Identification of Fungi and Bacteria Associated with Internally Discolored Horseradish Roots

IDENTIFICATION OF FUNGI AND BACTERIA ASSOCIATED WITH INTERNALLY DISCOLORED HORSERADISH ROOTS

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 surface- disinfected 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

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.

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

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

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

(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 after- birth, 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).

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

(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;

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 set- borne 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.

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. 81:356- 362.

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.

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.

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.

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.

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)

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)

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.

(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 macro- and 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.

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 (PCT-

200, 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

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 PTC-

200 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.

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.

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 A-

PDA 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 2-

1). 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 2-

1).

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

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 2-

12). 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).

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.

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 2-14-

3). The isolates in this group fit the description of F. equiseti.

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 2-14-

4). 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 foot- shaped 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 2-

14-6). The isolates in this group fit the description of F. solani.

Molecular identification of Fusarium species

The ITS region of all isolates of Fusarium were amplified with primers ITS5-

ITS4, 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

(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 R-

6678, 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,

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

(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.

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

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.

FIGURES AND TABLES

Table 2.1 Detection of fungi in horseradish roots in 2008 and 2009

Roots with fungal isolates [no. (%)]b No. of No. of Other roots roots with Verticillium Fusarium Unknown All fungi identified a tested fungi (%) spp. spp. fungi Location Cultivar fungic 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. bTwo 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. dRoots collected from Ancaster, ON. eRoots collected from Brantford, ON.

Table 2.2 Nucleotide sequences of primers used for amplifying target genes of isolated fungi

Locus Primer Nucleotide sequence (5’-3’) Target fungi ITSa ITS 4 TCCTCCGCTTATTGATATGC All fungi ITS 5 GGAAGTAAAAGTCGTAACAAGG cox3b* COX 3A TGATTTAGAGATSTAATATCAGAAG Verticillium COX 3B CCGTGGAAACCTGTSCCAAAATA nad1c* NAD 1A ATGGCSAGTATGCAAAGAAGA Verticillium NAD 1B GCATGTTCTGTCATAAASCCACTAAC mtSSU- NMS1 CAGCAGTGAGGAATATTGGTCAATG Fusarium rDNAd NMS2 GCGGATCATCGAATTAAATAACAT

EF1- alphae† EF-1 ATGGGTAAGGARGACAAGAC Fusarium EF-2 GGARGTACCAGTRATCATGTT

a ITS1-5.8S rDNA-ITS2 region (Internal Transcribed Spacer). b 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

Table 2.3 Fungal genera isolated from internally discolored horseradish roots in 2008 and 2009

No. of isolates Phylum Genus California Illinois Ontario Wisconsin Total Ascomycota Alternaria 3 9 11 7 30 Arthopyreniaceae 3 - - - 3 Chaetomium - - 4 - 4 Cylindrocarpon 6 - - - 6 Colletotrichum - 21 - - 21 Cordyceps 1 - 15 - 16 Fusarium 28 100 26 75 229 Neonectria 1 - 8 - 9 Penicillium - 6 24 3 33 Phoma 1 - - - 1 Phomopsis 3 - - - 3 Pyrenochaeta 7 11 5 - 23 Rhizopycnis - 4 - - 4 Trichurus 4 - - - 4 Ulocladium 3 - - - 3 Verticillium 36 146 83 12 277

Basidiomycota Rhizoctonia 1 - - - 1

Zygomycota Rhizopus - - - 7 7

Table 2.4 Verticillium species isolated from horseradish roots in 2008 and 2009a

No. of Verticillium species [no. (%)] Location Verticillium VD groupb VL groupb isolates V. dahliae type Ac V. dahliae type Bc V. longisporum c California 36 0 (0) 2 (6) 34 (94) Illinois 146 121 (83) 21 (14) 4 (3) Ontario 83 77 (93) 6 (7) 0 (0) Wisconsin 12 12 (100) 0 (0) 0 (0) a The roots were cultured on potato dextrose agar (PDA) b Morphological characterization; VD=V. dahliae group, VL=V. longisporum group c Molecular identification

Table 2.5 Fusarium species isolated from horseradish roots in 2008 and 2009*

Fusarium species [no. (%)] No.of F. Location Fusarium F. F. F. F. F. equiseti isolates acuminatum commune oxysporum proliferatum solani -like species California 28 10 (35) 0 (0) 0 (0) 3 (10) 0 (0) 15 (55) Illinois 100 29 (29) 3 (3) 12 (12) 13 (13) 0 (0) 43 (43) Ontario 26 13 (50) 0 (0) 0 (0) 6 (23) 0 (0) 7 (27) Wisconsin 75 43 (57) 4 (5) 0 (0) 10 (13) 3 (4) 16 (21)

* The roots were cultured on potato dextrose agar (PDA)

Table 2.6 Occurrence of the internal root discoloration of horseradish plants (n=6) following inoculation with Verticillium species

Disease development Disease 1573a BTWa Fungus occurrence 1b 2b 3b 4b LSDc 1b 2b 3b 4b LSD

Verticillium dahliaed Incidencee 0.0 af 50.0 b 83.3 bc 100.0 c 41.4 0.0 a 50.0 b 83.3 b 83.3 b 47.9 (Type A) Severityg 0.0 a 1.8 ab 21.7 bc 37.6 c 16.3 0.0 a 10.2 a 18.3 a 37.9 b 18.6

Verticillium dahliaeh Incidence 16.7 a 66.7 ab 83.3 b 83.3 b 52.7 0.0 a 50.0 ab 66.7 b 83.3 b 51.2 (Type A) Severity 0.8 a 5.5 ab 19.5 bc 25.0 c 14.1 0.0 a 4.6 a 25.3 b 37.9 b 19.1

Verticillium dahliaei Incidence 33.3 a 66.7 ab 83.3 ab 100.0 b 50.4 16.7 a 50.0 ab 66.7 ab 100.0 b 51.6 (Type B) Severity 1.7 a 5.5 a 18.3 b 34.7 c 9.7 0.8 a 2.5 a 25.3 b 28.0 b 13.7

Verticillium longisporumj Incidence 0.0 a 33.3 a 83.3 b 100 b 39.6 0.0 a 33.3 a 83.3 b 100.0 b 39.6 Severity 0.0 a 6.0 a 21.7 b 31.3 b 12.1 0.0 a 3.8 ab 18.3 b 37.6 c 15.2

Control Incidence 0 0 16.7 16.7 NSk 0 0 0 16.7 NS Severity 0 0 0.8 0.8 NS 0 0 0 0.8 NS

a Horseradish cultivars (BTW=Big Top Western). b 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.

Table 2.7 Occurrence of the internal root discoloration and root rot of horseradish (n=12) following inoculation with Fusarium species Disease development Disease 1573a BTWa Fungus Disease occurrence 1b 2b 3b 4b LSDc 1b 2b 3b 4b LSD Fusarium IRDd Incidencee 16.7 af 66.7 b 66.7 b 91.7 b 34.9 25.0 a 33.3 a 50.0 a 91.7 b 34.6 acuminatum Severityg 0.8 a 5.5 ab 10.9 b 18.8 c 7.8 1.7 a 2.3 a 9.6 ab 16.7 b 8.7 Fusarium IRD Incidence 33.3 ab 58.3 b 58.3 b 0.0 a 36.2 33.3 25 33.3 16.7 NSh commune Severity 1.7 ab 8.9 bc 11.7 c 0.0 a 7.7 2.1 4 14.5 3 NS RRi Incidencej 0.0 a 16.7 ab 41.7 b 100.0 c 26.6 0.0 a 33.3 ab 50.0 ab 83.3 c 33.6 Fusarium IRD Incidence 0 8.3 16.7 16.7 NS 0 0 16.7 8.3 NS equiseti -like species Severity 0 6.4 1.9 1.9 NS 0 0 0.8 0.4 NS Fusarium IRD Incidence 16.7 a 58.3 b 83.3 c 91.7 53.9 16.7 a 41.7 ab 66.7 bc 83.3 c 37 oxysporum Severity 0.8 a 7.3 ab 15.1 b 25.4 c 8.7 0.8 a 6.4 a 12.3 a 33.7 b 14.6 Fusarium IRD Incidence 0 16.7 16.7 25 NS 0 0 16.7 16.7 NS proliferatum Severity 0 0.8 0.8 1.3 NS 0 0 0.8 0.8 NS Fusarium IRD Incidence 33.3 a 66.7 ab 66.7 ab 83.3 b 38.6 16.7 a 66.7 b 66.7 b 83.3 c 36.5 solani Severity 1.7 a 7.7 a 18.7 a 21.7 b 9.4 0.8 a 5.5 a 15.3 b 31.7 c 8.7 Control IRD Incidence 0 0 8.3 16.7 NS 0 0 0 8.3 NS Severity 0 0 0.4 0.8 NS 0 0 0 1.5 NS RR Incidence 0 0 0 0 NS 0 0 0 0 NS a 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.

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.

Figure 2.2 Frequency of fungal genera isolated from horseradish roots collected from California, Illinois, Ontario (Canada), and Wisconsin in 2008 and 2009

Figure 2.3 Percentage of the fungal isolates identified as Verticillium and Fusarium species in horseradish growing areas in 2008 and 2009.

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.

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.

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.

ITS SDV1025 65 637 characters SDV1002 90 parsimony-informative SDV1006 characters 1000 trees V. dahliae M5 (AY555948) 313 steps V. dahliae UZ132 (AY555949) CI = 0.8179 CBV1046 RI = 0.7957 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) V. dahliae Vd292 (AF363994) 67 CBV2034 SCV2008 52 SDV2066 SDV2064 SDV2041 100 V. albo-atrum V90 (AY555954) V. albo-atrum 1776 (AY555953) 65 67 V. nubilum 278734 94 V. tricorpus 22 56 V. tricorpus188

100 V. nigrescens 193 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.

cox3 V. dahliae M5 (AY555921) 376 characters V. dahliae UZ132 (AY555922) 86 parsimony-informative SDV1025 characters 15 trees SDV1002 249 steps SDV1006 CI = 0.6466 RI = 0.6318 CBV1046

CBV1014 dahliae V. CBV1006

60 CAV1004 CAV1003

SCV1010 SCV1005 V. dahliae T9 (AY555920) CBV2034 SCV2008 SDV2066 SDV2064

93 SDV2041 V. longisporum G19 (AY555923)

V. longisporum K2 (AY555924) longispor V. V. longisporum K12 (AY555925) TBV3014 TAV3001 90

VBV3004 um

VAV3021 VAV3003 64 58 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

78 V. catenulaulatum 113078 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.

V. dahliae M5 (AY555977) nad1 523 characters V. dahliae UZ132 (AY555978) 120 parsimony-informative SDV1025 characters SDV1002 dahliae V.

4 trees A Type 385 steps SDV1006 CI = 0.7143 64 CBV1046 RI = 0.6927 CBV1014

CBV1006 CAV1004 CAV1003 SCV1010 SCV1005 V. dahliae T9 (AY555976) dahliae V.

Type Type CBV2034 SCV2008

97 B

SDV2066

SDV2064 SDV2041

V. longisporum G19 (AY555979) longisporum V. 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 V. tricorpus 22 56 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.

V. dahliae M5 cox3 + nad1 899 characters V. dahliae UZ132 206 parsimony-informative SDV1025 characters SDV1002 3 trees 637 steps SDV1006 CI = 0.6845 63 CBV1046 RI = 0.6683 A Type CBV1014 dahliae V. CBV1006

CAV1004 CAV1003

68 SCV1010 SCV1005 V. dahliae T9

CBV2034 Type SCV2008

SDV2066 B

SDV2064 100 SDV2041 V.longisporum G19

V. longisporum V. V. longisporum K2 V. longisporum K12

83 TBV3003 TAV3001 100 70 VBV3004

VAV3021 VAV3003

98 V. albo-atrum V90 73 V. albo-atrum 1776

100 V. tricorpus 22 74 V. tricorpus188 77 . nubilum 278734 V. theobromae 225818 Acrostalagmus luteo-albus 868 V. nigrescens 193

100 V. catenulaulatum 113078 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.

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.

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.

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.

SD5115 ITS region 429 characters 100 SD5030 80 parsimony- 67 WA5039 informative characters 696 trees 97 F. equiseti (AB425996) 163 steps F. cerealis NRRL25491 (AF006340) CI = 0.7914 83 RI = 0.9470 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

99 SC1042 100 SC1013

61 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.

NMS 562 characters 59 parsimony- informative characters 4 trees F. commune 9245H (DQ016145) 168 steps CI = 0.9167 F. redolens NRRL28058 (AF324293) 92 RI = 0.9381 SD4139 WA4023 WB4031

100 F. proliferatum 31X4 (DQ831947) WA6005 F. oxysporum f. sp. melonis TX388 (DQ831943)

85 SD3152 CA3022 93 CA3008 WA3037 SD5115 100 SD5030 WA5039

100 F. cerealis (U85551) F. venenatum (U85560) F. tricinctum NZ62 (EU919401) 70 F. acuminatatum NZ50 (EU919400) 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.

EF1-alpha F. oxysporum f. sp. melonis TX388(DQ837696) 380 characters 60 116 parsimony- SD3152 informative characters 95 CA3008 59 trees 100 344 steps CA3022 CI = 0.8052 WA3037 71 RI = 0.9193 F. commune NRRL28387 (AF246832) 64 WB4031 98 92 SD4139 WA4023 53 F. proliferatum NRRL43666(EF452997) WA6005 F. redolens NRRL28412 (AF324300) 100 WA1072 CA1018 58 SC1018 SD1184 F. solani f. sp. batatas NRRL22402 (AF178344) 91 SD1212

100 CB1005

68 N. haematococca MAFF 840047 (DQ452423) 54 59 SC1013 F. solani NRRL20547 (DQ247544) SC1042

62 SD5115 97 SD5030 94 WA5039 71 F. equiseti DAOM236361 (DQ842099)

88 F. cerealis NRRL13393 (AF212467) F. venenatum VI01179 (AJ543635) F. acuminatum R-6678 (FJ154737) SD2018 96 WA2077 99 CA2060 F. tricinctum PDD17HB (EU327337) C. cylindroides CBS 324.61 5 changes (DQ789732)

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.

F. commune ITS / NMS / EF 1-alpha 64 1422 characters WB4031 95 249 parsimony- F. commune informative characters SD4139 18 trees WA4023 715 steps CI = 0.8028 54 F. redolens RI = 0.9126 100 F. proliferatum F. proliferatum WA6005

62 SD3152 64 CA3008 100 F. oxysporum F. oxysporum 100 CA3022 complex

WA3037

WA1072 60 CA1018 94 SC1018

73 SD1184 SD1212 F. solani

80 CB1005 complex 100 85 SC1042 50 SC1013

58 F. solani

N. haematococca

F. acuminatum

94 SD2018 WA2077 F. acuminatum 100 CA2060

F. tricinctum

61 SD5115 100 SD5030 F. sp.

100 WA5039

99 F. cerealis 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.

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., Eranthodi, A., Jurgens, A., Hippard, K., and Wahle, E. 2007. Thermo- Therapy 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 albo- atrum 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 3-

3). 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 3-2-

1). 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 No. of roots with No. of bacterial Locationa Cultivar tested bacteria (%) isolates studiedb

California Czechoslovakia 80 57 (71) 93 (Tule Lake) 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 (Eau Clair) 15K 80 71 (89) 114 a Roots 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.

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 Genus California Illinois Ontario Wisconsin Actinobacteria Achromobacter 2 (3.5) 14 (4.3) 2 (1.6) 1 (0.9) Arthrobacter - * - 10 (7.8) - Curtobacterium - - 2 (1.6) - Microbacterium 2 (3.5) - 6 (4.7) 2 (1.9) Plantibacter - - 3 (2.3) - Sanguibacter - 6 (1.9) - - Bacilli Bacillus 12 (21.2) 104 (32.2) 44 (34.1) 25 (23.4) Paenibacillus 10 (17.5) 36 (11.1) 3 (2.3) - Alphaproteobacteria Agrobacterium - - - 2 (1.9) Ochrobactrum 2 (3.5) 6 (1.9) - - Betaproteobacteria Sphingobacterium 2 (3.5) - - - Variovorax - - 2 (1.6) - Gammaproteobacteria Enterobacter - 6 (1.9) - 4 (3.7) Kluyvera 2 (3.5) - - - Pantoea - - 8 (6.2) 13 (12.1) Pseudomonas 6 (10.5) 126 (39.0) 27 (20.9) 45 (42.1) Serratia 10 (17.5) - - - Stenotrophomonas - 6 (1.9) - - Rahnella - 6 (1.9) 15 (11.6) 11 (10.3) Yersinia - - 2 (1.6) - Unidentified 9 (15.8) 13 (4.0) 5 (3.9) 4 (3.7) * - = No bacterium was isolated.

80 Table 3.3 Bacterial species isolated from horseradish roots from different horseradish growing areas during 2008 and 2009

Location Class Genus species California Illinois Ontario Wisconsin Actinobacteria Achromobacter inoslitus - * 6 - - piechaudii - 4 - 1 spanius 2 4 2 - Arthrobacter nicotinovorans - - 10 - Curtobacterium flaccumfaciens - - 2 - Microbacterium hydrocarbonoxydans - - 1 2 profundi 2 - 4 - xylanilyticum - - 1 - Plantibacter auratus - - 2 - flavus - - 1 - Sanguibacter suarezii - 6 - - Bacilli Bacillus amyloliquefaciens - - 12 - anthracis - - 17 2 atrophaeus - - 9 - cereus - 68 - 14 megaterium - 10 4 1 thuringiensis 12 26 - 8 subtilis - - 2 - Paenibacillus amylolyticus - 6 3 - lautus - 2 - - massiliensis - 8 - - pabuli - 20 - - tylopili 10 - - -

* - = No bacterium was isolated.

81 Table 3.3 (Cont.)

Location Class Genus species California Illinois Ontario Wisconsin Alphaproteobacteria Agrobacterium radiobacter - - - 2 Ochrobactrum pseudogrignonense 2 6 - - Betaproteobacteria Sphingobacterium kitahiroshimense 2 - - - Variovorax boronicumulans - - 1 - paradoxus - - 1 - Gammaproteobacteria Enterobacter asburiae - - - 3 cowanii - - - 1 numupressuralis - 6 - - Kluyvera cryocrescens 2 - - - Pantoea agglomerans - - 3 13 eucalypti - - 1 - vagans - - 4 - Pseudomonas brassicacearum - 2 - - extremorientalis - 26 - 17 fragi - 54 - - gessardii - - 1 - grimontii - 4 3 17 libaniensis - - 1 - marginalis - 6 - - mediterranea - - 2 - monteilii - - 18 - proteolytica 4 - - - rhodesiae - 22 - - simiae 2 - - - thivervalensis - 12 2 11 Serratia proteamaculans 10 - - - Stenotrophomonas rhizophila - 6 - - Rahnella aquatica - 6 15 11 Yersinia kristensenii - - 2 -

82 Table 3.4 Antifungal activity of bacterial isolates from discolored horseradish roots

a Bacteria Fungi Verticillium Verticillium Fusarium Fusarium Genus Species dahlaie longisporum oxysporum solani Achromobacter spanius ++ b + + - Arthrobacter nicotinovorans - + - - Bacillus amyloliquefaciens +++ +++ +++ +++ anthracis ++ ++ + + atrophaeus ++ ++ +++ +++ cereus + + ++ ++ megaterium - - + + thuringiensis ++ ++ ++ + subtilis +++ +++ +++ +++ Curtobacterium flaccumfaciens - - - + Enterobacter asburiae + ++ + ++ Microbacterium hydrocarbonoxydans ++ ++ + + xylanilytiucm + + + + Paenibacillus lautus + + + + pabuli + ++ + ++ tylopili + - ++ +++ Pantoea agglomerans + + + + vagans + + ++ + Plantibacter flavus + - + + Pseudomonas extremorientalis - + - + fragi + - + + gessardii - - - - grimontii ++ ++ + ++ libaniensis - ++ - ++ marginalis - - + - mediterraneae ++ ++ ++ ++ montelli ++ +++ +++ ++ rhodesiae ++ ++ + + thivervalensis - + ++ + Rahnella aquatica ++ + - + Serratia proteamaculans ++ + + ++ Stenotrophomonas rhizophila - - + - Variovorax parardoxus - + - - Yersinia kristensenii + ++ - -

a Causal agents of internal discoloration of horseradish roots. bAntifungial 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)].

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 80 Kluyvera cryocrescens ATCC 33435T

100 Enterobacter aerogenes NCTC 10006T

Klebsiella oxytoca JCM 1665T

Serratia grimesii DSM 30063T

67 LH15-16S27F 77 Pseudomonas simiae Oli T 58 80 LH10-16S27F 100 Pseudomonas trivialis DSM 14937T

64 Pseudomonas proteolytica CMS 64(T) 94 LH11-16S27F

Achromobacter spanius LMG 5911T 67 100 LH5-16S27F

60 Achromobacter piechaudii ATCC 43552T Ochrobactrum thiophenivorans DSM 7216(T) 70 100 Ochrobactrum pseudogrignonense CCUG 30717T 79 LH17-16S27F

Sphingomonas adhaesiva GIFU 11458T 99 LH2-16S27F-22 98 95 100 Sphingobacterium kitahiroshimense 10C(T) Sphingobacterium faecium DSM 11690T

Bacillus megaterium IAM 13418T 57 96 Bacillus subtilis subsp. subtilis NCDO 1769T Bacillus thuringiensis gene for 16S IAM 12077T 97 LH7-16S27F 5 7 95 Paenibacillus tylopili strain MK2 98 64 LH4-16S27F Paenibacillus amylolyticus NRRL NRS-290T

Microbacterium keratanolyticum IFO 13309(T) 95 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 59 Pseudomonas brassicacearum CFBP 11706T 51 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 67 SC8-16S27F 100 Pseudomonas extremorientalis KMM 3447T 92 SC1-16S27F Pseudomonas simiae Oli T 78 100 SC12-16S27F Stenotrophomonas rhizophila DSM 14405T 87 Enterobacter nimipressuralis LMG 10245-T 100 SD12-16S27F 100 79 Enterobacter amnigenus JCM 1237(T) 100 Rahnella aquatica DSM 4594T SD9-16S27F 50 Achromobacter insolitus LMG 6003T 100 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) 76 Bacillus thuringiensis IAM 12077T 100 SD5-16S27F Bacillus anthracis ATCC 14578T 64 100 SC2-16S27F 100 100 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 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.

86 100 CA13-16S27F-77 53 Pseudomonas mediterranea CFBP 5447T 67 Pseudomonas thivervalensis CFBP 11261(T) CB24-27F-42 91 CB3-16S27F 84 Pseudomonas monteilii CIP 104883T 100Pseudomonas grimontii CFML 97-514T 100 CB17-27F-37 100 CB11-27F-32 80 Pseudomonas libaniensis CIP 105460(T) CA15-16S27F-79 100 Pantoea vagans LMG 24199T Pantoea eucalypti LMG 24197 CA20-16S27F 83 100 100 CA16-16S27F 99 Rahnella aquatica DSM 4594T 98 CA10-16S27F 87 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-68 100 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

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 87 Pantoea agglomerans DSM 3493T Pantoea ananatis LMG 2665T

100 Enterobacter asburiae JCM 6051T 60 WII4 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 WII-11 96 100 83 Pseudomonas reinekei Mt-1(T) Pseudomonas grimontii CFML 97-514T WI-7-16s27F-44

69 Achromobacter spanius LMG 5911T 79 100 Achromobacter piechaudii ATCC 43552T WII-5 WI-11-16s27F 77 99 Agrobacterium radiobacter ATCC 19358 Rhizobium sullae IS 123T Bacillus thuringiensis IAM 12077T 100 98 WI5-16S27F

99 Bacillus anthracis Sterne 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 two- parameter distance model. The bootstrap analysis was performed with 1,000 replications. The bar represents 0.1 substitutions per site.

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.

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. 16:111- 120.

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.

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.