the Petri
plates.

Synthetischer Nährstoffarmer agar (SNA)
17
SNA was prepared by adding 1 g of KH2PO4, 1 g of KNO3, 0.5 g of MgSO4*7
H2O, 0.5 g of KCL, 0.2 g of sucrose, 0.2 g of glucose, and 20 g of agar to distilled
water in a conical flask, and then bringing the volume to one liter by adding
distilled water. The mixture was first boiled to dissolve the compounds, then was
autoclaved at 15 psi for 25 min, cooled down to approximated 50 C and then
poured into the Petri plates.
Morphological identification
Cultures were examined periodically and identified when they sporulated. The
cultures were separated into groups based on their morphological characteristics
including growth pattern, colony texture, pigmentation, and growth rate of the colonies
on PDA (Promputtha et al., 2005). When fungal colonies sporulated on V8A and PDA,
small plaques from the edge and the center of each growing colony were transferred onto
glass slides, and then were examined using a compound light microscope (Olympus
BX41 system microscope, Olympus America Inc., Melville, NY) for characteristics of
their vegetative and reproductive structures such as hyphal color and structures, shape
and size of conidia, conidiophores, and microsclerotia. Isolates of Verticillium were
grown and screened on VSM in Petri plates at 22-24 C with 12 h light/12 h darkness for
7-10 days.
All selected Verticillium isolates were grown on PDA at 22-24 C with 12 h
light/12 h darkness for 7-10 days and identified according to the procedures reported by
Babadoost et al. (2004), Hawksworth and Talboys (1970 a and b), and Karapapa et al.
18
(1997). Texture and color of colony and morphology of conidiopohores, conidia, and
microsclerotia produced on PDA were evaluated, and the length and width of 15 conidia
of each Verticillium isolate were measured.
Fusarium isolates were grown on PDA and SNA in Petri plates at 22-24C with
12 h light/12 h darkness for 10 days. Species of the Fusarium isolates were identified
based on their color and growth pattern of colonies on PDA and morphology of macroand micro-conidia and chlamydospores on SNA according to the guidelines reported by
Nelson and Toussoun (1983), Skovgaard et al. (2003) and Leslie et al. (2006).
DNA extraction
Based on the morphological characteristics, the isolates were separated into 20, 21,
15 and 13 groups from California, Illinois, Ontario, and Wisconsin, respectively. Five
representative isolates from each group (or all of them with fewer than five isolates) were
used in sequence analysis.
A small plaque of each isolate was transferred into 150 ml potato dextrose broth
(PDB) in a 250 ml flask, and the flask was placed onto a shaking incubator (150 rpm) at
25 C for 7-10 days. PDB was prepared by adding 24 g of dehydrated PDB (Difco™,
Sparks, MD) to one liter of distilled water in a conical flask. After mixing it thoroughly,
the medium was autoclaved for 25 min, cooled down to room temperature (22 C), and
then inoculated with fungal hyphae. Fungal mycelia were recovered from the PDB
culture by centrifugation (12,000 x g) and filtration with sterilized filter paper.
19
Total DNA was extracted from fungal mycelia using the Fast DNA Spin kit (MP
Bio., Solon, OH), according to the manufacturer‟s recommended protocol. The amount of
DNA was determined by comparing with pre-calculated λ-DNA (Promega Corp.,
Madison, WI) and 1-kb molecular weight marker by electrophoresis through 1% agarose
gel in 1× Tris-acetate EDTA (TAE) buffer stained with ethidium bromide. The agarose
gel was visualized under UV light and documented after electrophoresis. The DNA was
stored at -20 C until used.
Polymerase chain reaction (PCR) amplification
For identification of the fungal isolates, DNA of each isolate was amplified by
polymerase chain reaction (PCR) using two different Internal Transcribed Spacer (ITS)
primers sets (ITS 5 and ITS 4, Table 2-2). A 50-µl PCR reaction mixture was prepared,
which contained final concentrations of 10x PCR buffer, 2.0 mM MgCl2, 2.5 mM dNTPs,
10 µM of each primers, 2 units of Taq polymerase (Promega Corp., Madison, WI), and 2
µl of DNA extract. The PCR amplifications were conducted using a thermal cycler (PCT200, MJ Research Inc., Waltham, MA). The PCR cycles included initial denaturing for 2
min at 95 °C; 35 cycles of denaturation at 94 °C for 45 sec, annealing at 54 °C for 1 min
and extension at 72 °C for 1.5 min; followed by final extension at 72 °C for 5 min.
For phylogenetic identification of Verticillium species, PCR was performed to
amplify the Internal Transcribed Spacer (ITS) ITS1-5.8s-ITS2 region using primer ITS5
and ITS 4 (White et al., 1990), two mitochondrial genes of cytochrome oxidase subunit
20
III (cox3) using cox3A and cox3B (Kouvelis et al., 2004; Pantou et al., 2005), and NADH
dehydrogenase subunit I (nad1) using nad1A and nad1B (Kouvelis et al., 2004; Pantou et
al., 2005) (Table 2-2). The PCR reactions were conducted in a PCT-200 thermal cycler.
A total volume of 50 µl PCR reaction mixture was prepared, which contained 10x PCR
buffer, 2.0 mM MgCl2, 2.5 mM dNTPs, 10 µM of each primers, 30 ng of Taq polymerase
(Promega Corp., Madison, WI), and 2 µl of DNA extract. PCR conditions for ITS were
the same as above. For cox3 and nad1, PCR conditions were initial denaturing at 94 °C
for 3 min; 30 cycles of denaturing at 94 °C for 1 min, annealing at 48 °C for cox3 primers
and 50 °C for nad1 primers for 1 min, and extension at 74 °C for 2 min; and final
extension at 74 °C for 5 min.
For phylogenetic identification of Fusarium species, amplification of the ITS,
mitochondrial small subunit rDNA (mtSSU), and translation elongation factor 1-alpha
(EF1-alpha) was accomplished by using primers ITS5 and ITS4 (White et al., 1990),
NSM1 and NSM2 (Li et al., 1994), and EF-1H and EF-2T (O‟Donnell et al., 1998),
respectively (Table 2-2). Total DNA from each isolate was subjected to PCR in a PTC200 thermal cycler. PCR conditions for ITS region were the same as the described above.
For mtSSU, PCR conditions were initial denaturing at 94 °C for 3min; 35 cycles of
denaturing at 94 °C for 35 sec, annealing at 52 °C for 55 sec, extension at 72 °C for 2
min; and final extension at 72 °C for 5 min. For EF1-alpha, PCR conditions were 30
cycles of DNA denaturing at 92 °C for 1 min, annealing at 55 °C for 1 min, and extension
at 72 °C for 1 min. Initial denaturing at 94 °C was extended to 2 min and the final
extension was at 72 °C for 5 min.
21
All PCR amplifications were verified by electrophoresis with 10 μl of each PCR
product in a 1% agarose gel with 1X TAE buffer stained with ethidium bromide. The gels
were visualized and documented under UV light. The lengths of the amplification
products were estimated by comparing with a 100-bp DNA ladder.
DNA sequencing and data analysis
PCR products were purified using the Wizard SV gel and PCR Clean-Up system
(Promega Corp., Madison, WI), according to the manufacturer‟s protocol. Purified PCR
products were sequenced in an automated sequencer at the Core DNA Sequencing
Facility of the University of Illinois at Urbana-Champaign (UIUC). Sequencing was
performed using the same forward and reverse primer sets which were used for PCR
amplification (Table 2-2).
Partial forward sequences were used as query sequences to find similar sequences
in NCBI GenBank (http://www.ncbi.nlm.nih.gov/) for all of the fungal isolates. The most
similar reference sequences with the query sequences were obtained and used to identify
the genus or species of the fungus. The isolates were assigned to a species if the sequence
was ≥ 99% similar to a valid species sequence deposited with NCBI GenBank, and to a
genus if the species identity was not conclusive, but the similarity was ≥ 97% (Thomas et
al., 2008). The sequences obtained from the primer sets (Table 2-3) of Fusarium and
Verticillium were compared to the reference sequences available in GenBank using the
Blast search.
22
Phylogenetic analysis
The sequence results were first edited using Clustal X (Thompson et al., 1997)
and BioEdit version 7.0.9 (Hall, 1999), and then the alignment was refined using
PHYDIT version 3.2 (Chun, 1995; http://plaza.snu.ac.kr/~jchun/phydit/). The edited
sequences were compared with the reference sequences from GenBank. Maximum
parsimony (MP) analysis was estimated using heuristic searches consisting of random
addition order and tree bisection-reconnection (TBR) branch swapping. Bootstrap
analysis using 1000 replications was performed to assess the relative stability of the
branches.
Pathogenecity tests
Pathogenecity tests were conducted using root-dip inoculation methods, which
were reported by Babadoost et al. (2004) and Eastburn & Chang (1994). In this
experiment, two Verticillium species [V. dahliae (group A and group B) and V.
longisporum] and seven Fusarium species (F. acuminatum, F. commune, F. equiseti-like
species, F. oxysporum, F. proliferatum and F. solani) were tested for their pathogenicity
using two horseradish cultivars 1573 and Big Top Western (BTW), which are susceptible
to internal root discoloration (Babadoost et al., 2004).
Asymptomatic horseradish sets, obtained from commercial growers, were used in
the pathogenicity test. To eliminate possible set-borne inoculum, the sets were treated at
47 °C for 20 min (Babadoost et al., 2007) prior to inoculation. The sets were then dipped
23
in 70% ethanol for 25 sec, soaked in 0.6 % NaOCl for 5 min, and rinsed in tap water.
Segments were then planted in 50 x 35 x 10 cm pots containing a pasteurized mix of
soil:peat:perlite (1:1:1) (Babadoost et al., 2004; Eastburn and Chang, 1994), and were
grown in a greenhouse at 20-26 C. After 30 days, plants were removed from the pots,
soil was gently cleaned from roots, and roots were inoculated with Verticillium and
Fusarium isolates as described below. One isolate from each species was used in the
pathogenicity test.
For producing Verticillium inoculum, two-week-old Vertciliium species cultures
on PDA in Petri plates were macerated with 15 ml of SDW and the conidia were
dislodged using a soft brush. One milliliter of the conidial suspension was spread onto the
surface of PDA in each Petri plate. The inoculated plates were incubated at 22-24 C
under 12 hr fluorescent light/ 12 hr darkness for 3 weeks. The cultures were then blended
(Laboratory Blender 51BL30, Waring Comm., Torrington, CT) with 200 ml DW at high
speed for 30 sec. The suspension was passed through a series of 500-, 180-, and 38-µm
sieves. The material (microsclerotia) caught on the 38-µm sieve was rinsed with DW, and
collected material was resuspend in 0.5% sterilized agar solution to prepare a sticky
inoculum (Babadoost et al. 2004). An inoculum suspension of 500 microsclerotia per ml
was prepared using a spore-counting chamber.
For producing Fusarium inoculum, a conidial suspension was prepared from 7- to
10-day-old Fusarium cultures on PDA plates by adding 10 ml SDW to each plate and
dislodging the conidia using a soft brush. The concentration of conidia (micro- and
24
macro-conidia) was adjusted to 105 conidia per 1 ml using a hemocytometer (Babadoost
et al., 2004). To prepare the sticky inoculum suspension, Fusarium inoculum was
prepared in a sterilized 0.5% agar solution (Atibalentja and Eastburn, 1998; Babadoost et
al., 2004).
Roots of plants grown in the greenhouse were dipped in the 0.5% agar suspension
of microsclerotia (for Verticillium) and conidia (for Fusarium) for 15 sec, and then
inoculated plants were planted in plastic pots containing the pasteurized soil:peat:perlite
(1:1:1) mix. Roots of control plants were dipped in sterilized 0.5 % agar solution. Pots
were placed in the greenhouse at 20-26 C under a combination of 1,000 W high pressure
sodium and mercury vapor lamps with a 12 h per day photoperiod. Each inoculation
treatment included four replications, each with six plants. The inoculation experiments
were repeated once.
Plants were removed at four weeks intervals until four month after inoculation.
Roots were washed, cut, and evaluated for severity of internal discoloration at the cross
section at the middle of the root. Severity of discoloration was rated in their medium
discolored percentage ranges as 0 = no discoloration; 5 = 1-10; 18 = 11-25; 38 = 26-50;
and 75.5 = 51-100% discoloration of vascular (Babadoost et al., 2004). Root sections
were surface sterilized by soaking in 6% NaOCl solution for 1 min, dipping in 95%
ethanol for 2 min, and rinsing in SDW three times. Root sections were then placed on APDA in Petri plates for isolation of the pathogens. Plates were incubated at 22-24 C
25
under 12 hr fluorescent light/ 12 hr darkness. Growing fungal colonies were transferred
onto PDA and the species were identified based on their morphological characteristics.
Statistical analyses were performed using SAS (Version 9.2, SAS Institute. Cary,
NC). Disease incidence and severity of the roots were analyzed using ANOVA
procedures.
RESULTS
Identification of fungal isolates
Fungi were isolated from 528 roots from a total of 616 roots (86%), which
included 65 of 75 (87%), 250 of 306 (82%), 106 of 115 (92%), and 107 of 120 (89%) of
roots from California, Illinois, Ontario (Canada), and Wisconsin, respectively (Table 21). A total of 742 fungal isolates were collected, which were comprised 106, 328, 192,
and 116 isolates from California, Illinois, Ontario, and Wisconsin, respectively (Table 21).
Based on partial ITS sequence analysis, 18 genera of fungi, including Alternaria,
Arthopyreniaceae, Chaetomium, Colletotrichum, Cordyceps, Cylindrocarpon, Fusarium,
Neonectria, Penicillium, Phoma, Phomopsis, Pyrenochaeta, Rhizoctonia, Rhizopus,
Rhizopycnis, Trichurus, Ulocladium and Verticillium were identified (Table 2-3, and
Figure 2-2). The most commonly isolated genera were Verticillium and Fusarium.
Verticillium species were isolated from 34, 45, 43, and 10% of roots collected from
California, Illinois, Ontario, and Wisconsin, respectively. Similarly, Fusarium species
26
were isolated from 26, 31, 14, and 65% of roots collected from California, Illinois,
Ontario, and Wisconsin, respectively (Table 2-3, and Figure 2-3).
Morphological characterization of Verticillium species
All Verticillium isolates developed verticillate conidiophores from globes,
swollen, primary cells and later they formed microsclerotia as reported by Karapapa et al.
(1997). One group (group VD) of isolates with spherical and more compact
microsclerotia (Figure 2-4) fit the characteristics of V. dahliae (Karapapa et al., 1997). A
total of 210 of 277 (76%) Verticillium isolates (24% of total fungal isolates) were
identified as V. dahliae, which included 0, 83, 93, and 100% of Verticillium isolates from
California, Illinois, Ontario, and Wisconsin, respectively (Table 2-4). Another group
(group VL) of Verticillium isolates with elongate and irregular-shaped microsclerotia
(Figure 2-5) fit the characteristics of V. longisporum (Karapapa et al., 1997). A total 67 of
277 (14%) Verticillium isolates were identified as V. longisporum, which included 100,
17, 7, and 0% of Verticillium isolates from California, Illinois, Ontario, and Wisconsin,
respectively (Table 2-4). V. dahliae isolates produced conidiophores with 4-5 phialides
(Figure 2-6), while V. longisporum isolates produced conidiophores with 2-3 phialides
(Figure 2-7) (Karapapa et al., 1997). Also, isolates of V. dahliae produced conidia with
spore length ≤ 5.5 µm (mean length 4.4± 0.23 μm), while V. longisporum formed conidia
measuring ≥ 7 µm (mean length 7.9 ±0.08 μm) (Babadoost et al., 2004; Hawksworth and
Talboys, 1970b; Karapapa et al., 1997) (Figure 2-8 and 2-9).
Phylogenetic analysis of Verticillium species
27
Amplification of the ITS1-5.8S-ITS2 region of Verticillium isolates generated
532-534 bp fragments and alignment of the sequences resulted in a dataset of 637
characters. Based on 90 parsimony-informative characters, 1,000 most parsimonious trees
were generated with tree length of 313 steps (CI=0.8179, RI=0.7957) (Figure 2-10). In
the maximum parsimony analysis, the Verticillium isolates from horseradish were divided
into two clades. Ten isolates of Verticillium including SDV1025 referred to as group VD
based on morphological characteristics and five isolates of Verticillium including
TBV3014 assigned to group VL based on morphological characteristics clustered
together in a clade along with strains of V. dahliae (M5, UZ132, and T9) and V.
longisporum (G18). Five isolates of Verticillium including CBV2034 assigned to group
VL based on morphological characteristics clustered together in a clade along with strains
of V. dahliae (Vd292) and V. longisporum (K2 and K12) (Figure 2-10). V. dahliae and V.
longisporum could not be differentiated by sequences of this gene.
The cytochrome oxidase subunit III (cox3) of all isolates of Verticillium were
amplified with primers cox3A-cox3B and the sequences from PCR products were
combined with retrieved sequences from GenBank. Amplification of the cox3 gene
generated 408-413 bp fragments, and alignment of the sequences resulted in a dataset of
376 characters. Based on 86 parsimony-informative characters, 15 most parsimonious
trees were generated with tree length of 249 steps (CI=0.6466, RI=0.6318) (Figure 2-11).
In the maximum parsimony analysis, the isolates from horseradish were divided into two
clades, V. dahliae clade and V. longisporum clade. Ten isolates of Verticillium including
SDV1025 referred to as group VD based on morphological characteristics and five
28
isolates of Verticillium including CBV2034 referred to as group VL based on
morphological characteristics were clustered together in the V. dahliae clade along with V.
dahliae strains M5, UZ132, and T9, which was supported by a bootstrap value of 60%.
Five isolates of Verticillium including TBV3014 assigned to group VL and V.
longisporum strains G19, K2, and K12 were clustered together in a V. longisporum clade
(Figure 2-11). V. dahliae group differed only at two positions from V. longisporum group.
The NADH dehydrogenase subunit 1 (nad1) of all isolates of Verticillium was
amplified with primers nad1A-nad1B, and the sequences from PCR products were
combined with retrieved sequences from GenBank. Amplification of the nad1 gene
generated 536-542 bp fragments and alignment of the sequences resulted in a dataset of
523 characters. Based on 120 parsimony-informative characters, 4 most parsimonious
trees were generated with a tree length of 385 steps (CI=0.7143, RI=0.6927) (Figure 212). In the maximum parsimony analysis, the Verticillium isolates from horseradish were
divided into three clades, two V. dahliae clades and one V. longisporum clade. Ten
isolates of Verticillium including SDV1025 referred to as group VD based on
morphological characteristics and V. dahliae strain UZ132, M5 were clustered together in
a clade, which was supported by a bootstrap value of 64%. Five isolates of Verticillium
including CBV2034 belong to morphological group VL and V. dahliae strain T9 were
clustered together in a clade, and five other isolates of Verticillium including TBV3014
belong to morphological group VL and V. longisporum strains G19, K2, and K12 were
clustered together in a V. longisporum clade which was supported by a bootstrap value of
68% (Figure 2-12).
29
Phylogenetic analysis based on combined sequence of cox3 and nad1 yielded
three most parsimonious tree (steps=637, CI=0.6845, RI=0.6683). Parsimony analysis of
the combined dataset revealed a tree with similar topology as that revealed in
independent analyses of the cox3 and nad1 data. The tree topology distinctively
discriminated V. dahliae and V. longisporum, and the V. dahliae clade was divided into
two subclades, V. dahliae type A (morphological group VD) and type B (morphological
group VL) (Figure 2-13). As in the nad1 tree, V. dahliae type A included ten isolates of
Verticillium and V. dahliae strains M5 and UZ132. The V. dahliae type B included five
isolates of Verticillium and V. dahliae strain T9. The two subgroups formed the
monophyletic clade of V. dahliae supported by moderate bootstrap values (68%). The V.
longisporum clade included five isolates of Verticillium assigned to morphological group
VL and V. longisporum strains G19, K2, and K12. All isolates of V. longisporum formed
a monophyletic clade with strong bootstrap support (83%) (Figure 2-13).
Based on morphological and molecular characteristics, the isolates of Verticillium
associated with internally discolored horseradish roots were identified as two species of
Verticillium, V. dahliae (type A and B) and V. longisporum. The V. dahliae type A which
has morphological characteristics of typical V. dahliae was the most prevalent in root
samples from Illinois, Ontario, and Wisconsin; however, V. longisporum was the most
prevalent in roots from California. V. dahliae type B which has morphological
characteristics of V. longisporum was isolated only from California, Illinois and Ontario.
30
Morphological characterization of Fusarium species
Based on morphological characteristics, Fusarium isolates were divided into six
species: F. acuminatum, F. commune, F. equiseti, F. oxysporum, F. proliferatum, and F.
solani.
The first group (group FA) of isolates grew slower than other Fusarium groups.
The colonies were red to burgundy with abundant mycelium on PDA. Sporodochia
formed in the center of the colony and were pale to dark brown. Macroconidia were
moderately curved with a, distinct foot-shaped basal cell and a relatively elongated apical
cell, and 3-5 septate (Figure 2-14-1). The isolates in this group fit the description of F.
acuminatum.
The second group (group FC) of isolates produced whitish mycelium on PDA,
and grayish yellow to violet pigment in the agar. Macroconidia were cylindrical, straight
to slightly curved, with a slightly curved apical cell and foot-shaped basal cell that bent
equally at both ends, and 3-5 septate (Figure 2-14-2). The isolates in this group fit the
description of F. commune.
The third group (group FE) of isolates produced abundant mycelium that were
initially white, but became brown with age on PDA. Sporodochia were orange and had
long and slender-shape macroconidia that were sickle-shaped with a distinctive curvature,
with whip-like elongated apical cell, and prominent foot-shaped basal cell (Figure 2-143). The isolates in this group fit the description of F. equiseti.
31
The fourth group (group FO) of isolates produced pale purple pigments in the
agar with abundant whitish mycelium on PDA. Macroconidia were abundant, only
slightly sickle-shaped, with an attenuated apical cell and a foot-shaped basal cell, and 3
septate. Microconidia were abundant, single-celled, oval or kidney-shaped (Figure 2-144). This group of isolates fit the description of F. oxysporum.
The fifth group (group FP) of isolates had abundant mycelium with white colonies
on PDA, which became purple-violet with age. The isolates produced tan to pale orange
sporodochia. Macroconidia were slightly sickle-shaped to almost straight, with a footshaped basal cell and curved apical cell, and 3-5 sepatate. Microconidia were club-shaped
and single-celled (Figure 2-14-5). This group of isolates fit the description of F.
proliferatum.
The sixth group (group FS) of isolates produced white to pale, sparse mycelium,
and sporodochia were produced in abundance with cream to yellow color on PDA.
Macroconidia were abundant, relatively wide, stout, slightly curved, with a rounded
apical cell and a rounded or foot-shaped basal cell, and 5-7 septate. Microconidia were
oval to kidney-shaped, generally single celled and sometimes one with septum (Figure 214-6). The isolates in this group fit the description of F. solani.
Molecular identification of Fusarium species
32
The ITS region of all isolates of Fusarium were amplified with primers ITS5ITS4, and the sequences from PCR products were combined with retrieved sequences
from GenBank. Amplification of the ITS region generated 521-569 bp fragments, and
alignment of the sequences resulted in a dataset of 429 characters. Based on 80
parsimony-informative characters, 696 most parsimonious trees were generated with a
tree length of 163 steps (CI=0.7914, RI=0.9470) (Figure 2-15). In the maximum
parsimony analysis, three isolates of Fusarium including SD5115 and F. equiseti
(AB425996) were clustered together in a monophyletic clade which was supported by
bootstrap value 67%, however sequences of three Fusarium isolates and F. equiseti have
2-5 base pairs differences (Figure 2-15). Four isolates of Fusarium including SD3152 and
F. oxysporum were clustered together in a same monophyletic clade of F. oxysporum
complex. Eight Fusarium isolates including SD1184, F. solani S-0900 and Nectaria
haematococca were clustered together in a clade which divided into three sub-clades of F.
solani complex. Three isolates of Fusarium including SD2018, and F. acuminatum
IBE000017 were clustered together in a clade which was supported by a bootstrap value
of 80%, and three Fusarium isolates including SD4139, and F. commune 9245H were
clustered together in a clade with identical sequence. Isolate WA6005 and F.
proliferatum 9223F were clustered together in a clade which was supported by a
bootstrap value of 68%.
The mitochondrial small subunit rDNA (mtssu rDNA) regions of all Fusarium
isolates were amplified with primers NSM1 and NSM2, and the sequences from PCR
products were combined with retrieved sequences from GenBank. Amplification of the
33
mtssu rDNA region generated 690-730 bp fragments and alignment of the sequences
resulted in a dataset of 562 characters. Based on 59 parsimony-informative characters,
four most parsimonious trees were generated with a tree length of 168 steps (CI=0.9167,
RI=0.9381) (Figure 2-16). In the maximum parsimony analysis, the Fusarium isolates
from horseradish were divided into six clades. Three isolates of Fusarium including
SD4139 were clustered together in a clade with F. commune 9245H and F. rodolens
NRRL28058, which was supported by a high bootstrap value of 91%. Fusarium isolates
WA6005 was located on a same clade with F. proliferatum 31X4 with 100% bootstrap
value. Also four isolates of Fusarium including SD3152 and F. oxysporum f. sp. melonis
were grouped together in a same clade which was supported by a 94% bootstrap value.
Three isolates of Fusarium including SD2018, F. acuminatum NZ50 and F. tricinctum
NZ62 were clustered together in a clade with 69% bootstrap support, and eight isolates of
Fusarium including SD1212 were clustered together with F. solani NRRL22382 which
was supported by 98% bootstrap value. Three isolates of Fusarium including SD5115
were clustered together in a clade, but there was no matched reference sequence in the
Genbank database.
The translation elongation factor 1-alpha region (EF 1-alpha) of all isolates of
Fusarium was amplified with primers EF-1H and EF-1T, and sequences from PCR
products were combined with the retrieved sequences from GenBank. Amplification of
the EF 1-alpha region of all isolates generated 664-708 bp fragments and alignment of the
sequences resulted in a dataset of 380 characters. Based on 116 parsimony-informative
characters, 59 most parsimonious trees were generated with a tree length of 344 steps
34
(CI=0.8052, RI=0.9193) (Figure 2-17). In the maximum parsimony analysis, Fusarium
isolates from horseradish were divided into seven different clades with F. acuminatum R6678, F. commune NRRL28387, F. equiseti DAOM236361, F. oxysporum f. sp. melonis
TX388, F. proliferatum NRRL43666, and F. solani complex of NRRL22402,
NRRL20547 and N. haematococca MAFF840047 (Figure 2-17). All of morphologically
identified Fusarium isolates were clustered together and located in the monophyletic
clade with their reference strains. Isolates of Fusarium including SD5115 and F. equiseti
DAOM236361 were clustered together, but they had 2-5 difference sequences.
Phylogenetic analysis based on combined sequences of ITS, NMS, and EF1-alpha
yielded eighteen most parsimonious trees. The length of the tree was 715 steps
(CI=0.8028, RI=0.9126) and the number of parsimony informative characters was 1422.
Parsimony analysis of the combined dataset revealed a tree with similar topology as that
revealed in independent analyses of the ITS, NMS, and EF1-alpha data (Figure 2-18).
The phylogenetic tree revealed six clades corresponding to the six reference strains of
Fusarium used in the investigation. Fusarium isolates clustered in the same clade along
with their reference strains of F. acuminatum, F. commune, F. oxysporum, F.
proliferatum, and F. solani. However, isolates of SD5115, SD5030 and WA5039 which
have morphological characteristics of F. equiseti were not clustered with any of the
Fusarium species.
Based on the morphological characteristics and molecular analysis of the
Fusarium isolates, six species were identified, which were F. acuminatum, F. commune,
35
F. equiseti-like species, F. oxysporum, F. proliferatum, and F. solani. F. acuminatum, F.
oxysporum, and F. solani were isolated from samples from California, Illinois, Ontario,
and Wisconsin. Two most parsimonious trees showed that the group of isolates that had
the same morphological description of F. equiseti was clustered together, but these
isolates had 2-5 sequence changes compared to the reference sequence of F. equiseti in
Genbank. F. acuminatum, F. oxysporum, and F. solani were isolated from horseradish
roots collected from all four horseradish growing areas (California, Illinois, Ontario, and
Wisconsin) (Table 2-5). F. commune was isolated from roots collected from Illinois and
Wisconsin. F. equiseti–like species was isolated only from roots from Illinois, and F.
proliferatum was isolated only from roots collected from Wisconsin (Table 2-5).
Pathogenicity test
Verticillium spp.
Symptoms of the internal discoloration of roots developed over a period of four
months in the inoculated plants (Table 2-6). There was no significant difference in the
incidence (percentage of root discolored) or severity (percentage of root area discolored
at cross section) of root discoloration between two horseradish cultivars „BTW‟ and
„1573‟. Also, there were no significant differences in the incidence or severity of internal
discoloration of roots inoculated with V. dahliae and V. longisporum. Initial discoloration
symptoms were observed one or two months after inoculation of roots with Verticillium
isolates. Both incidence and severity of the discoloration of the roots gradually increased
over the four month period. After four months from inoculation, 83-100% of the
36
inoculated roots exhibited symptoms of internal discoloration. The same Verticillium
species used to inoculate in roots was reisolated from the discolored roots.
Fusarium spp.
In the Fusarium trial, plants inoculated with F. acuminatum, F. commune, F.
oxysporum, and F. solani developed symptoms of internal discoloration of roots. Internal
discoloration was observed one month after inoculation of roots. Four months after
inoculation, more than 83% of the plants had internally discolored root. In most of the
pathogenicity tests, significantly higher percentages of roots were discolored four months
after inoculation compared to those after one month from inoculation (Table 2-7). Roots
inoculated with F. commune also started rotting two months after inoculation, and all
roots were completely rotted four months after inoculation.
Less than 25% of plants inoculated with either F. equiseti-like species or F.
proliferatum exhibited internal discoloration in the roots, with very low discoloration
severity ≤ 0.5% (Table 2-7). Both fungi were isolated from the discolored roots. Also,
both of the fungi were isolated from inoculated asymptomatic roots, indicating that the
fungi entered roots but did not induce discoloration in most of the root.
DISCUSSION
Although all of the 616 collected roots showed symptoms of internal root
discoloration, no fungal colony grew out from 88 (14%) of the roots. Some of the
cultured root segments with no fungal colony were from severely discolored roots
37
(severity of discoloration > 50%). Colonies of bacteria such as Pseudomonas and
Bacillus species developed on the cultured root segments with no fungal colony. It is
believed that the invasion of internally discolored roots by Pseudomonas and Bacillus
species resulted in a suppression of fungal growth. In this study, Pseudomonas and
Bacillus isolates collected from internally discolored horseradish roots strongly
suppressed growth of Verticillium and Fusarium species in vitro (data presented in
chapter 3).
Eastburn and Chang (1994) reported V. dahliae as the causal agent of the internal
discoloration of horseradish roots in Illinois. Percich and Johnson (1990) reported that, in
addition to V. dahliae, F. roseum ‘acuminatum’ caused internal discoloration in
horseradish roots in Wisconsin. Babadoost et al. (2004) reported that the internal
discoloration of horseradish root is a disease complex, which is caused by at least three
fungi, V. dahliae, V. longisporum, and F. solani. The result of this study expands on the
findings of Babadoost et al. (2004) and Percich and Johnson (1990) that the internal
discoloration of horseradish root is a disease complex that is caused by V. dahliae, V.
longisporum, F. acuminatum, F. commune, F. oxysporum, and F. solani. This is the first
report of F. acuminatum and F. oxysporum as causal agents of internal root discoloration
of horseradish roots. Also, this is the first report of F. commune causing horseradish root
rot.
This study was the first investigation to identify fungi associated with internally
discolored horseradish roots using molecular methods. Although careful examination of
38
morphological characteristics provided information for identification of fungi associated
with horseradish roots, using molecular methods was helpful in determining accuracy of
identification of fungi at the species level. The results showed that accurate identification
of Verticillium species without using molecular methods may not be possible.
V. longisporum is identified from V. dahliae by its longer conidia (≥7 μm vs. ≤
5.5 µm) and irregular-shaped microsclerotia (Babadoost et al., 2004; Karapapa et al.,
1997). However, molecular analysis appears to be necessary to accurately differentiate
these two species from each other. The nuclear ribosomal DNA (rDNA) has been
commonly used for phylogenetic studies of fungi because it is multi-copied and contains
highly conserved and universally-present coding genes (Bruns et al., 1991; Butler and
Metzenberg, 1989; Pantaou et al., 2003). However, phylogenetic analysis using rDNA
gene sequences along with morphological characteristics may not lead to accurate
identification of fungal species (Berbee and Taylor, 2001). Some reports (Carder and
Barbara, 1999; Pantaou et al., 2005) show that identifying species of Verticillium, based
on morphological characteristics alone, or in combination with analysis of sequences of
single gene of rDNA, may lead to misidentification of species. Therefore, analysis of
mtDNA sequences has become an alternative to using rDNA sequences for phylogenetic
studies (Bullerwell et al., 2003; Kouvelis et al., 2004). In this study, using morphological
characteristics along with analysis of sequences of ITS region and mtDNA genes (nad1
and cox3) helped to accurately separate Verticillium isolates into their species.
39
The results of this study showed that the causal agents of root discoloration were
different in different horseradish growing areas. For example, V. longisporum was the
dominant species in roots from California, while V. dahliae was isolated from most of the
roots collected from Illinois, Ontario, and Wisconsin. Similarly, F. solani was more
prevalent in roots from California and Illinois, while F. acuminatum was isolated from
most of the roots from Ontario and Wisconsin. It is not clear whether the differences in
occurrence of pathogens of internal discoloration of horseradish roots are related to the
climate or horseradish cultivars. However, since the greenhouse studies showed that
inoculation of roots of both cultivars (BTW and 1573) resulted in root discoloration, it is
likely that occurrences of different pathogens in different horseradish growing areas is
related to the climate rather than horseradish cultivars. Further studies are needed to
precisely elaborate interactions among Verticillium and Fusarium species and horseradish
cultivars in different climates.
Based on the morphological characteristics of conidia and colony patterns,
Fusarium isolates were separated into six species, F. acuminatum, F. commune, F.
equiseti, F. oxysporum, F. proliferatum, and F. solani. Analysis of ITS sequences
confirmed identification of Fusarium species based on morphological characteristics of
conidial and colony characteristics. However, analysis of the sequences resulting from
using translational elongation factor 1-alpha (EF1-alpha) and mitochondrial small subunit
rDNA (mtssu rDNA) regions (Geiser et al., 2004; Yli-Mattila et al., 2002) showed that F.
commune and F. oxysporum are more closely related. Also, analyses of the sequences
resulting from using EF1-alpha and mtssu rDNA gene revealed that isolates identified as
40
F. equiseti had 2-5 bases different than the reference sequence of F. equiseti
(DAOM236361), even though those isolates were fit with morphological description of F.
euqiseti Thus, the isolates are classified as F. equiseti-like species. Additional
investigations are needed to clarify the classification of these isolates, which may confirm
the existence of a new Fusarium species in horseradish roots.
In addition to Verticillium and Fusarium species, fungi in at least 16 genera were
isolated from internally discolored roots. Since the occurrence of these fungi in the roots
was very low, it is believed that these fungi are secondary colonizers of the roots infected
by Verticillium and/or Fusarium species. No pathogenicity test of fungal species other
than Verticillium and Fusarium species was conducted. Among the identified fungi,
Alternaria occurs on horseradish plants, causing leaf spot (Laemmlen, 2001).
Sixty-two (8.4%) of the fungal isolates, including 7 (6.6%), 30 (9.1%), 14 (7.3%),
and 11 (9.5%) isolates from California, Illinois, Ontario, and Wisconsin, respectively,
were not identified. None of these isolates produced reproductive structures. Morphology
of the mycelia and colony characteristics did not provide sufficient information for
identification of these fungi. Similarly, information generated through molecular testing
was not adequate for determining their genera or species. No pathogenicity test was
conducted for any of these isolates. Considering their low occurrence in the roots, it is
believed that these fungi were not pathogenic on horseradish, but were rather secondary
colonizers.
41
FIGURES AND TABLES
Table 2.1 Detection of fungi in horseradish roots in 2008 and 2009
No. of
roots
tested
No. of
roots with
fungi (%)
Roots with fungal isolates [no. (%)]b
All fungi
Verticillium
spp.
Fusarium
spp.
Other
identified
fungic
Unknown
fungi
Locationa
Cultivar
California
Czechoslovakia
45
37 (82)
63
21 (33)
19 (30)
19 (30)
4 (7)
(Tule Lake)
Tule Lake
30
28 (93)
43
15 (35)
9 (21)
16 (37)
3 (7)
Illinois
Big Top Western
72
67 (93)
132
65 (49)
34 (26)
26 (20)
7 (5)
(Collinsville)
1573
234
183 (78)
196
81 (42)
66 (33)
26 (13)
23 (12)
Ontario, Canada
1590 Md
75
71 (95)
105
52 (50)
17 (16)
30 (29)
6 (5)
(Ancaster/Brantford)
1590 We
40
35 (88)
87
51 (59)
10 (12)
18 (20)
8 (9)
Wisconsin
Big Top Western
40
36 (90)
43
5 (12)
30 (70)
6 (14)
2 (4)
(Eau Clair)
15K
80
71 (89)
73
7 (10)
45 (62)
12 (16)
9 (12)
aRoots
from California, Illinois, and Ontario were collected from commercial fields. Roots from Wisconsin were from
research plots.
b
Two or more different fungal isolates grew out of some roots.
cOther identified fungal genera included Alternaria, Arthopyreniaceae, Chaetomium, Cylindrocarpon, Colletotrichum,
Cordyceps, Neonectria, Penicillium, Phoma, Phomopsis, Pyrenochaeta, Pythium, Rhizoctonia, Rhizopus, Rhizopycnis,
Trichurus, and Ulocladium.
d
Roots collected from Ancaster, ON.
e
Roots collected from Brantford, ON.
42
Table 2.2 Nucleotide sequences of primers used for amplifying target genes of isolated fungi
Locus
ITSa
Primer
ITS 4
ITS 5
Nucleotide sequence (5’-3’)
TCCTCCGCTTATTGATATGC
GGAAGTAAAAGTCGTAACAAGG
Target fungi
All fungi
cox3b*
COX 3A
COX 3B
TGATTTAGAGATSTAATATCAGAAG
CCGTGGAAACCTGTSCCAAAATA
Verticillium
nad1c*
NAD 1A ATGGCSAGTATGCAAAGAAGA
Verticillium
NAD 1B GCATGTTCTGTCATAAASCCACTAAC
mtSSUrDNAd
NMS1
NMS2
CAGCAGTGAGGAATATTGGTCAATG
GCGGATCATCGAATTAAATAACAT
Fusarium
EF1- alphae†
EF-1
EF-2
ATGGGTAAGGARGACAAGAC
GGARGTACCAGTRATCATGTT
Fusarium
a
ITS1-5.8S rDNA-ITS2 region (Internal Transcribed Spacer).
Mitochondrial cytochrome oxidase subunit III gene.
c
Mitochondrial NADH dehydrogenase subunit I gene.
d
Mitochondrial Small Subunit rDNA gene.
e
Elongation Factor 1-α.
*
S=C or G
†
R=A or G
b
43
Table 2.3 Fungal genera isolated from internally discolored horseradish roots in 2008 and 2009
No. of isolates
Phylum
Genus
Ascomycota
Alternaria
Arthopyreniaceae
Chaetomium
Cylindrocarpon
Colletotrichum
Cordyceps
Fusarium
Neonectria
Penicillium
Phoma
Phomopsis
Pyrenochaeta
Rhizopycnis
Trichurus
Ulocladium
Verticillium
California
Illinois Ontario
Wisconsin
Total
3
3
6
1
28
1
1
3
7
4
3
36
9
21
100
6
11
4
146
11
4
15
26
8
24
5
83
7
75
3
12
30
3
4
6
21
16
229
9
33
1
3
23
4
4
3
277
Basidiomycota Rhizoctonia
1
-
-
-
1
Zygomycota
-
-
-
7
7
Rhizopus
44
Table 2.4 Verticillium species isolated from horseradish roots in 2008 and 2009a
Location
No. of
Verticillium
isolates
California
Illinois
Ontario
Wisconsin
36
146
83
12
VD group
b
Verticillium species [no. (%)]
VL groupb
V. dahliae type Ac
V. dahliae type Bc
0 (0)
121 (83)
77 (93)
12 (100)
V. longisporum c
2 (6)
21 (14)
6 (7)
0 (0)
34 (94)
4 (3)
0 (0)
0 (0)
a
The roots were cultured on potato dextrose agar (PDA)
Morphological characterization; VD=V. dahliae group, VL=V. longisporum group
c
Molecular identification
b
Table 2.5 Fusarium species isolated from horseradish roots in 2008 and 2009*
Location
California
Illinois
Ontario
Wisconsin
Fusarium species [no. (%)]
No.of
F.
Fusarium F.
F.
F.
F.
equiseti
isolates
acuminatum commune
oxysporum proliferatum
-like species
28
10 (35)
0 (0)
0 (0)
3 (10)
0 (0)
100
29 (29)
3 (3)
12 (12)
13 (13)
0 (0)
26
13 (50)
0 (0)
0 (0)
6 (23)
0 (0)
75
43 (57)
4 (5)
0 (0)
10 (13)
3 (4)
* The roots were cultured on potato dextrose agar (PDA)
45
F.
solani
15 (55)
43 (43)
7 (27)
16 (21)
Table 2.6 Occurrence of the internal root discoloration of horseradish plants (n=6) following inoculation with Verticillium
species
Disease development
a
Fungus
Disease
occurrence
1b
Verticillium dahliaed
(Type A)
Incidencee
Severityg
0.0 af
0.0 a
Verticillium dahliaeh
(Type A)
Incidence
Severity
Verticillium dahliaei
(Type B)
Incidence
Severity
BTWa
3b
4b
50.0 b
10.2 a
83.3 b
18.3 a
83.3 b
37.9 b
47.9
18.6
0.0 a
0.0 a
50.0 ab
4.6 a
66.7 b
25.3 b
83.3 b
37.9 b
51.2
19.1
50.4
9.7
16.7 a
0.8 a
50.0 ab
2.5 a
66.7 ab
25.3 b
100.0 b
28.0 b
51.6
13.7
100 b
31.3 b
39.6
12.1
0.0 a
0.0 a
33.3 a
3.8 ab
83.3 b
18.3 b
100.0 b
37.6 c
39.6
15.2
16.7
0.8
NSk
NS
0
0
0
0
0
0
16.7
0.8
NS
NS
1573
3b
4b
50.0 b
1.8 ab
83.3 bc
21.7 bc
100.0 c
37.6 c
41.4
16.3
0.0 a
0.0 a
16.7 a
0.8 a
66.7 ab
5.5 ab
83.3 b
19.5 bc
83.3 b
25.0 c
52.7
14.1
33.3 a
1.7 a
66.7 ab
5.5 a
83.3 ab
18.3 b
100.0 b
34.7 c
Verticillium longisporumj Incidence
Severity
0.0 a
0.0 a
33.3 a
6.0 a
83.3 b
21.7 b
Control
0
0
0
0
16.7
0.8
Incidence
Severity
2b
a
LSDc
1b
2b
LSD
Horseradish cultivars (BTW=Big Top Western).
Month after inoculation of roots with the pathogen.
c
Least significant difference (P=0.05).
d
V. dahliae type A isolate from Ontario.
e
Percentage of roots affected.
f
Values within each row and for each cultivar with a letter in common are not significantly different from each other according
of Fisher‟s protected LSD (P=0.05).
g
Percentage of root area discolored at the cross sections (5=1-10%; 18=11-25%; 38=26-50%; 75.5=51-100%).
h
V. dahliae type A isolate from Illinois.
i
V. dahliae type B isolate from Illinois.
j
V. longisporum isolate from California.
k
NS = not significant.
b
46
Table 2.7 Occurrence of the internal root discoloration and root rot of horseradish (n=12) following inoculation with Fusarium
species
Disease development
Fungus
Fusarium
acuminatum
Fusarium
commune
Fusarium
equiseti
-like species
Fusarium
oxysporum
Fusarium
proliferatum
Fusarium
solani
Control
Disease
IRDd
IRD
RRi
IRD
IRD
IRD
IRD
IRD
RR
Disease
occurrence
Incidencee
Severityg
Incidence
Severity
Incidencej
Incidence
1b
16.7 af
0.8 a
33.3 ab
1.7 ab
0.0 a
0
Severity
Incidence
Severity
Incidence
Severity
Incidence
Severity
Incidence
Severity
Incidence
0
16.7 a
0.8 a
0
0
33.3 a
1.7 a
0
0
0
2b
66.7 b
5.5 ab
58.3 b
8.9 bc
16.7 ab
8.3
1573a
3b
66.7 b
10.9 b
58.3 b
11.7 c
41.7 b
16.7
4b
91.7 b
18.8 c
0.0 a
0.0 a
100.0 c
16.7
LSDc
34.9
7.8
36.2
7.7
26.6
NS
1b
25.0 a
1.7 a
33.3
2.1
0.0 a
0
6.4
58.3 b
7.3 ab
16.7
0.8
66.7 ab
7.7 a
0
0
0
1.9
83.3 c
15.1 b
16.7
0.8
66.7 ab
18.7 a
8.3
0.4
0
1.9
91.7
25.4 c
25
1.3
83.3 b
21.7 b
16.7
0.8
0
NS
53.9
8.7
NS
NS
38.6
9.4
NS
NS
NS
0
16.7 a
0.8 a
0
0
16.7 a
0.8 a
0
0
0
a
2b
33.3 a
2.3 a
25
4
33.3 ab
0
BTWa
3b
50.0 a
9.6 ab
33.3
14.5
50.0 ab
16.7
4b
91.7 b
16.7 b
16.7
3
83.3 c
8.3
LSD
34.6
8.7
NSh
NS
33.6
NS
0
41.7 ab
6.4 a
0
0
66.7 b
5.5 a
0
0
0
0.8
66.7 bc
12.3 a
16.7
0.8
66.7 b
15.3 b
0
0
0
0.4
83.3 c
33.7 b
16.7
0.8
83.3 c
31.7 c
8.3
1.5
0
NS
37
14.6
NS
NS
36.5
8.7
NS
NS
NS
Horseradish cultivars (BTW=Big Top Western).
b
Months after inoculation of roots with the pathogen.
c
Least significant difference (P=0.05).
d
IRD= internal root discoloration.
e
Percentage of roots affected.
f
Values within each row and for each cultivar with a letter in common are not significantly different from each other according
of Fisher‟s protected LSD (P=0.05).
g
Percentage of root area discolored at the cross section (5=1-10%; 18=11-25%; 38=26-50%; 75.5=51-100%).
h
NS = not significant.
i
RR = root rot.
j
Percentage of plants with rotted root.
47
A
B
C
D
Figure 2.1 Cross and longitudinal sections of horseradish roots. A, cross section of an
asymptomatic horseradish root; B, cross section of a horseradish root showing internal
discoloration symptoms; C, longitudinal sections of an asymptomatic horseradish root;
D, longitudinal sections of an internally discolored horseradish root.
48
Figure 2.2 Frequency of fungal genera isolated from horseradish roots collected from California, Illinois, Ontario (Canada),
and Wisconsin in 2008 and 2009
49
Figure 2.3 Percentage of the fungal isolates identified as Verticillium and Fusarium
species in horseradish growing areas in 2008 and 2009.
50
50 µm
Figure 2.4 Microsclerotia of Verticillium dahliae
produced on potato dextrose agar. Note the spherical
shape of the microsclerotia.
50 µm
Figure 2.5 Microsclerotia of Verticillium longisporum
produced on potato dextrose agar. Note the irregular
shape of the microsclerotia.
51
Figure 2.6 Phialides (4- to 5-per whorl) of Verticillium
dahliae produced on potato dextrose agar.
Figure 2.7 Phialides (2-to 3-per whorl) of Verticillium
longisporum produced on potato dextrose agar.
52
20 µm
Figure 2.8 Conidia of Verticillium dahliae (mean length
4.4±0.23 μm) produced on potato dextrose agar.
20 µm
Figure 2.9 Conidia of Verticillium longisporum (mean
length 7.9±0.08 μm) produced on potato dextrose agar.
53
SDV1025
ITS
637 characters
90 parsimony-informative
characters
1000 trees
313 steps
CI = 0.8179
RI = 0.7957
65
SDV1002
SDV1006
V. dahliae M5 (AY555948)
V. dahliae UZ132 (AY555949)
CBV1046
CBV1014
CBV1006
CAV1004
96 CAV1003
SCV1010
SCV1005
V. dahliae T9 (AY555947)
V. longisporum G19 (AY555950)
TBV3014
TAV3001
VBV3004
VAV3021
VAV3003
V. longisporum K2(AY555951)
V. longisporum K12 (AY555952)
67
V. dahliae Vd292 (AF363994)
CBV2034
52
SCV2008
SDV2066
SDV2064
SDV2041
100
V. albo-atrum V90 (AY555954)
65
V. albo-atrum 1776 (AY555953)
V. nubilum 278734
67
94
56
V. tricorpus 22
V. tricorpus188
V. nigrescens 193
100
V. theobromae 225818
Acrostalagmus luteo-albus 868
59
V. catenulaulatum 113078
V. fungicola 88936
Neurospora crassa
5 changes
Figure 2.10 Maximum parsimony tree generated from reference sequences of internal
transcribed spacer (ITS) ITS1-5.8S rDNA-ITS2 with sequences of Verticillium isolates
from horseradish roots from North America. Numbers next to branches represent
bootstrap values of 1000 replicates calculated with maximum parsimony. Only bootstrap
values higher than 50% are shown.
54
V. dahliae M5 (AY555921)
cox3
376 characters
86 parsimony-informative
characters
15 trees
249 steps
CI = 0.6466
RI = 0.6318
V. dahliae UZ132 (AY555922)
SDV1025
SDV1002
SDV1006
CBV1046
CBV1006
60 CAV1004
CAV1003
V. dahliae
CBV1014
SCV1010
SCV1005
V. dahliae T9 (AY555920)
CBV2034
SCV2008
SDV2066
SDV2064
93
SDV2041
V. longisporum G19 (AY555923)
V. longisporum K12 (AY555925)
TBV3014
TAV3001
90
VBV3004
VAV3021
64
58
V. longisporum
V. longisporum K2 (AY555924)
VAV3003
V. albo-atrum V90 (AY555927)
V. albo-atrum 1776 (AY555926)
V. nubilum 278734
100
V. tricorpus 22
V. tricorpus 188
V. theobromae 225818
61
Acrostalagmus luteo-albus 868
V. nigrescens 193
V. catenulaulatum 113078
78
V. fungicola 188936
Neurospora crassa
5 changes
Figure 2.11 Maximum parsimony tree generated from reference sequences of
mitochondrial gene, cytochrome oxidase subunit III (cox3) with sequences of Verticillium
isolates from horseradish roots from North America. Numbers next to branches represent
bootstrap values of 1000 replicates calculated with maximum parsimony. Only bootstrap
values higher than 50% are shown.
55
V. dahliae M5 (AY555977)
nad1
523 characters
120 parsimony-informative
characters
4 trees
385 steps
CI = 0.7143
RI = 0.6927
V. dahliae UZ132 (AY555978)
SDV1025
V. dahliae
Type A
SDV1002
SDV1006
64 CBV1046
CBV1014
CBV1006
CAV1004
CAV1003
SCV1010
SCV1005
CBV2034
SCV2008
97
SDV2066
V. dahliae
Type B
V. dahliae T9 (AY555976)
SDV2064
SDV2041
V. longisporum K2 (AY555980)
V. longisporum K12 (AY555981)
60 TBV3014
99
TAV3001
VBV3004
VAV3021
68
VAV3003
69
93
V. albo-atrum V90 (AY555983)
V. albo-atrum 1776 (AY555982)
74
56
V. tricorpus 22
80
V. tricorpus188
V. nubilum 278734
72
V. theobromae 225818
V. nigrescens 193
Acrostalagmus luteo-albus 868
100
V. catenulaulatum 113078
V. fungicola 188936
Neurospora crassa
5 changes
Figure 2.12 Maximum parsimony tree generated from reference sequences of
mitochondrial gene, NADH dehydrogenase subunit I (nad1) with sequences of
Verticillium isolates from horseradish roots from North America. Numbers next to
branches represent bootstrap values of 1000 replicates calculated with maximum
parsimony. Only bootstrap values higher than 50% are shown.
56
V. longisporum
V. longisporum G19 (AY555979)
V. dahliae M5
cox3 + nad1
899 characters
206 parsimony-informative
characters
3 trees
637 steps
CI = 0.6845
RI = 0.6683
V. dahliae UZ132
SDV1025
SDV1002
SDV1006
63 CBV1046
CBV1006
CAV1004
CAV1003
68
V. dahliae
Type A
CBV1014
SCV1010
SCV1005
V. dahliae T9
Type B
CBV2034
SCV2008
SDV2066
SDV2064
100
SDV2041
V.longisporum G19
V. longisporum
V. longisporum K2
V. longisporum K12
83 TBV3003
TAV3001
100
70
VBV3004
VAV3021
VAV3003
98
73
V. albo-atrum V90
V. albo-atrum 1776
100
V. tricorpus 22
74
77
V. tricorpus188
. nubilum 278734
V. theobromae 225818
Acrostalagmus luteo-albus 868
V. nigrescens 193
V. catenulaulatum 113078
100
V. fungicola 188936
Neurospora crassa
10 changes
Figure 2.13 Maximum parsimony tree generated from combined sequences of
mitochondrial genes, nad1 and cox3 genes with sequences of Verticillium isolates from
horseradish roots from North America. Numbers next to branches represent bootstrap
values of 1000 replicates calculated with maximum parsimony. Only bootstrap values
higher than 50% are shown.
57
Figure 2.14.1 Colony pattern on PDA (left) and macrocondia on SNA (right) of
Fusarium acuminatum.
Figure 2.14.2 Colony pattern on PDA (left) and mycelium and macrocondia on SNA
(right) of Fusarium commune.
58
Figure 2.14.3 Colony pattern on PDA (left) and macrocondia on SNA (right) of
Fusarium equiseti.
Figure 2.14.4 Colony pattern on PDA (left) and macrocondia and microconidia
on SNA (right) of Fusarium oxysporum.
59
Figure 2.14.5 Colony pattern on PDA (left) and macrocondia on SNA (right) of
Fusarium proliferatum.
Figure 2.14.6 Colony pattern on PDA (left) and macrocondia on SNA (right) of
Fusarium solani.
60
SD5115
ITS region
429 characters
80 parsimonyinformative characters
696 trees
163 steps
CI = 0.7914
RI = 0.9470
100 SD5030
67
WA5039
F. equiseti (AB425996)
97
83
F. cerealis NRRL25491 (AF006340)
F. venenatum (AY188922)
99
F. oxysporum f. sp. melonis 348 (DQ016233)
68`
SD3152
CA3022
99
CA3008
WA3037
F.solani S-0900 (EF152426)
SD1184
99
SC1018
WA1072
67
CA1018
100
61
99 SC1042
SC1013
Nectria haematococca (AY310442)
SD1212
CB1034
F. tricinctum (AF008921)
F. acuminatum IBE000017 (EF531233)
100
SD2018
WA2077
CA2060
F. commune 9245H (DQ016195)
F. redolens aurim1114 (DQ093672)
80
SD4139
WA4023
92
WB4031
68
F. proliferatum 9223F (DQ016238)
WA6005
Cylindrocarpon cylindroides CBS 503.67 (AY677261)
5 changes
Figure 2.15 Maximum parsimony tree generated from reference sequence of Internal
transcribed spacer (ITS) ITS1-5.8S-ITS2 with sequences of Fusarium isolates from
horseradish roots from North America. Numbers next to branches represent bootstrap
values of 1000 replicates calculated with maximum parsimony. Only bootstrap values
higher than 50% are shown.
61
NMS
562 characters
59 parsimonyinformative characters
4 trees
168 steps
CI = 0.9167
RI = 0.9381
F. commune 9245H (DQ016145)
F. redolens NRRL28058 (AF324293)
92
SD4139
WA4023
WB4031
100 F. proliferatum 31X4 (DQ831947)
WA6005
F. oxysporum f. sp. melonis TX388 (DQ831943)
85 SD3152
93
CA3022
CA3008
WA3037
SD5115
100
SD5030
WA5039
100
F. cerealis (U85551)
F. venenatum (U85560)
F. tricinctum NZ62 (EU919401)
F. acuminatatum NZ50 (EU919400)
70
SD2018
WA2077
CA2060
F. solani NRRL22382(AF125023)
SD1212
SD1184
SC1018
98
SC1042
SC1013
96
WA1072
CA1018
CB1005
N. haematococca AFTOL-ID 161 (FJ713623)
C. cylindroides (AF315201)
5 changes
Figure 2.16 Maximum parsimony tree generated from reference sequence of
mitochondrial small subunit rDNA genes (mtssu rDNA) with sequences of Fusarium
isolates from horseradish roots from North America. Numbers next to branches represent
bootstrap values of 1000 replicates calculated with maximum parsimony. Only bootstrap
values higher than 50% are shown.
62
EF1-alpha
380 characters
116 parsimonyinformative characters
59 trees
344 steps
CI = 0.8052
RI = 0.9193
F. oxysporum f. sp. melonis TX388(DQ837696)
60
95
SD3152
CA3008
100
CA3022
WA3037
71
F. commune NRRL28387 (AF246832)
64
92
WB4031
98
SD4139
WA4023
53
F. proliferatum NRRL43666(EF452997)
WA6005
F. redolens NRRL28412 (AF324300)
100
WA1072
CA1018
58
SC1018
SD1184
91
F. solani f. sp. batatas NRRL22402 (AF178344)
SD1212
CB1005
100
68
54
59
N. haematococca MAFF 840047 (DQ452423)
SC1013
F. solani NRRL20547 (DQ247544)
SC1042
62 SD5115
97
94
SD5030
WA5039
71
F. equiseti DAOM236361 (DQ842099)
88
F. cerealis NRRL13393 (AF212467)
F. venenatum VI01179 (AJ543635)
F. acuminatum R-6678 (FJ154737)
96
99
SD2018
WA2077
CA2060
F. tricinctum PDD17HB (EU327337)
C. cylindroides CBS 324.61
(DQ789732)
5 changes
Figure 2.17 Maximum parsimony tree generated from reference sequence of translation
protein Elongation Factor 1-alpha region (EF1-alpha) with sequences of Fusarium
isolates from horseradish roots from North America. Numbers next to branches represent
bootstrap values of 1000 replicates calculated with maximum parsimony. Only bootstrap
values higher than 50% are shown.
63
64 F. commune
ITS / NMS / EF 1-alpha
1422 characters
249 parsimonyinformative characters
18 trees
715 steps
CI = 0.8028
RI = 0.9126
WB4031
95
F. commune
SD4139
WA4023
54
F. redolens
F. proliferatum
100
F. proliferatum
WA6005
62
SD3152
64
CA3008
100
F. oxysporum
100
CA3022
F. oxysporum
complex
WA3037
WA1072
60
CA1018
94
SC1018
73
SD1184
F. solani
complex
SD1212
100
CB1005
80
85 SC1042
SC1013
50
58
F. solani
N. haematococca
F. acuminatum
94 SD2018
F. acuminatum
WA2077
100
CA2060
F. tricinctum
61 SD5115
100
F. sp.
SD5030
WA5039
100
F. cerealis
99
F. venenatum
C.cylindro
10 changes
Figure 2.18 Maximum parsimony tree generated from combined sequences of ITS, NMS,
and EF-1α genes with sequences of Fusarium isolates from horseradish roots from North
America. Numbers next to branches represent bootstrap values of 1000 replicates
calculated with maximum parsimony. Only bootstrap values higher than 50% are shown.
64
LITERATURE CITED
Atibalentja, N., and Eastburn, D.M. 1998. Verticillium dahliae resistance in horseradish
germ plasm from the University of Illinois collection. Plant Dis. 82:176-180.
Babadoost, M., Chen, W., Bratsch, A.D., and Eastman, C.E. 2004. Verticillium
longisporum and Fusarium solani: two new species in the complex of internal
discoloration of horseradish roots. Plant Pathol. 53:669-676.
Babadoost, M., Eranthodi, A., Jurgens, A., Hippard, K., and Wahle, E. 2007. ThermoTherapy and use of biofungicides and fungicides for management of internal
discoloration of horseradish roots,2006. pp 25-31. In 2007 Horseradish Res. Rev. & Proc.
Horseradish Growers School.
Berbee, M. L., and Taylor, J.W. 2001. Fungal molecular evolution: gene trees and
geologic time. In The Mycota. Vol. VII. Part B. pp 229-245. In Systemics and Evolution
(D.J. McLaughlin, E.G. McLaughlin, and P.A. Lemke (eds). Springer-Verlag, Hidelberg,
Germany.
Bruns, T.D., White, T.J., and Taylor, J.W. 1991. Fungal molecular systematics. Annu.
Rev. Ecol. and Syst. 22:525-564.
Bullerwell, C.F., Forget, L. and Lang, B.F. 2003. Evolution of monoblepharidalean fungi
based on complete mitochondrial genome sequences. Nucleic Acid Res. 31:1614-1623.
Butler, D.K., and Metzenberg, R.L. 1989. Premeiotic change of nucleolus organizer size
in Neurospora. Genetics 122:783-791.
Carder, J.H., and Barbara, D.J. 1989. Taxonomic status of putative Verticillium alboatrum isolates. FEMS Microbiol. Letter 170:211-219.
Chun J. 1995. Computer assisted classification and identification of actinomycetes. Ph.D.
Thesis. University of Newcastle. Newcastle upon Tyne, UK.
Eastburn, D.M., and Chang, R.J. 1994. Verticillium dahliae: a causal agent of root
discoloration of horseradish in Illinois. Plant Dis. 78:496-498.
Geiser, D.M., del Mar Jimenez-Gasco, M., Kang, S., Makalowska, I., Veeraraghavan, N.,
Ward, T.J., Zhang, N., Kuldau, G.A., and O'donnell, K. 2004. FUSARIUM-ID v. 1.0: A
DNA sequence database for identifying Fusarium. Eur. J. Plant Pathol. 110:473-479.
Gerber, J.M., Doll, C.C., Simons, R.K., and Fillingim, K.E. 1983. Internal discoloration
of horseradish-development of symptoms. Univ. Ill. Veg. Res. Rep. Hortic. Ser. 47:34-37.
65
Hall, T.A. 1999. BioEdit: a user-friendly biological sequence alignment editor and
analysis program for Windows 95/98/NT. Nucleic Acid. Symp. Ser. 41:95-98.
Hawksworth, D.L., and Talboys, P.W. 1970. Verticillium albo-atrum. CMI, descriptions
of pathogenic fungi and bacteria No. 255. CABI Publishing, Wallingford, UK.
Hawksworth, D.L., and Talboys, P.W. 1970. Verticillium dahliae . CMI, descriptions of
pathogenic fungi and bacteria No. 256. CABI Publishing, Wallingford, UK.
Karapapa, V.K., Bainbridge, B.W., and Heale, J.B. 1997. Morphological and molecular
characterization of Verticillium longisporum comb. nov., pathogenic to oilseed rape.
Mycol. Res. 101:1281-1294.
Kouvelis, V.N., Ghikas, D.V., and Typas, M.A. 2004. The analysis of the complete
mitochondrial genome of Lecanicillium muscarium (synonym Verticillium lecanii)
suggests a minimum common gene organization in mtDNAs of Sordariomycetes:
phylogenetic implications. Fungal Genetics and Biol. 41:930-940.
Laemmlen, F. 2001. Alternaria Diseases. University of California, Division of
Agriculture and Natural Resources. Publication No. 8040:1-5.
Leslie, J.F., Summerell, B.A., and Bullock, S. 2006. The Fusarium Laboratory Manual.
Blackwell Publishing. Ames, IA. 388 pp.
Li, K.N., Rouse, D.I. and German, T.L. 1994. Differentiation of ascomycetes with PCR
primers. App. Env. Microbiol. 60:4321-4331.
Nelson, P.E., and Toussoun, T.A. 1983. Fusarium Species: An Illustrated Manual for
Identification. The Pennsylvania State University Press, University Park, PA. 206 pp.
O‟Donnell, K., Kistler, H.C., Cigelnik, E. and Ploetz, R.C. 1998. Multiple evaluationary
origin of the fungus causing Panama disease of banana: concordant evidence from
nuclear and mitochondrial gene genealogies. Proc. Natl. Acad. Sci. 95:2044-2049
Pantou, M.P., Mavridou, A. and Typas, M.A. 2003. IGS sequence variation, group-I
introns and the complete nuclear ribosomal DNA of the entomopathogenic fungus
Metarhiziu: excellent tools for isolate detection and phylogenetic analysis. Fungal
Genetics and Biol. 38:159-174.
Pantou, M.P., Strunnikova, O.K., Shakhnazarova, V.Y., Vishnevskaya, N.A., Papalouka,
V.G., and Typas, M.A. 2005. Molecular and immunochemical phylogeny of Verticillium
species. Mycol. Res. 109:889-902.
Percich, J.A., and Johnson, D.R. 1990. A root rot complex of horseradish. Plant Dis.
74:391-393.
66
Promputtha, I., Jeewon, R., Lumyong, S., McKenzie, E.H.C., and Hyde, K.D. 2005.
Ribosomal DNA fingerprinting in the identification of non sporulating endophytes from
Magnolia liliifera (Magnoliaceae). Fungal Diversity 20:167-186.
Skovgaard, K., Rosendahl, S., O'Donnell, K., and Nirenberg, H.I. 2003. Fusarium
commune is a new species identified by morphological and molecular phylogenetic data.
Mycologia 95:630-636.
Thomas, P., Swarna, G.K., Roy, P.K., and Patil, P. 2008. Identification of culturable and
originally non-culturable endophytic bacteria isolated from shoot tip cultures of banana
cv. Grand Naine. Plant Cell, Tissue and Organ Cult. 93:55-63.
Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., and Higgins, D.G. 1997.
The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment
aided by quality analysis tools. Nucleic Acids Res. 25:4876-4882.
White, T.J., Bruns, T., Lee, S., and Taylor, J. 1990. Amplification and direct sequencing
of fungal ribosomal RNA genes for phylogenetics. pp 315-322. In PCR Protocols: A
Guide to Methods and Applications. M.A. Innis, D.H. Gelfand, J.J. Sninsky, and T.J.
White (eds). Academic Press. San Diego, CA. 482 pp.
Yli-Mattila, T., Paavanen-Huhtala, S., Bulat, S.A., Alekhina, I.A., and Nirenberg, H.I.
2002. Molecular, morphological and phylogenetic analysis of the Fusarium avenaceum/F.
arthrosporioides/F. tricinctum species complex–a polyphasic approach. Mycol. Res.
106:655-669.
67
CHAPTER 3
BACTERIAL SPECIES ASSOCIATED WITH INTERNALLY DISCOLORED
HORSERADISH ROOTS
MATERIALS AND METHODS
Sample collection
A total of 621 discolored horseradish roots, including 80, 306, 115, and 120 from
California, Illinois, Ontario (Canada), and Wisconsin, respectively, were collected and
tested for the presence of bacteria during 2008 and 2009 (Table 3-1). The roots from
California, Illinois, and Ontario were collected from commercial fields, while roots from
Wisconsin were from research plots. Roots were either tested immediately or were stored
at 4 C until they were tested.
Bacterial isolation
The roots were washed with tap water and a 5-cm segment from the middle of
each root was collected. The segments were peeled to remove outer layers of the cortex to
minimize contamination. Peeled-segments were immersed into 6% NaClO solution for 1
min followed by immersing into 95% ethanol for 3 min and washing with sterile-distilled
water (SDW) three times (each time for 1 min). Five small segments (5 mm thick) of
each surface-sterilized root were cut in a sterile hood, and the segments were placed onto
nutrient agar (NA) in Petri plates for evaluation of the presence of bacteria in the roots.
NA was prepared by adding 23 g of nutrient agar (Difco™, Sparks, MD) to 1 L distilled
68
water and autoclaving at 121 C for 25 min. The plates were incubated at 22 C under 12
h light and 20 C with 12 h darkness. The plates were observed for development of
bacterial colonies 5, 10, and 15 days from the time of incubation. A loop-full of bacterial
cells from the edge of each colony was streaked onto another NA plate for single-cell
purification and identification of the bacteria.
DNA extraction, PCR amplification and sequencing
Isolated bacteria were grown in nutrient broth (NB) overnight. NB was prepared
by dissolving 8 g of nutrient broth (Difco™, Sparks, MD) in 1 L distilled water, and then
autoclaving it at 121 °C for 25 min. Bacterial cells were recovered from cell suspension
by centrifugation for 2 min at 11,400 x g. Total DNA was extracted from bacterial cells
using a Fast DNA Spin kit (MP. Solon, OH). Polymerase chain reaction (PCR) was
performed to amplify the 16S rRNA genes from the extracted DNA using the primer set
27f (5‟-AGA GTT TGA TCM GGC TCA G-3‟) and 1492r (5‟GGT TAC CTT GTT ACG
ACT T-3‟) (Lane, 1991).
The 50 µl PCR reaction mixture contained a final
concentration of 10X PCR buffer, 2.0 mM MgCl2, 2.5 mM dNTPs, 0.5 pM of each
primers, 2 units of Taq polymerase (Promega Corp., Madison, WI), and 100 ng of DNA
extract. Cycling condition for PCR (Model PCT-200; MJ Research Inc., Waltham, MA)
included initial denaturing for 2 min at 95 °C; 35 cycles of denaturation at 94 °C for 45 s,
annealing at 55 °C for 30 s, and extension at 72 °C for 1.5 min; followed by a final
extension at 72 °C for 5 min. Amplification was verified by running 10 μl of the PCR
product in a 1% agarose gel with 1X Tris-acetate EDTA (TAE), staining with ethidium
bromide, and visualizing under ultraviolet light. PCR products were purified with the
69
Wizard SV gel and PCR Clean-Up system (Promega Madison, WI) for sequencing.
Purified 16S rDNAs were sequenced at the University of Illinois Core DNA Sequencing
Facility.
RFLP analysis
The amplified 16S rDNAs were subjected to restriction fragment length
polymorphism (RFLP) analysis for screening their ligased fragment patterns. Ten
microliters of each PCR product was digested directly by 3 units of the restriction
enzyme Hae III by overnight incubation at 37 ºC. Digested products were separated in a
4% agarose gel at 115 V for 2 hours with 1 kb DNA ladder used as a marker. Restriction
patterns were visualized with ethidium bromide staining using a gel documentation
system under UV light. The RFLP analysis was performed at least twice for all isolates.
The isolates were grouped based on their RFLP patterns, and the DNAs of the
representative isolates were sequenced.
Phylogenetic analysis
Single-end sequence was performed with the same forward primer for PCR
amplification. The sequences were proofread and edited using Clustal X (Thompson et al.,
1997) and BioEdit version 7.0.9 (Hall, 1999). The edited sequences were compared with
the 16S rDNA sequences from the NCBI GenBank database and identified to genera. The
sequences
from
the
isolates
and
the
reference
sequences
from
GenBank
(http://www.ncbi.nlm.nih.gov/) were aligned using CLUSTAL X (Thompson et al.,
1997), and then the alignment was refined manually using PHYDIT version 3.2 program
70
(Chun, 1995; http://plaza.snu.ac.kr/~jchun/ phydit/). Since BLAST search results showed
the most similar reference sequence with query sequences, detailed species level
identifications were conducted by phylogenetic analysis. The neighbor-joining
phylogenetic tree was constructed using PHYLIP 3.57 c software package (Felsenstein,
1980) and the Kimura‟s 2-parameter distance model (Kimura, 1980) for distance matrix
calculations. A bootstrap analysis using 1,000 replications was performed to assess the
relative stability of branches.
In vitro antifungal activity tests
Isolates of the identified 18 genera and 52 species of bacteria were tested in a dual
culture study with Verticillium dahliae, V. longisporum, Fusarium oxysporum, and F.
solani to determine the effect of the bacteria on development of fungal colonies (Berg et
al., 2001). These fungi were isolated from internally discolored horseradish roots during
2008 and 2009.
The fungal species were grown on PDA in Petri plates at 22-24 C with 12 h
light/12 h darkness. When colonies were 7-10 days old, a 3-mm-diameter plug was taken
from the edge of the colony and transferred onto the center of Waksman agar (WA) in
Petri plates. WA was prepared by adding 5 g proteose-peptone, 10 g glucose, 3 g beef
extract, 5 g NaCl, and 20 g agar to 1 L distilled water, and adjusting its pH to 6.8 (Berg et
al., 2001). Three bacterial isolates were lined at 30 mm distance from the plug of the test
fungus. Since Verticillium species grow slowly, plates with plugs of Verticillium species
were pre-inoculated for 2 days at 22-24 C with 12 h light/12 h darkness prior to
71
inoculation with bacteria (Alstrom, 2000). Zones of inhibition were measured 5 days after
inoculation with bacteria and incubation at 22-24 C with 12 h light/12 h darkness.
Antifungal activity of the bacterium was evident as mycelial growth of the test fungus
was prohibited in the direction toward the bacterial line (Paul et al., 2007). The level of
inhibition was determined by the distance from the edge of fungal colony to the edge of
the bacterial colony (inhibition zone). The width of inhibition zones between fungal
colony and bacterial colony line was used to determine the inhibition efficacy of the
bacterium on the test fungus, which include: > 8 mm (+++, strong inhibition), 2-8 mm
(++, moderate inhibition) and < 2 mm (+, weak inhibition) as suggested by Paul et al.
(2007).
RESULTS
Identification of bacterial isolates
Bacterial colonies developed on the cultured segments of 509 of 621 roots. The
ratios of roots with bacteria to the number of root cultures were 57/80, 264/306, 81/115,
and 107/120 from California, Illinois, Ontario, and Wisconsin, respectively (Table 3-1).
A total of 824 bacterial isolates were collected, which included 93, 416, 138, and 177
from California, Illinois, Ontario, and Wisconsin, respectively (Table 3-1). Two or more
different bacterial colonies developed on the cultured segments of some roots. The
isolates were grouped according to the RFLP patterns of their 16S rDNA. The isolated
bacterial groups included 12, 16, 12, 18, 18, 12, and 10 groups from „Czechoslovakia‟
(Califronia), „Big Top Western‟ (Illinois), „1573‟ (Illinois), „1590‟ (Brandford, Ontario),
„1590‟ (Ancaster, Ontario), „Big Top Western‟ (Wisconsin), and „15K‟ (Wisconsin),
72
respectively (Figure 3-1). Isolates with the same RFLP patterns were considered the same
species.
Several isolates from the same group with the same RFLP pattern were sequenced
for confirmation of identification. Identification of the bacterial genera and species was
based on the partial sequences of their 16S rDNA (Tables 3-2 and 3-3). A total of 216
isolates were sequenced using their partial forward 16S rDNA sequences, and were
compared with reference sequences from the NCBI GenBank database. As a result 52
different groups, representing 52 species in 20 genera, were identified (Tables 3-2 and 33). The bacterial isolates were assigned to five classes including Actinobacteria, Bacilli,
Alphaproteobacteria, Betaproteobacteria, and Gammaproteobacteria (Tables 3-2 and 3-3).
The majority of isolates were grouped in bacterial classes Gammaproteobacteria (49.9%)
and Bacilli (39.5%).
Occurrence of bacteria in different areas
California isolates
Bacterial isolates from California were placed in five classes: Actinobacteria,
Bacilli, Alpahproteobacteria, Betaproteobacteria, and Gammaproteobacteria (Figure 3-21). The highest number (37.8%) of isolates belonged to the class Bacilli (Table 3-2).
Nine genera of bacteria were identified which included: Achromobacter (3.5%) and
Microbacterium (3.5%) from the class Actinobacteria; Ochrobactrum (3.5%) from the
class Alphaproteobacteria; Bacillus (21.2%) and Paenibacillus (17.5%) from the class
Bacilli; Sphingobacterium (3.5%) from the Betaproteobacteria; and Kluyvera (3.5%),
73
Pseudomonas (10.5%), and Serratia (17.5%) from the class Gammaproteobacteria (Table
3-2). Altogether ten bacterial species were identified. The most prevalent species were
Bacillus thuringiensis (six roots), Paenibacillu tylopili (five roots), and Serratia
proteamaculans (five roots) (Table 3-3). Nine isolates (15.8%) were not identified.
Illinois isolates
The bacterial isolates from Illinois were assigned to four classes, including
Actinobacteria, Bacilli, Alphaproteobacteria, and Gammaproteobacteria (Figure 3-2-2).
Most of the isolates belonged to the classes Bacilli (43.3%) and Gammaproteobacteria
(44.7%). Genera Bacillus and Pseudomonas were the major taxa (32.2 and 39% of total
isolates, respectively). Identified bacterial genera included Achromobacter (4.3%) and
Sanguibacter (1.9%) in the class Actinobacteria; Ochrobactrum (1.9%) in the class
Alphaproteobacteria; Bacillus (32.2%) and Paenibacillus (11.1%) in the class Bacilli; and
Enterobacter (1.9%), Pseudomonas (39%), Rahnella (1.9%) and Stenotrophomanas
(1.9%), in the class Gammaproteobacteria (Table 3-2). A total of 21 species were
identified, and the most prevalent species were Bacillus cereus, B. thuringiensis, and
Pseudomonas extremorientalis with 34, 12, and 12 isolates (representing 12 roots),
respectively (Table 3-3). Thirteen isolates (4%) were not identified.
Ontario isolates
Isolates from Ontario (Canada) were placed in four classes including
Actinobacteria, Bacilli, Betaproteobacteria, and Gammaproteobacteria (Figure 3-2-3).
The dominant class was Gammaproteobacteria with 40.3% of the isolates (Table 3-2). A
74
total of 12 genera were identified which were: Achromobacter (1.6%), Arthrobacter
(7.8%), Curtobacterium (1.6%), Microbacterium (4.7%), Plantibacter (2.3%) in the class
Actinobacteria; Bacillus (34.1%) and Paenibacillus (2.3%) in the Bacilli; Variovorax
(1.6%) in the Betaproteobacteria; and Pantoea (6.2%), Pseudomona (20.9%), Rahnella
(11.6%), and Yersinia (1.6%) in the class Gammaproteobacteria (Table 3-2). The most
frequently isolated species were Arthrobacter nicotinovorans, Bacillus anthracis,
Pseudomonas monteilii, and Rahnella aquatic with 10, 17, 18, and 15 isolates
(representing 15 roots), respectively (Table 3-3). Five isolates (3.9%) were not identified.
Wisconsin isolates
Four bacterial classes were identified from the isolates from Wisconsin, which
included Actinobacteria, Bacilli, Alphaproteobacteria, and Gammaproteobacteria (Figure
3-2-4). The dominant class was Gammaproteobacteria with 68.2% of the isolates (Table
3-2). The class Bacilli with 23.4% of the isolates was the second most abundant group. A
total of eight genera were identified, which included: Achromobacter (0.9%) and
Microbacterium (1.9%) in the class Actinobacteria; Bacillus (23.4%) in the class Bacilli;
Agrobacterium (1.9%) in the class Alphaproteobacteria; and Enterobacter (3.7%),
Pantoea (12.1%), Pseudomonas (42.1%), and Rahnella (10.3%) in the class
Gammaproteobacteria (Table 3-2). Fourteen species of bacteria were identified from
Wisconsin (Table 3-3). Bacillus cereus (14 roots), Pantoea agglomerans (13 roots),
Pseudomonas grimontii (17 roots), and P. extremorientalis (17 roots) were the most
prevalent species (Table 3-3). Four isolates (3.7%) were not identified.
75
In vitro antifungal activity
The results of in vitro tests of screening of the isolated bacteria for their antifungal
activities against V. dahliae, V. longisproum, F. oxysporum, and F. solani are presented
in Table 3-4. Thirty four species out of 52 identified species exhibited antifungal activity
against at least one of the above-mentioned fungi. Three species of Bacillus, (B.
amyloliquefaciens, B. atrophaeus, and B. subtilis) exhibited the strongest antifungal
activities. Also Pseudomonas species, P. grimontii, P. mediterraneae, and P. montelli
exhibited strong antifungal activity. In addition, bacteria in the genera of Enterobacter,
Microbacterium, Paenibacillus, and Serratia moderately suppressed the growth of the
fungi.
DISCUSSION
Although the occurrence of bacteria in internally discolored roots of horseradish
has already been reported (Atibalentja and Eastburn, 1998; Babadoost et al., 2004;
Eastburn and Chang, 1994; Percich and Johnson, 1990; Potschke, 1923), species of the
bacteria in the roots and their roles have not been determined. This is the most
comprehensive study to accurately identify species of the bacteria associated with
internal discoloration of horseradish roots and to determine their interactions with the
pathogens causing the discoloration.
Occurrence of 52 species in 20 bacterial genera in five classes indicates that
horseradish roots harbor diverse bacterial communities in their roots. Although
pathogenecity tests of the isolated bacteria were not conducted, it is unlikely any of the
76
identified bacterial species causes discoloration in horseradish roots for following
reasons: (1) bacteria have been routinely isolated from asymptomatic roots (Babadoost et
al., 2004); (2) only Verticillium and Fusarium species have been reported to cause the
internal discoloration (Atibalentja and Eastburn, 1998; Babadoost et al., 2004; Percich
and Johnson, 1990); and (3) whenever only bacteria have been isolated from the
discolored root, they are usually Pseudomonas or Bacillus species, which are antagonistic
to fungi and likely the secondary invaders of roots. However, pathogenecity tests are
needed to clarify if any of the isolated bacteria is involved in causing internal
discoloration of horseradish roots.
Studying the relationships between the isolated fungal and bacterial species from
internally discolored horseradish roots showed that some of the bacterial isolates are
antagonistic against V. dahliae, V. longisporum, F. oxysporum, and F. solani (the causal
agents of the internal root discoloration), in vitro. Most of the bacterial isolates with
antifungal activity belong to the genera Bacillus and Pseudomonas. Several species of
Bacillus and Pseudomonas have been reported to be antagonist to fungal pathogens (Berg
et al., 2001; Ziedan, 2006). Some of the bacterial isolates in this study may be good
biocontrol agents for control of soil-borne pathogens of Verticillium and Fusarium
species.
Identification of bacterial isolates in this study was based on analyzing sequences
of the 16S rDNA. The small subunit of rDNA exists universally among bacteria and
includes regions with species-specific variability, which makes it possible to identify
77
bacteria to the levels of genera and species (Vandamme et al., 1996). In this study,
however, sequences of some isolates were similar to two different species. Constructing
and analyzing phylogenetic trees helped to precisely identify species of bacteria, as
reported by Park et al. (2005).
The composition of bacterial species identified from different horseradish
growing areas differed from each other. For example, Bacillus cereus, B. thuringiensis,
Pseudomonas fragi, and P. rhodesiae were dominant species in Illinois, whereas the
prevalent species in Wisconsin were Bacillus cereus, Pantoea agglomerans,
Pseudomonas grimontii, and P. extremorientalis. Additional studies are needed to
identify factors related to the existence of different species in different horseradish
growing areas.
Previous investigators reported the presence of Erwinia species and P. fluorescens
in internally discolored horseradish roots (Eastburn and Chang, 1994; Percich and
Johnson, 1990). In this study, neither Erwinia species nor P. fluorescens was identified.
This difference is likely related to the procedures used for identification of the species.
The previous investigators used non-molecular methods for identifying bacterial species
while molecular methods were employed in this study to identify the species.
78
FIGURES AND TABLES
Table 3.1 Horseradish roots tested for presence of the bacteria during 2008 and 2009
No. of roots
tested
No. of roots with No. of bacterial
bacteria (%)
isolates studiedb
Locationa
Cultivar
California
Czechoslovakia
80
57 (71)
93
Illinois
Big Top Western
72
67 (93)
145
(Collinsville)
1573
234
197 (84)
271
Ontario, Canada
1590 Mc
75
46 (61)
71
(Ancaster/Brantford)
1590 Wd
40
35 (88)
67
Wisconsin
Big Top Western
40
36 (90)
63
(Tule Lake)
(Eau Clair)
15K
80
71 (89)
114
from California, Illinois, and Ontario were collected from commercial fields. Roots
from Wisconsin were from research plots.
b
Two or more different bacterial isolates were present in some roots.
c
Roots collected from Ancaster, Ontario.
d
Roots collected from Brantford, Ontario.
a Roots
79
Table 3.2 Genera of bacteria isolated from internally discolored horseradish roots from different
horseradish growing areas during 2008-2009
Roots with bacteria (%)
Class
Actinobacteria
Bacilli
Alphaproteobacteria
Betaproteobacteria
Gammaproteobacteria
Genus
Achromobacter
California
2 (3.5)
14 (4.3)
2 (1.6)
1 (0.9)
Arthrobacter
Curtobacterium
Microbacterium
Plantibacter
Sanguibacter
-*
2 (3.5)
-
6 (1.9)
10 (7.8)
2 (1.6)
6 (4.7)
3 (2.3)
-
2 (1.9)
-
12 (21.2)
10 (17.5)
2 (3.5)
2 (3.5)
2 (3.5)
6 (10.5)
10 (17.5)
-
104 (32.2)
36 (11.1)
6 (1.9)
6 (1.9)
126 (39.0)
6 (1.9)
6 (1.9)
-
44 (34.1)
3 (2.3)
2 (1.6)
8 (6.2)
27 (20.9)
15 (11.6)
2 (1.6)
25 (23.4)
2 (1.9)
4 (3.7)
13 (12.1)
45 (42.1)
11 (10.3)
-
9 (15.8)
13 (4.0)
5 (3.9)
4 (3.7)
Bacillus
Paenibacillus
Agrobacterium
Ochrobactrum
Sphingobacterium
Variovorax
Enterobacter
Kluyvera
Pantoea
Pseudomonas
Serratia
Stenotrophomonas
Rahnella
Yersinia
Unidentified
* - = No bacterium was isolated.
80
Illinois
Ontario
Wisconsin
Table 3.3 Bacterial species isolated from horseradish roots from different horseradish growing areas during 2008 and 2009
Location
Class
Actinobacteria
Genus
Achromobacter
Arthrobacter
Curtobacterium
Microbacterium
Plantibacter
Bacilli
Sanguibacter
Bacillus
Paenibacillus
species
inoslitus
piechaudii
spanius
nicotinovorans
flaccumfaciens
hydrocarbonoxydans
profundi
xylanilyticum
auratus
flavus
suarezii
amyloliquefaciens
anthracis
atrophaeus
cereus
megaterium
thuringiensis
subtilis
amylolyticus
lautus
massiliensis
pabuli
tylopili
* - = No bacterium was isolated.
81
California
-*
2
2
12
10
Illinois
6
4
4
6
68
10
26
6
2
8
20
-
Ontario
2
10
2
1
4
1
2
1
12
17
9
4
2
3
-
Wisconsin
1
2
2
14
1
8
-
Table 3.3 (Cont.)
Class
Alphaproteobacteria
Betaproteobacteria
Gammaproteobacteria
Genus
Agrobacterium
Ochrobactrum
Sphingobacterium
Variovorax
Enterobacter
Kluyvera
Pantoea
Pseudomonas
Serratia
Stenotrophomonas
Rahnella
Yersinia
species
radiobacter
pseudogrignonense
kitahiroshimense
boronicumulans
paradoxus
asburiae
cowanii
numupressuralis
cryocrescens
agglomerans
eucalypti
vagans
brassicacearum
extremorientalis
fragi
gessardii
grimontii
libaniensis
marginalis
mediterranea
monteilii
proteolytica
rhodesiae
simiae
thivervalensis
proteamaculans
rhizophila
aquatica
kristensenii
82
California
2
2
2
4
2
10
-
Location
Illinois Ontario
6
1
1
6
3
1
4
2
26
54
1
4
3
1
6
2
18
22
12
2
6
6
15
2
Wisconsin
2
3
1
13
17
17
11
11
-
Table 3.4 Antifungal activity of bacterial isolates from discolored horseradish roots
Bacteria
Genus
Species
Achromobacter
Arthrobacter
Bacillus
spanius
nicotinovorans
amyloliquefaciens
anthracis
atrophaeus
cereus
megaterium
thuringiensis
subtilis
flaccumfaciens
asburiae
hydrocarbonoxydans
xylanilytiucm
lautus
pabuli
tylopili
agglomerans
vagans
flavus
extremorientalis
fragi
gessardii
grimontii
libaniensis
marginalis
mediterraneae
montelli
rhodesiae
thivervalensis
aquatica
proteamaculans
rhizophila
parardoxus
kristensenii
Curtobacterium
Enterobacter
Microbacterium
Paenibacillus
Pantoea
Plantibacter
Pseudomonas
Rahnella
Serratia
Stenotrophomonas
Variovorax
Yersinia
Verticillium
dahlaie
++ b
+++
++
++
+
++
+++
+
++
+
+
+
+
+
+
+
+
++
++
++
++
++
++
+
a
Fungia
Verticillium Fusarium
longisporum oxysporum
+
+
+++
++
++
+
++
+++
++
++
+
+
++
+
+
+
++
++
++
+++
++
+
+
+
+
++
+
+++
+
+++
++
+
++
+++
+
+
+
+
+
++
+
++
+
+
+
+
++
+++
+
++
+
+
-
Fusarium
solani
+++
+
+++
++
+
+
+++
+
++
+
+
+
++
+++
+
+
+
+
+
++
++
++
++
+
+
+
++
-
Causal agents of internal discoloration of horseradish roots.
Antifungial effects, calculated as the inhibition zone between the edge of fungal colony and the
edge of the bacterial colony [+++ = > 8 mm (strong inhibition); ++ = 2-7 mm (moderate
inhibition); + = < 2 mm (weak inhibition); - = (no inhibition)].
b
83
Cultivar Big Top Western, Illinois
Cultivar Czechoslovakia, California
Cultivar 1573, Illinois
Cultivar 1590, Ontario (Ancster)
Cultivar 1590, Ontario (Brantford)
Cultivar Big Top Western, Wisconsin
Figure 3.1
RFLP patterns of 16s rDNA of
bacterial isolates from different
horseradish cultivars from California,
Illinois, Ontario, and Wisconsin.
RFLP patterns resulted by ligasing 16s
rDNA by restriction enzyme Hae III.
Each lane represents different group of
isolated bacteria. Both ends are 1 kb
DNA molecular weight marker.
Cultivar 15K, Wisconsin
84
LH1-16S27F
70
Kluyvera cryocrescens ATCC 33435T
80
100 Enterobacter aerogenes NCTC 10006T
Klebsiella oxytoca JCM 1665T
Serratia grimesii DSM 30063T
67
77
LH15-16S27F
Pseudomonas simiae Oli T
58
80 LH10-16S27F
100 Pseudomonas trivialis DSM 14937T
64
Pseudomonas proteolytica CMS 64(T)
94
LH11-16S27F
67
Achromobacter spanius LMG 5911T
100 LH5-16S27F
Achromobacter piechaudii ATCC 43552T
60
Ochrobactrum thiophenivorans DSM 7216(T)
70
100 Ochrobactrum pseudogrignonense CCUG 30717T
79
LH17-16S27F
Sphingomonas adhaesiva GIFU 11458T
99
98
95
100
LH2-16S27F-22
Sphingobacterium kitahiroshimense 10C(T)
Sphingobacterium faecium DSM 11690T
57
96
Bacillus megaterium IAM 13418T
Bacillus subtilis subsp. subtilis NCDO 1769T
Bacillus thuringiensis gene for 16S IAM 12077T
97
5
7
LH7-16S27F
95 Paenibacillus tylopili strain MK2
98
64 LH4-16S27F
Paenibacillus amylolyticus NRRL NRS-290T
95
Microbacterium keratanolyticum IFO 13309(T)
99 Microbacterium profundi Shh49
LH9-16S27F
Planctomyces brasiliensis DSM 5305T
0.1
Figure 3.2.1 Phylogenetic relationships among bacterial isolates from horseradish
cultivar Czechoslovakia from California. The tree was constructed based on the partial
16S rDNA sequences by neighbor-joining method using Kimura‟s two-parameter
distance model. The bootstrap analysis was performed with 1,000 replications. The bar
represents 0.1 substitutions per site.
85
100 SC17-16S27F-48
Pseudomonas brassicacearum CFBP 11706T
Pseudomonas thivervalensis CFBP 11261(T)
SC6-16S27F-37
100 Pseudomonas fragi ATCC 4973T
SD1-16S27F-52
Pseudomonas psychrophila E-3(T)
100
Pseudomonas marginalis ATCC 10844T
64
SD3-16S27F-54
87
Pseudomonas grimontii CFML 97-514T
SC8-16S27F
100
Pseudomonas extremorientalis KMM 3447T
92 SC1-16S27F
Pseudomonas simiae Oli T
100 SC12-16S27F
Stenotrophomonas rhizophila DSM 14405T
87 Enterobacter nimipressuralis LMG 10245-T
100 SD12-16S27F
100
Enterobacter amnigenus JCM 1237(T)
100 Rahnella aquatica DSM 4594T
SD9-16S27F
50 Achromobacter insolitus LMG 6003T
SC10-16S27F
100 Achromobacter piechaudii ATCC 43552T
SD11-16S27F-62
97 Ochrobactrum pseudogrignonense CCUG 30717T
100
SD17-16S27F
Ochrobactrum thiophenivorans DSM 7216(T)
Paenibacillus pabuli JCM 9074(T)
SD2-16S27F
100 Paenibacillus taichungensis BCRC 17757(T)
SC18-16S27F-49
Paenibacillus amylolyticus NRRL NRS-290T
69
SD13-16S27F
100
100 Paenibacillus massiliensis 2301065(T)
SD8-16S27F
SC19-16S27F
100
Paenibacillus lautus DSM 5162(T)
Bacillus thuringiensis IAM 12077T
100 SD5-16S27F
Bacillus anthracis ATCC 14578T
64
100
SC2-16S27F
100 SC13-16S27F
99
Bacillus megaterium IAM 13418T
Bacillus flexus IFO 15715(T)
85 Sanguibacter suarezii ST50T
100 SD4-16S27F
Sanguibacter keddieii ST74
Planctomyces brasiliensis DSM 5305T
59
51
67
78
79
100
76
100
0.1
Figure 3.2.2 Phylogenetic relationships among bacterial isolates from horseradish
cultivars Big Top Western and 1573 from Illinois. The tree was constructed based on the
partial 16S rDNA sequences by neighbor-joining method using Kimura‟s two-parameter
distance model. The bootstrap analysis was performed with 1,000 replications. The bar
represents 0.1 substitutions per site.
86
53
100 CA13-16S27F-77
Pseudomonas mediterranea CFBP 5447T
Pseudomonas thivervalensis CFBP 11261(T)
CB24-27F-42
91
CB3-16S27F
84
Pseudomonas monteilii CIP 104883T
100 Pseudomonas grimontii CFML 97-514T
CB17-27F-37
100
100 CB11-27F-32
Pseudomonas libaniensis CIP 105460(T)
80
CA15-16S27F-79
Pantoea vagans LMG 24199T
100
Pantoea eucalypti LMG 24197
CA20-16S27F
83
100
100 CA16-16S27F
Rahnella aquatica DSM 4594T
99
98 CA10-16S27F
Yersinia kristensenii ATCC 33638T
100 CB14-27F-35
Pseudomonas geniculata ATCC 19374T
94 CA22-16S27F
100 Variovorax boronicumulans BAM-48(T)
CB10-27F-31
93 Variovorax paradoxus IAM 12373T
99 Plantibacter auratus IAM 18417(T)
100 CA12-16S27F
59 CB13-27F-34.
89
Plantibacter flavus DSM 14012T
100 CB22-27F-41
78
Microbacterium xylanilyticum S3-E(T)
100 CA6-16S27F
Microbacterium hydrocarbonoxydans DSM 16089T
100 62
CA21-16S27F-85
100 Arthrobacter nicotinovorans DSM 420T
100
CA7-16S27F
53
Arthrobacter methylotrophus DSM 14008T
100 CA4-16S27F-68
Curtobacterium flaccumfaciens pv. flaccumfaciens LMG 3645T
100 Bacillus amyloliquefaciens ATCC 23350T
CA2-16S27F-66
61
100 Bacillus anthracis ATCC 14578T
100
CA17-16S27F-81
100 Bacillus megaterium IAM 13418T
100
CA23-16S27F
Paenibacillus amylolyticus NRRL NRS-290T
100 CA5-16S27F
Paenibacillus pabuli JCM 9074(T)
Planctomyces brasiliensis DSM 5305T
0.1
67
87
100
Figure 3.2.3 Phylogenetic relationships among bacterial isolates from horseradish
cultivars 1590 from Ancaster and Brantford, Ontario (Canada). The tree was constructed
based on the partial 16S rDNA sequences by neighbor-joining method using Kimura‟s
two-parameter distance model. The bootstrap analysis was performed with 1,000
replications. The bar represents 0.1 substitutions per site.
87
100 WII-6
Pantoea agglomerans DSM 3493T
87
Pantoea ananatis LMG 2665T
100 Enterobacter asburiae JCM 6051T
WII4
60
92 Enterobacter cowanii CIP 107300(T)
80
98 WII-12
WI-16-16s27F-53
100 WI-9-16s27F-46
Rahnella aquatica DSM 4594T
94
74 WI-6-16s27F-43
Pseudomonas extremorientalis KMM 3447T
100
96
WII-11
Pseudomonas reinekei Mt-1(T)
83
Pseudomonas grimontii CFML 97-514T
WI-7-16s27F-44
69
100
79
Achromobacter spanius LMG 5911T
Achromobacter piechaudii ATCC 43552T
WII-5
WI-11-16s27F
77
99
Agrobacterium radiobacter ATCC 19358
Rhizobium sullae IS 123T
100
98
Bacillus thuringiensis IAM 12077T
WI5-16S27F
Bacillus anthracis Sterne
99
Bacillus cereus IAM 12605T
96
CA17-16S27F-81
100 Bacillus megaterium IAM 13418T
CA18-16S27F-82
WI-3-16s27F-40
98
100 Microbacterium hydrocarbonoxydans DSM 16089T
100
WI-15-16s27F-52
Microbacterium profundi Shh49
Planctomyces brasiliensis DSM 5305T
0.1
Figure 3.2.4 Phylogenetic relationships among bacterial isolates from horseradish
cultivars Big Top Western and 15K from Wisconsin. The tree was constructed based on
the partial 16S rDNA sequences by neighbor-joining method using Kimura‟s twoparameter distance model. The bootstrap analysis was performed with 1,000 replications.
The bar represents 0.1 substitutions per site.
88
LITERATURE CITED
Alstrom, S. 2000. Root-colonizing fungi from oilseed rape and their inhibition of
Verticillium dahliae. J. Phytopathol 148:417-423.
Atibalentja, N., and Eastburn, D.M. 1998. Verticillium dahliae resistance in horseradish
germ plasm from the University of Illinois collection. Plant Dis. 82:176-180.
Babadoost, M., Chen, W., Bratsch, A.D., and Eastman, C.E. 2004. Verticillium
longisporum and Fusarium solani: two new species in the complex of internal
discoloration of horseradish roots. Plant Pathol. 53:669-676.
Berg, G., Fritze, A., Roskot, N., and Smalla, K. 2001. Evaluation of potential biocontrol
rhizobacteria from different host plants of Verticillium dahliae Kleb. J. Appl. Microbiol.
91:963-971.
Chun J. 1995. Computer assisted classification and identification of actinomycetes. Ph.D.
Thesis. University of Newcastle. New Castle upon Tyne. UK.
Eastburn, D.M., and Chang, R.J. 1994. Verticillium dahliae: A causal agent of root
discoloration of horseradish in Illinois. Plant Dis. 78:496-498.
Hall, T.A. 1999. BioEdit: a user-friendly biological sequence alignment editor and
analysis program for Windows 95/98/NT. Nucleic Acid Symp Ser 41:95-98.
Felsenstein, J. 1985. Confidence limits on phylogenies: An approach using the bootstrap.
Evol. 39:783-791.
Kimura, M. 1980. A simple method for estimating evolutionary rates of base
substitutions through comparative studies of nucleotide sequences. J. Mol. Evol. 16:111120.
Lane, D.J. 1991. 16S/23S rRNA sequencing. pp 115-175. In Nucleic Acid Techniques in
Bacterial Systematics. E. Stackebrandt and M. Goodfellow (eds). J.Wiley and Sons Ltd.,
Chichester, UK. 329 pp.
Paul, N.C., Kim, W.K., Woo, S.K., Park, M.S., and Yu, S.H. 2007. Fungal endophytes in
roots of Aralia species and their antifungal activity. The Plant Pathol. J. 23:287-294.
Percich, J.A., and Johnson, D.R. 1990. A root rot complex of horseradish. Plant Dis.
74:391-393.
89
Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., and Higgins, D.G. 1997.
The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment
aided by quality analysis tools. Nucleic Acids Res. 25:4876-4882.
Vandamme, P.,Pot, B., Gillis, M., De Vos, P., and Kersters, K., and Swings, J. 1996.
Polyphasic taxonomy, a consensus approach to bacterial systematics, Microbiol. Rev.
60:407–438.
Ziedan, E.H.E. 2006. Manipulating Endophytic Bacteria for biological control to soil
borne diseases of peanut. J. Appl. Sci. Res. 2:497-502.
90