Genetics, Genomics and Breeding of Late Blight and Early Blight Resistance in Tomato

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Genetics, Genomics and Breeding of Late Blight and Early Blight Resistance inTomatoMajid R. Foolad a; Heather L. Merk a; Hamid Ashrafi a

a Department of Horticulture and The Intercollege Graduate Degree Program in Genetics, The PennsylvaniaState University, University Park, PA, USA

Online Publication Date: 01 March 2008

To cite this Article Foolad, Majid R., Merk, Heather L. and Ashrafi, Hamid(2008)'Genetics, Genomics and Breeding of Late Blight andEarly Blight Resistance in Tomato',Critical Reviews in Plant Sciences,27:2,75 — 107

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Critical Reviews in Plant Sciences, 27:75–107, 2008Copyright c© Taylor & Francis Group, LLCISSN: 0735-2689 print / 1549-7836 onlineDOI: 10.1080/07352680802147353

Genetics, Genomics and Breeding of Late Blight and EarlyBlight Resistance in Tomato

Majid R. Foolad, Heather L. Merk, and Hamid AshrafiDepartment of Horticulture and The Intercollege Graduate Degree Program in Genetics, ThePennsylvania State University, University Park, PA 16802 USA

Referees: Dr. Dennis R. Decoteau, Professor, Departments of Horticulture and Plant Pathology, The Pennsylvania State University, University Park,PA 16802 USADr. David R. Huff, Associate Professor, Department of Crop and Soil Sciences, The Pennsylvania State University, University Park, PA16802 USADr. David M. Geiser, Associate Professor, Department of Plant Pathology, The Pennsylvania State University, University Park, PA 16802USA

Table of Contents

I. INTRODUCTION ..............................................................................................................................................76

II. LATE BLIGHT ..................................................................................................................................................77A. Background ..................................................................................................................................................77B. Disease Symptoms ........................................................................................................................................78C. The Pathogen ...............................................................................................................................................79

1. Disease Cycle .......................................................................................................................................792. Clonal Lineage .....................................................................................................................................793. Possible Mechanism of Pathogenicity .....................................................................................................80

D. Possible Mechanisms of Host Resistance ........................................................................................................80E. Disease Screening Methods ...........................................................................................................................81

1. Inoculum Preparation ............................................................................................................................812. Field Screening .....................................................................................................................................813. Greenhouse Screening ...........................................................................................................................824. Growth Chamber Screening Using Detached Leaflets ...............................................................................82

F. Nongenetic Measures of Disease Control ........................................................................................................821. Cultural Practices ..................................................................................................................................822. Chemical Applications ..........................................................................................................................83

G. Genetic Measures of Disease Control .............................................................................................................831. Late Blight Resistance in Potato .............................................................................................................832. Late Blight Resistance in Tomato ...........................................................................................................86

III. EARLY BLIGHT ...............................................................................................................................................87A. Background ..................................................................................................................................................87B. Disease Symptoms ........................................................................................................................................88C. The Pathogen ...............................................................................................................................................88

1. Disease Cycle .......................................................................................................................................882. Variability of Alternaria Isolates and Causes of Variability .......................................................................89

a. Variability in Morphology ..........................................................................................89b. Variability at the Molecular Level ...............................................................................89c. Variability in Toxin Production ...................................................................................89

3. Physiological Races ..............................................................................................................................904. Mechanisms of Pathogenicity .................................................................................................................90

Address correspondence to Prof. Majid R. Foolad, Department of Horticulture, The Pennsylvania State University, University Park, PA 16802USA. E-mail: [email protected]

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76 M. R. FOOLAD ET AL.

a. Enzymatic Reactions .................................................................................................90b. Toxin Production ......................................................................................................90

D. Mechanisms of Host Resistance .....................................................................................................................91E. Disease Screening Methods ...........................................................................................................................92

1. Inoculum Preparation ............................................................................................................................922. Field Screening .....................................................................................................................................923. Greenhouse Screening ...........................................................................................................................924. Growth Chamber Screening Using Detached Leaflets ...............................................................................93

F. Nongenetic Measures of Disease Control ........................................................................................................93G. Genetic Measures of Disease Control .............................................................................................................93

1. Genetic Sources of Resistance ................................................................................................................93a. Solanum lycopersicum (formerly Lycopersicon esculentum) ..........................................93b. Solanum habrochaites (formerly Lycopersicon hirsutum) ..............................................94c. Solanum pimpinellifolium (formerly Lycopersicon pimpinellifolium) .............................94d. Solanum peruvianum (formerly Lycopersicon peruvianum) ..........................................95

2. Inheritance Studies ................................................................................................................................953. Genetic Mapping ..................................................................................................................................95

a. S. lycopersicum × S. habrochaites ..............................................................................96b. S. lycopersicum × S. pimpinellifolium .........................................................................96c. S. lycopersicum × S. peruvianum ...............................................................................99

4. Breeding for Early Blight Resistance ......................................................................................................99

IV. CONCLUSION AND FUTURE PROSPECTS .................................................................................................. 100A. Late Blight ................................................................................................................................................. 100B. Early Blight ............................................................................................................................................... 100

REFERENCES .......................................................................................................................................................... 101

Late blight (LB), caused by the oomycete Phytophthora infes-tans, and early blight (EB), caused by the fungi Alternaria solaniand A. tomatophila, are two common and destructive foliar dis-eases of the cultivated tomato (Solanum lycopersicum) and potato(Solanum tuberosum) in the United States and elsewhere in theworld. While LB can infect and devastate tomato plants at any de-velopmental stages, EB infection is usually associated with plantphysiological maturity and fruit load where older senescing plantsexhibit greater susceptibility and a heavy fruit set enhances the dis-ease. At present, cultural practices and heavy use of fungicides arethe most common measures for controlling LB and EB. Genetic re-sources for resistance have been identified for both diseases, largelywithin the tomato wild species, in particular the red-fruited speciesS. pimpinellifolium and the green-fruited species S. habrochaites. Afew race-specific major resistance genes (e.g., Ph-1, Ph-2 and Ph-3)and several race-nonspecific resistance QTLs have been reportedfor LB. Ph-3 is a strong resistance gene and has been incorporatedinto many breeding lines of fresh market and processing tomato.However, new P. infestans isolates have been identified which over-come Ph-3 resistance. Recently, a new resistance gene (Ph-5) hasbeen identified, which confers resistance to several pathogen iso-lates including those overcoming the previous resistance genes.Advanced breeding lines including Ph-5 alone and in combina-tions with Ph-2 and Ph-3 are being developed. Genetic controls ofEB resistance have been studied and advanced breeding lines andcultivars with improved resistance have been developed throughtraditional breeding. Additionally, QTLs for EB resistance havebeen identified, which can be utilized for marker-assisted resistancebreeding. Currently, new inbred lines and cultivars of tomato with

good levels of EB resistance and competitive yield performance arebeing developed at the Pennsylvania State University. This reviewwill focus on the current knowledge of both LB and EB with respectto the causal pathogens, host resistances, and genetics and breedingprogresses.

Keywords disease resistance, horizontal resistance, pathogenicity, R-gene, QTL, Solanum lycopersicum, S. pimpinellifolium,vertical resistance

I. INTRODUCTIONThe cultivated tomato, Solanum lycopersicum L. (formerly

Lycopersicon esculentum Mill.), a fruit that is often treated as avegetable, is the second most consumed vegetable in the worldafter potato (FAOSTAT, 2005). It is the third most economi-cally important vegetable (with a total farm value of $2.062B) in the U.S. following potato ($2.564 B) and lettuce ($2.064B) (http://www.usda.gov/nass/pubs/agr05/agstats2005.pdf). Inaddition to tomatoes that are consumed as raw vegetables oradded to other food items, a variety of processed products suchas pastes, whole peeled tomatoes, diced products, and variousforms of juice, sauces, and soups have gained significant ac-ceptance. Although a tropical plant, the tomato is grown in al-most every region of the world, from the tropics to within a fewdegrees of the Arctic Circle. When outdoor production is

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BLIGHT RESISTANCE IN TOMATO 77

restricted due to cold temperatures, the tomato is grown ingreenhouses. Major tomato-producing countries (in descend-ing order of tonnage) include China, the U.S., India, Turkey,Egypt, and Italy (http://faostat.fao.org/). Other leading coun-tries include Spain, Brazil, Iran, Mexico, Greece, and Russia.Tomatoes are an important part of a diverse and balanced diet(Willcox et al., 2003). Although the tomato does not rank high innutritional value, by virtue of volume consumed, it contributessignificantly to the dietary intake of vitamins A and C, as wellas essential minerals and other nutrients. For example, tomatoranks first among all fruits and vegetables as a source of vitaminsand minerals in the U.S. diet (Rick, 1980) and phenolic antiox-idants (Vinson et al., 1998). In addition, fresh and processedtomatoes are the richest sources of the antioxidant lycopene(Nguyen and Schwartz, 1999), which arguably protects cellsfrom oxidants that have been linked to cancer (Giovannucci,1999).

Diseases are the number one concern to processing and freshmarket tomato industries throughout the world. The tomato issusceptible to over 200 diseases caused by pathogenic fungi, bac-teria, viruses, and nematodes (Lukyanenko, 1991). The greatestcontribution of modern plant breeding to tomato improvementhas been the development of cultivars with improved diseaseresistance. Resistance has been identified, and in many cases,characterized for more than 30 major tomato diseases. Mostcommercial cultivars possess up to 6 (in true breeding lines) or10 (in hybrids) disease-resistance attributes. Wild tomato specieshave been utilized as the source of disease resistance, except ina few cases. Resistance sources have been identified mainly inwild species S. pimpinellifolium L. [formerly L. pimpinellifolium(L.) Mill.], S. peruvianum L. [formerly L. peruvianum (L.) Mill.]and S. habrochaites S. Knapp & D. Spooner (formerly L. hirsu-tum Dunal).

The original characterization, disease evaluation, and incor-poration of resistance genes were conducted using phenotypicselection and traditional breeding protocols. Contemporary dis-ease resistance breeding is done primarily through the use ofsimilar protocols. However, the difficulties encountered whentransferring resistance from wild species to the cultivated tomatovia traditional protocols have restricted transfer of resistanceto many elite tomato lines. This is in addition to the inade-quate screening facilities and expertise in many tomato breed-ing programs to develop cultivars with multiple disease resis-tances. Thus, breeders have consistently sought more effectiveapproaches for resistance breeding.

During the past two decades, the use of molecular markersand marker-assisted selection (MAS) techniques have facili-tated the processes of identifying, mapping, and transferringof disease resistance genes and quantitative trait loci (QTLs).Furthermore, breeding for disease resistance remains a majorgoal of most public and private tomato breeding programs asnew diseases achieve significance and new races of existingpathogens become established. The ultimate goal of breedingprograms with respect to disease is to eliminate or significantly

reduce the use of chemicals in tomato production through theuse of host resistance.

Knowledge of the genetic diversity of a pathogen is bene-ficial to the understanding of its evolutionary past, as well asits future (McDonald and Linde, 2002). Such knowledge canbe used to implement successful strategies for host resistancebreeding. For example, McDonald and Linde (2002) proposed adecision-making model for choosing resistance-breeding strate-gies based on the knowledge of pathogen variability. Accordingto their model, asexually reproducing pathogens, such as EB,pose a low risk to breakdown of genetic resistance in host plants.Conversely, for sexually reproducing pathogens with high geneflow in the population, a breeding strategy involving major genepyramiding (MGP) may be more effective in developing durableresistance.

Two of the most common and destructive foliar diseases oftomato worldwide are late blight (LB) and early blight (EB).Both diseases have the potential to drastically damage the tomatoplant, render fruit unmarketable, reduce yield and cause signifi-cant economic losses. Currently, both diseases are primarily con-trolled through cultural practices and heavy use of fungicides,however, the use of genetic resistance has been sought by tomatobreeders and the industry. To date, simple and/or complex re-sistance to both diseases have been identified and characterized,and considerable progress has been made toward developmentof resistant tomatoes. In this review, we provide an overviewof these two diseases, focusing on the biology of the pathogensand their mechanisms of pathogenicity, mechanisms of host re-sistance, and genetics and breeding efforts toward developingtomato breeding lines and cultivars with improved resistance.

II. LATE BLIGHT

A. BackgroundLate blight (LB), caused by the oomycete Phytophthora in-

festans (Mont.) de Bary, occurs throughout tomato and potatogrowing regions of North America and elsewhere in the worldwith varying frequency, depending on weather conditions. LBhas been identified as a major disease of tomato and potato andis one of the most devastating plant diseases for four reasons(Fry and Goodwin, 1997b). First, LB can destroy tomato andpotato crops within several days of occurrence in the field. Sec-ond, low levels of P. infestans are difficult to detect in the fieldand by the time the disease is detected it is often too late to savethe crop through fungicide application. This is largely becausemost current P. infestans isolates are resistant to metalaxyl, theonly available systemic fungicide to combat the disease. Third,each LB lesion can produce as many as 300,000 sporangia perday, contributing to a quick spread of the disease. Finally, theasexual disease cycle consisting of pathogen penetration, colo-nization, sporulation, and dispersal can occur in fewer than fivedays (Fry and Goodwin, 1997b). In the United States, LB hasbeen ranked as the eight most important disease of tomato basedon a weighted average crop loss per acre (Davis et al., 2000). It

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78 M. R. FOOLAD ET AL.

has been suggested that potato yield would increase by 5% andpotato storage loss would decrease by 17% if LB did not exist(Guenthner et al., 2001).

The origin of LB is not precisely known, with the major theo-ries proposing that it originated in the South American Andes orthe Toluca Valley of Central Mexico (Bourke, 1964; Andrivon,1996; Grunwald and Flier, 2005; Gomez-Alpizar et al., 2007).The Andean theory was first suggested by de Bary in the 19thcentury, shortly after the Irish potato famine (de Bary, 1876). deBary and other proponents of this theory argued that P. infestansoriginated in the same place as its hosts, the potato and otherSolanaceous species. More recently, isozyme and pathogenicitysimilarities among Peruvian, American and European isolatesof P. infestans also suggest a common origin for the host andpathogen populations (Tooley et al., 1989). However, a lack ofdiversity in Andean populations compared to Mexican popula-tions led few to support this theory (Tooley et al., 1989; Perezet al., 2001). As early as 1939, the Andean theory was reputedby arguing that wild potato species were also found outside theAndes, there were wild potato species in Mexico resistant toLB, and that species other than potato could serve as the hostfor P. infestans (Reddick, 1939). With the advent of molecularmarkers and DNA fingerprinting, the comparatively high geneticdiversity of P. infestans in Mexico further suggested a Mexicanorigin (Goodwin et al., 1994), though recent sequence analysisof two nuclear genes and three mitochondrial genes supports theAndean theory (Gomez-Alpizar et al., 2007).

LB was first identified outside of Central and South Americain 1843, when it appeared in the northeastern U.S., and wasfirst identified in Europe in June 1845, when it was reportedin Belgium (Bourke, 1964). LB was reported in France and theNetherlands soon after, and by September 1845 it was reported inIreland marking the beginning of the Irish potato famine, whichled to the deaths of over one million people and emigration ofa similar number, mostly to the east coast of the U.S. (Ristaino,2002).

Until the late 1970s, LB was relatively well controlledthrough the use of cultural practices, heavy application of fungi-cides, and the use of somewhat resistant potato cultivars. How-ever, in the early 1980s in Europe and late 1980s in NorthAmerica, LB re-emerged as an important disease of potato andtomato. LB re-emergence causes two major concerns (Fry andGoodwin, 1997b). First, prior to the 1980s, only the A1 matingtype was observed outside Mexico, so sexual reproduction didnot play a significant role in the disease cycle. Phytophthorainfestans is a heterothallic organism with two known matingtypes, A1 and A2, both of which are required for sexual repro-duction (Judelson, 1997). Early in the 1980s, A2 mating typewas identified outside Mexico (Fry and Goodwin, 1997a; b),first in Europe in 1984 and then in North America in 1991 (Hohland Iselin, 1984; Deahl et al., 1991). The presence of the A2mating type outside Mexico created the opportunity for sex-ual reproduction and creation of new, more aggressive isolates.This concern was realized with the appearance of the US-11

clonal lineage, a highly aggressive sexually derived lineage thathas devastated tomato crops in the Pacific Northwest, the NorthEast (mainly New York), and California (Gavino et al., 2000). Inaddition to creating new isolates, sexual reproduction producesoospores. Unlike asexual spores, known as zoospores, oosporescan overwinter in the field and survive well under harsh condi-tions. Oospores are large, thick-walled spores, which enable thefungus to survive in plant debris or soil outside the living hostplant. A concern is that the oospores may become an inoculumsource for the next year crop (Gavino et al., 2000).

The second concern associated with LB re-emergence is thatmany new, complex lineages exhibit metalaxyl resistance. Theappearance of metalaxyl resistance nearly coincided with theobservation of the A2 mating type outside Mexico, however,no genetic correlation has been documented between matingtype and metalaxyl resistance (Gisi and Cohen, 1996). Meta-laxyl resistance poses great threat to tomato and potato growersas these are the only available systemic fungicides (Gisi andCohen, 1996). Systemic fungicides, also known as therapeuticfungicides, inhibit or reduce disease progress once symptomsare apparent. This is in contrast to protectant fungicides, whichare applied before the presence of disease symptoms. Presenceof metalaxyl resistance suggests that if LB is observed, it islikely too late to use protectant fungicides to save the crop. Tocombat problems associated with metalaxyl resistance, the in-dustry began distributing these fungicides only in combinationwith at least one other fungicide with a different mode of ac-tion (Russell, 2005). The second fungicide is often mancozeb orchlorothalonil. Metalaxyl has also been replaced by an opticalisomer, metalaxyl-m (mefenoxam) (Russell, 2005). However,the concern with fungicide resistance has urged tomato breed-ers to look for sources of host genetic resistance, as discussedbelow.

B. Disease SymptomsPhytophthora infestans can quickly devastate tomato and

potato crops at any time during plant ontogeny. It caninfect all above-ground parts of the plant, causing leafand stem necrosis, fruit rot and eventual plant death(http://www.nysipm.cornell.edu/publications/blight/). Phytoph-thora infestans can also infect tomato seed (Rubin et al., 2001;Rubin and Cohen, 2004) and potato tubers (Park et al., 2005d).

Late blight lesions first appear near leaflet margins. The pur-ple, dark brown or black water-soaked lesions often have a paleyellowish-green border that blends into the healthy tissue. Asthe disease progresses, lesions may occur elsewher on the leaves(http://www.cropinfo.net/Potatoblight.htm#Late%20Blight%20Information). Fluffy, white sporangia may grow on the lower(abaxial) leaflet surface in moist weather. As LB progresses,leaflets shrivel and die and the disease spreads to the restof the foliage, leading to extensive defoliation. Dark brownLB stem lesions often first appear at the top of the stem orat a node and may progress down the stem. Firm, brown,

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and greasy tomato fruit lesions are often located at thestem end and sides of green fruit, rendering them unmar-ketable. Infected tomato fruit may be invaded by secondaryorganisms, causing soft-rot disease. LB can affect potatotubers before or after harvest, with affected tubers havingdry rot and brown, or purple, sunken or depressed lesions(http://plantclinic.cornell.edu/Old%20FactSheets/latebl4.htm).

C. The PathogenPhytophthora infestans, as translated from Greek for “plant

destroyer,” is a member of the oomycetes. The oomycetesare a group of fungus-like microorganisms sometimes referredto as the “water molds.” Fungus-like characteristics includ-ing heterotrophy and filamentous growth led to the belief thatoomycetes such as P. infestans were closely related to fungi;however, oomycetes are now considered to be more closely re-lated to photosynthetic microorganisms such as brown algaeand diatoms than the true fungi (de Bary, 1876; Lamour et al.,2007). The oomycete lineage contains a variety of saprophyticand pathogenic microorganisms, with Phytophthora species be-ing the most intensively studied and P. infestans one of the mostprominent pathogens. Phytophthora infestans was implicated asthe cause of the Irish potato famine in the 1840s.It can infectother members of the Solanaceae, including eggplant, night-shade and petunia.

1. Disease CycleThe life cycle of P. infestans includes rapid, asexual repro-

duction conducive to disease development as well as sexual re-production, which can lead to the generation of new pathogenraces. The success of P. infestans as a pathogen originates fromits effective asexual and sexual life cycles. Cool, humid, rainyor foggy conditions favor LB infection. Asexual reproduction,or the disease cycle, begins when sporangia, spore-producingstructures, land on host plant tissue, which must be coveredwith a film of water. Sporangia germinate at temperatures above21◦C (optimally at 25◦C) directly on host tissue in a processtaking between 8 and 48 hours. Below 21◦C, up to eight biflag-ellate zoospores are released from the sporangia, with optimalzoospore formation occurring at 12◦C. Biflagellate zoosporespenetrate through the film of water, lose their flagella and encystuntil they produce germ tubes. This occurs after approximatelytwo hours at an optimum temperature between 12 and 15◦C.Germ tubes differentiate into appressoria that invade the hostthrough the leaf cuticle, or less frequently, the stomata. Germtube differentiation occurs optimally between 21 and 24◦C. In-tercellular hyphae develop and travel inside the host betweencells, using haustoria to form biotrophic feeding relationshipsin the mesophyll. Rapid colonization occurs optimally between22 and 24◦C. Hyphae spread and sporangiophores eventuallyemerge from stomata, soon after LB symptoms are apparent,which is often between 5 and 10 days after inoculation. Sporu-lation occurs to produce 2N sporangia, which eventually release

zoospores to promote aerial transmission of LB and continuethe disease cycle (reviewed by Judelson, 1997). Disease devel-opment ceases if temperatures increase above 35◦C, though P.infestans can survive in living host tissue and the disease canprogress when conditions again become favorable.

The sexual life cycle of P. infestans requires the matingof individuals with different mating types, termed A1 and A2(Gallegly and Galindo, 1958; Brasier, 1992). The mating typesare not dimorphic forms of P. infestans, but are compatibilitytypes differentiated by mating hormones (reviewed by Judel-son, 1997). When mycelia of the two mating types interact,mating hormones induce gametangial formation in the oppos-ing mating types, resulting in sexual propagation by means ofoospore formation. During gametangia formation, vegetative,diploid mycelia undergo meiosis to form haploid antheridia andoogonia. During the sexual life cycle, an antheridium fuses withan oogonium to form a diploid oospore. Unlike sporangia, whichare airborne, fragile, and need live plants for survival, oosporescan survive for extended periods of time in harsh conditionsoutside the living host plant. Oospores can germinate under en-vironmentally favorable conditions and release diploid progenyof A1 or A2 mating type (Judelson, 1997).

2. Clonal LineageA clonal lineage is defined as the asexual descendents of

a genotype (Fry and Goodwin, 1997a). The mating type andgenotypes for several isozyme and restriction fragment lengthpolymorphism (RFLP) markers are often used to describe a P.infestans clonal lineage. Glucose-6-phosphate dehydrogenaseand peptidase are two commonly used isozymes, while RG57 isthe commonly used RFLP marker (Goodwin et al., 1992; Good-win et al., 1994). US-1 was the first clonal lineage (A1 matingtype) described and likely the only one present outside Mexicountil the late 1970s, when additional lineages emerged, someof which possessed the A2 matingtype (Goodwin et al., 1994).US-1 remained the dominant clonal lineage until the late 1980s,but by 1993 it represented less than 8% of the North AmericanP. infestans population (Goodwin et al., 1998). Since the declineof US-1, more than a dozen clonal lineages have been identifiedand described (Goodwin et al., 1998). The most relevant and im-portant lineages include US-6, US-7, US-8, US-11 and US-17(Gavino et al., 2000). US-8 has been particularly aggressive onpotato tubers and foliage. In 1991, US-8 was found in only onecounty in New York, whereas by 1995 it was found in nearlyall potato production areas in the United States and Canada (Fryand Goodwin, 1997a). The significance of US-8 has led to its usein many potato LB resistance screening and breeding programs(Douches et al., 1997; Douches et al., 2001; Kirk et al., 2001;Chen et al., 2003). US-11 is a particularly important clonal lin-eage as it is a highly aggressive, sexually-derived clonal lineagethat has devastated tomato crops in the Pacific Northwest, NewYork and California (Gavino et al., 2000). Similarly, US-17 wasalso very destructive for commercial tomato production in themid- to late 1990s.

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80 M. R. FOOLAD ET AL.

The occurrence of both A1 and A2 mating types outside Mex-ico sparked great interest in describing genetic variability in P.infestans populations throughout the world, including Canadaand the U.S. (Goodwin et al., 1998), France and Switzerland(Knapova and Gisi, 2002), Hungary (Bakonyi et al., 2002),Nepal (Ghimire et al., 2003), Israel (Cohen, 2002), Peru (Perezet al., 2001) and Taiwan (Deahl et al., 2002). Genetic characteri-zation of isolates from various regions as well as the same regionover time has indicated that P. infestans populations are diverseand have changed over time. Genetic variation in populations ofP. infestans has increased in recent years mainly due to the pres-ence of both mating types and occurrence of sexual reproduction(Rubin and Cohen, 2004). Many of the new recombinant isolatesmay be more aggressive (Gavino et al., 2000) or are resistantto the systemic fungicide metalaxyl, rendering chemical controlless effective.

On the basis of their reaction on tomato or potato plants, twophysiological races of P. infestans have been identified, namelyrace T-0 and T-1 (Conover and Walter, 1953; Gallegly, 1960).Race 0, known as the common LB race, was the original raceidentified in tomato and is considered no longer to be a problem.Race 1 is a more recent and much more aggressive race of thepathogen. Most of the current clonal lineages of P. infestans arebelieved to be of race 1.

3. Possible Mechanism of PathogenicityAlthough much is known about P. infestans, including

the genome sequence (http://www.broad.mit.edu/annotation/genome/phytophthora infestans/Home.html), the molecular ba-sis of P. infestans pathogenicity remains somewhat less un-derstood. In general, during infection, phytopathogens secretepeptides and proteins, broadly known as effectors (Lamouret al., 2007). Effectors can be of two types; apoplastic effectors,which target the extracellular space, and cytoplasmic effectors,which target subcellular compartments. Effectors include en-zymes involved in degrading the plant cell wall and suppressingextracellular plant defenses (Abramovitch and Martin, 2004;Vorwerk et al., 2004).

A computational analysis of over 2000 P. infestans expressedsequence tags (ESTs) resulted in the identification of 142 nonre-dundant extracellular proteins. Expression analysis of 63 ofthem led to the identification of two new necrosis-inducingcDNAs, crn1 and crn2 (Torto et al., 2003). It was furtherdetermined that crn2 induces tomato defense response genes.A later In Planta study of P. infestans’ secretome in tomatousing the yeast secretion trap technique identified proteinssecreted by the pathogen and the host during a disease inter-action (Lee et al., 2006). The P. infestans proteins identifiedincluded two elicitins, one of which is a well-characterized se-creted elicitor of the hypersensitive response, an extracellularmetallopeptide, a cutinase and a cysteine-rich, species-specificelicitor-like protein with similarity to PcF, a necrosis-inducingprotein secreted by P. cactorum (Lee et al., 2006). Interest-ingly, the majority of P. infestans’ genes encoding secretory pro-

teins had no identifiable function and were only detected duringpathogenesis.

In addition to the identification of P. infestans elicitors, thereis limited information on the gene-for-gene interaction betweenP. infestans and potato. A candidate gene approach, followedby association genetics, was used to identify Avr3a, an aviru-lence gene in P. infestans (Armstrong et al., 2005). Avr3a cor-responds to the R3a resistance gene in the wild potato speciesSolanum demissum and it was demonstrated that Avr3a encodesa protein recognized in the host cytoplasm that triggers R3a-dependent cell death. Using P. infestans as a model oomycetephytopathogen, Whisson et al. (2007) studied translocation ofeffector proteins into host cells. Using Avr3a, they demonstratedthat the RXLR-EER motif is required for translocation intopotato host cells (Whisson et al., 2007). Interestingly, the RXLR-EER resembles the RXLXE/D/Q motif required for transloca-tion of Plasmodium effectors to host red blood cells. Computa-tional analyses suggest there are approximately 200 candidateRXLR-EER effectors in the P. infestans genome (Lamour et al.,2007).

D. Possible Mechanisms of Host ResistancePlant cell walls function as a first line of defense against

pathogens by creating a physical barrier, which the pathogenmust overcome in order to cause infection. Currently little isknown about the role that various cell wall components mayplay in disease resistance. It has been suggested that polysac-charides have a role in disease resistance, however, there are fewdocumented genetic differences in cell wall composition withina species. And when differences exist, there is no way to examinewhether they play a role in disease interactions, as there are alsoother differences across genotypes (Vorwerk et al., 2004). Plantcells secrete a broad spectrum of compounds to combat disease,including constitutively expressed and inducible defense-relatedproteins. The functions of such proteins include degrading mi-crobial cell walls and inhibiting pathogen-released elicitors (Vor-werk et al., 2004; Thatcher et al., 2005; van Loon et al., 2006;Ferreira et al., 2007).

An In Planta study of secretomes in tomato using the yeast se-cretion trap technique identified proteins secreted by P. infestansand tomato during a compatible interaction (Lee et al., 2006).Almost half the secreted proteins in the tomato plants had knownassociations with defense, including pathogenesis-related (PR)proteins, structural proteins, glycosyl hydrolases (which targetglucans, typical constituents of microbial cell walls) and a puta-tive peroxidase. Other secreted proteins had less defined roles indefense, including those induced by wounding or elicitors. Ad-ditionally, about one third of the induced genes had no obviousfunctional domains or homology to known genes.

To date, no mechanisms for LB resistance have been proposedin either tomato or potato. A preliminary study was conductedto elucidate the mechanism of vertical LB resistance con-ferred by the S. bulbocastanum R-gene, RB, by investigating the

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effects of RarI and SgtI on LB resistance (Bhaskar et al., 2008).Unlike the resistance conferred by most major genes, RB slowsdisease progress but does not eliminate disease symptoms (seebelow). RarI and SgtI are known to regulate R gene expression(Shirasu et al., 1999; Hubert et al., 2003). Although proposedto be involved in forming or stabilizing R-protein associatedrecognition complexes, silencing RARI using RNAi had no ef-fect on RB resistance, indicating that it is not required in the LBresistance response. Conversely, silencing SGT1, which knownto be involved in NB-LRR and Pto-kinase mediated resistance,resulted in disease susceptibility, indicating that SGT1 plays arole in RB resistance.

With respect to horizontal resistance, global gene expressionpatterns in response to P. infestans inoculation have been stud-ied in potato. A recent cDNA microarray study that included1009 ESTs from a subtractive cDNA library identified 348 P.infestans responsive genes (Wang et al., 2005). Of these, ∼27%were associated with metabolism, ∼20% with plant defense,∼13% with signaling and transcriptional regulation and ∼33%had unknown function. Interestingly, genes involved in the hy-persensitive response were reported to be up-regulated followingP. infestans inoculation, even though the potato EST clone usedin the study was derived from an R-gene-free population. Thisstudy also identified three distinct stages of gene expression, in-cluding early [2–6 hours postinoculation, (hpi)], mid (8–24 hpi)and late (36–72 hpi). Subsequently, a second cDNA microarraystudy on 100 selected ESTs from the same subtractive cDNAlibrary was conducted (Tian et al., 2006). This study aimed toidentify the expression patterns of selected genes and to provideinsight into the molecular mechanism of horizontal LB resis-tance in potato. Based on expression levels at 24, 48 and 72 hpi,four groups of genes were identified. The first group consisted ofgenes upregulated at 72 hpi; most of these genes were associatedwith metabolism. The second group of genes was upregulatedat all three time intervals assessed; these included genes with avariety of functions. The third group of genes reached peak lev-els of expression 48 hpi; these genes were defense-related andmetabolic genes. The fourth group of genes was up-regulated24 hpi and decreased thereafter; these genes were transcriptionrelated. The authors concluded that there were multiple defensemechanisms involved in horizontal LB resistance in potato andthat alteration of metabolic pathways was one of the most im-portant disease defense responses.

E. Disease Screening MethodsThe three conditions of the disease triangle, including the

presence of pathogen, host, and environmental conditions con-ducive to disease progress, must be met to screen for LB resis-tance. The pathogen must be in a form that can infect the plant(see inoculum preparation below). The host tissue screened canbe whole plants, detached leaves (leaflets), tomato fruit or potatotubers (disks or slices). Typically, screening for LB resistance isdone with whole plants either under field or greenhouse condi-

tions, with leaflets and tubers under growth chamber conditions.To successfully screen for LB resistance, regardless of the typeof host tissue used, the environment should generally have mod-erate temperature (15–20◦C) and high humidity (Fry and Good-win, 1997b). In the case of tomato fruit, no particular screeningfor LB resistance has been published to date. This is a concernas the fruit is the marketable product of the tomato crop. It hasbeen observed that some tomato S. pimpinellifolium accessionsthat exhibited strong foliar resistance to LB were susceptible atthe fruit level (Irzhansky and Cohen, 2006; M.R. Foolad et al.,unpublished data). Similarly with potato, foliage resistance isnot always associated with tuber resistance (Kirk et al., 2001).

1. Inoculum PreparationTo prepare the pathogen that can infect the plant, P. infestans

is placed into suspension culture under conditions favorable forzoospore release. This can be induced by storing P. infestansat 4◦C for one to a few hours prior to inoculation (Malcolm-son, 1976). Zoospores produce germ tubes to penetrate hosttissue and infect the plant. The method of inoculation dependson the screening method, which may include field, greenhouseand growth chamber screenings.

2. Field ScreeningField evaluation allows for disease screening of whole-plant

on the largest scale possible, compared to greenhouse and growthchamber methods. Field screening is particularly desirable whenscreening must be done on large breeding populations. Whenscreening plants for LB resistance in the field, inoculation canbe either natural or artificial. Typically artificial inoculations aredone with one or a mixture of P. infestans isolates already presentin the area. Exotic isolates are avoided in field screenings so asnot to introduce, or allow for generation of new recombinant iso-lates. Artificial inoculation is typically done using a backpacksprayer, following sprinkler irrigation early in the evening whentemperature is cool and humidity is high. A possible drawbackto a large-scale field inoculation is reduced-uniformity of inoc-ulation across plants and throughout the field. This may lead todisease escape, a confounding factor in field screening. Otherdisadvantages of field screening include possible confoundingeffects by uncontrollable environmental and biological factorssuch as other foliar diseases and pests.

Following a successful inoculation and subsequent appear-ance of disease symptoms, an objective measure of evaluatingdisease response must be determined. For whole-plant evalua-tion in the field or greenhouse, this can include using a scalesuch as Malcolmson’s 1–9 level for assessing LB resistance(Malcolmson, 1976). Using this scale, a score of 1 indicatesthat greater than 90% of the tissue is necrotic, a score of 8 indi-cates that 10% of the tissue is necrotic and a score of 9 indicatesthat a plant is completely healthy. The composite form of Mal-colmson’s scale measures the number and size of LB lesions,in addition to providing an overall assessment of the disease.Cruickshank (1982) published drawings that depict each value

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82 M. R. FOOLAD ET AL.

on Malcolmson’s scale to aid in scoring (Cruickshank et al.,1982). Other possible measures of whole-plant disease responseinclude increasing the span of the scale to begin with a score of0 for a completely healthy plant to a score of 100 for a plantcompletely blighted and killed by the disease. Using this sys-tem, the area under disease progress curve (AUDPC) can bedetermined by scoring the disease progress over time, and in-tegrating host, pathogen and environmental effects on the dis-ease (Shaner and Finney, 1977; Johnson and Wilcoxson, 1982;Campbell and Madden, 1990). An AUDPC can also be adaptedfor use in growth chamber experiments with detached leafletsby measuring the rate of LB lesion expansion over time (Goth,1997). In addition to measuring lesion size and determining therate of lesion expansion in detached leaflets, infection efficiency(percentage of successful infections), latency period, and sporedensity can be used as measures of LB resistance (Birhman andSingh, 1995).

3. Greenhouse ScreeningIn this screening for LB resistance, whole plants must be

grown and inoculated in confined greenhouses to prevent intro-duction of new isolates and spread of the disease. Prior to inocu-lation, plants are misted under cool and humid conditions. Plantsare usually inoculated by atomizing them with a water suspen-sion of P. infestans (normally with a density of ∼10,000 spo-rangia/ml). Following inoculation, greenhouse conditions aremaintained with high humidity and 12 hours alternating light(∼21◦C) and dark (∼16◦C) to encourage pathogen infectionand growth. Disease ratings are usually made 7 to 10 days afterinoculation. Under confined greenhouse conditions, P. infestansisolates not present in the field may be examined without thefear of introducing new isolates to the environment. However,precautions must be taken during and after the experiment notto spread the pathogen, including restricted and monitored ac-cess to the greenhouse, autoclaving infected plants and grow-ing media, and sterilizing the greenhouse after the experiment.Greenhouse screening for LB allows control of environmentalfactors that influence the virulence of the pathogen and the re-action of the host while excluding other pathogens, which mayoccur in the field. Greenhouse evaluation is also independentof such factors as growth habit and fruit load, which may af-fect the plant’s reaction to the pathogen in the field. In spiteof these advantages, many research programs may not affordthe expenses associated with the required facilities to conductsuccessful greenhouse screening.

4. Growth Chamber Screening Using Detached LeafletsDetached-leaflet assay has been used for disease screening

with obligate parasites and to a lesser extent with facultative par-asites (Yarwood, 1946; Locke, 1948; Goth, 1997). It has beenused extensively to evaluate LB resistance in tomato and potatosince 1954, when it was used to test for the presence of R genes(Toxopeus, 1954). In this method, fully-expanded, but not oldleaflets (normally from the third or fourth leaf from the vine

apex) are detached and used for inoculation. Sampled leafletsshould be of similar physiological age across genotypes. Devi-ation from this requirement may make the screening unreliable.Inoculation can be done by dropping inoculum on the abax-ial (lower) leaflet surface using a micropipettor, or by dippingthe leaflet in a suspension of P. infestans. Inoculated leaflets areplaced in Petri dishes with water agar on one surface to maintainhumidity within the dishes. Petri dishes are normally placed inincubators maintained at 18–21◦C and 12 hrs light for 5 to 7 daysbefore scoring. The detached-leaflet method of disease screen-ing does not have the extensive space or facility requirementsof field or greenhouse screening. Furthermore, this method pro-vides for uniform inoculation of experimental objects with nodisease escape. However, there are mixed reports on the relia-bility of this screening technique and its correlation with fieldor greenhouse disease data (Malcolmson, 1969; Goth, 1997;Vleeshouwers et al., 1999). Our extensive experience with thedetached-leaflet screening method indicates that while this tech-nique provides a reasonable assessment of LB resistance, it isnot a fully reliable screening system (M.R. Foolad et al., un-published data). This is in agreement with assessments by manyother research groups.

F. Nongenetic Measures of Disease ControlTomato and potato LB costs growers an estimated $5 bil-

lion annually worldwide, including the cost of disease controland crop losses (Judelson and Blanco, 2005). Estimated cost offungicides and crop losses due to LB in the U.S. exceeds $210million annually (Guenthner et al., 2001). In addition to geneticresistance (discussed below), LB can be controlled by culturalpractices and chemical applications. Prior to the re-emergenceof LB in the late 1970s, cultural practices in combination withfungicide applications effectively controlled the disease. How-ever, these approaches alone are not expected to provide sus-tainable control of the disease in the future, as discussed below.

1. Cultural PracticesCultural practices are important components of growers’

strategy in disease management, and they can impact diseasedevelopment and control. The aims of cultural control of LB areto minimize inoculum buildup, prevent introduction of inocu-lum from nearby potato cull piles or from tomato transplants,minimize infection rate and generate conditions unfavorable fordisease development and spread. Specific cultural practices usu-ally employed to control LB include crop rotation and fallow,eliminating volunteer tomato and potato plants, planting nonin-fected seedlings and tubers, and eliminating sources of LB suchas potato cull piles. The latter is of particular importance becausecull piles can serve as a living host on which P. infestans myceliacan survive over the winter and produce tremendous amounts ofairborne spores at the beginning of the new field season. If thisoccurs, the next year crop is at risk of LB devastation.

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2. Chemical ApplicationsChemical control measures can be effective in managing LB,

especially when guided by disease forecast systems. Chemi-cal controls have included the use of both protectant and sys-temic fungicides. Protectant fungicides are usually applied be-fore emergence of the disease whereas systemic fungicides areapplied before or upon disease development. Current chemicalpractices to control LB include a mixture of fungicides designedto slow the disease progress (Gisi and Cohen, 1996). Metalaxylfungicides, a class of systemic fungicides, have been widelyemployed to control LB. Metalaxyl fungicides inhibit riboso-mal RNA (rRNA) polymerases in fungi by reducing incorpo-ration of uridine. However, unfortunately metalaxyl resistancein P. infestans populations appeared as early as 1980 (Gisi andCohen, 1996). Fungicide resistance is controlled by a single, in-completely dominant gene present at low levels in P. infestansnatural populations. In addition to the development of resis-tance, fungicides are expensive, harmful to the environment andhumans, and must be applied at the proper time. With the useof at least partially resistant cultivars, the number of fungicideapplications and/or the rate of application can be significantlyreduced (Shtienberg et al., 1994; Kirk et al., 2001; Stevensonet al., 2007), particularly when combined with blight forecast-ing (Grunwald et al., 2002). The more resistant the cultivar, thegreater the potential for fungicide reduction (Naerstad et al.,2007). It should be noted that copper fungicides have also beenused in organic fresh-market tomato production, but they havebeen shown to only suppress the LB disease.

G. Genetic Measures of Disease ControlResistance to P. infestans can be classified into two

categories: race-specific (or pathotype-specific) and race-nonspecific resistance, with races defined by disease interac-tions between the pathogen and different host genotypes. Race-specific resistance, also known as vertical resistance, single-generesistance or qualitative resistance, is typically controlled by asingle gene. This type of resistance was first described by Flor(1955) to explain interactions between flax and flax rust, when heobserved that the disease response depended on factors presentin both the host and the pathogen (Flor, 1955). This type of inter-action is also known as “gene-for-gene interaction” between hostplant and disease organism. The host resistance gene product,also known as the R gene product, interacts with the pathogen’spathogenicity gene product, also known as the Avr gene product.A classic example of gene-for-gene interaction in tomato is theresistance to bacterial speck, caused by Pseudomonas syringaepv.tomato, where the tomato Pto gene interacts with the bacterialspeck AvrPto gene (Martin et al., 1993). Single gene resistancetypically confers complete resistance to one or a limited num-ber of races of a pathogen. However, other races/isolates of thepathogen may completely overcome the host resistance. Single-gene resistance is often not durable and can be quickly brokendown by emergence of new races of the pathogen. Conversely,

race-nonspecific resistance, also known as horizontal resistance,polygenic resistance, quantitative resistance or field resistance,is usually governed by more than one gene and may be moredurable. Horizontal resistance usually confers partial resistanceto multiple isolates/races of the pathogen. This type of resis-tance slows, but does not stop disease progress. The polygenicnature of horizontal resistance makes it more difficult to breedfor, compared to vertical resistance, however, breeders find hori-zontal resistance more desirable because it is often more durable.

LB resistance in the form of vertical as well as horizontalresistance has been reported in wild species of both potato andtomato. As potato and tomato are closely related and have sim-ilar genomes, and because the LB resistance genes and QTLsidentified in potato may have relevance in tomato or correspondto tomato LB resistance genes and QTLs, it is useful to reviewand discuss LB genes and QTLs in potato as well.

1. Late Blight Resistance in PotatoMajor LB resistance genes have been reported in several

wild species of potato, in particular the Mexican hexaploidspecies S. demissum Lindl. (nightshade) and the Mexican self-incompatible diploid species S. bulbocastanum Dunal (orna-mental nightshade). At least 11 resistance (R) genes have beenidentified in S. demissum, several of which have been mappedand/or cloned (Naess et al., 2000; Gebhardt and Valkonen, 2001;Bradshaw et al., 2006a) or incorporated into potato cultivars(Malcolmson and Black, 1966; Gebhardt and Valkonen, 2001;Bradshaw et al., 2006a). All R genes of S. demissum exhibit arace-specific, hypersensitive resistance response (Niederhauserand Mills, 1953; Song et al., 2003). However, P. infestans iso-lates have been identified that overcome all of the reported Rgenes (Niederhauser and Mills, 1953; Bradshaw et al., 2006a).Conversely, three LB resistance genes have been reported in S.bulbocastanum, which do not seem to be race-specific (discussedbelow). A few R genes have also been reported in other wildspecies of potato, including S. berthaultii Hawkes (Ewing et al.,2000), S. mochiquense Ochoa, S. phureja Juz. & Bukasov., andS. pinnatisectum Dunal (Kuhl et al., 2001; Smilde et al., 2005;Sliwka et al., 2006).

Seven of the eleven R genes identified in S. demissum havebeen mapped. R1 has been mapped to chromosome 5, associ-ated with potato genomic clones GP21 and GP179 and closeto a dominant potato virus X resistance gene, Rx2 (Leonards-Schippers et al., 1992; El-Kharbotly et al., 1994). R1 confersfoliar as well as tuber LB resistance (Park et al., 2005d). R2 hasbeen mapped to chromosome 4, tightly associated with threeAFLP markers (Li et al., 1998).R3 has been mapped to distalend of chromosome 11, associated with tomato genomic cloneTG105a and potato genomic clones GP185 and GP250a (El-Kharbotly et al., 1994). Subsequently, GP185 and GP250a werealso linked to R6 and R7, indicating that these three R geneswere in a similar position (El-Kharbotly et al., 1996). It has beenproposed that R3, R6 and R7 are either alleles of the same lo-cus or three tightly linked genes. Subsequently, R10 and R11

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were also mapped to the location of R3 and were presumed tobe alleles of R3, though R11 acted more like a QTL explaining∼57% of the variation in LB response (Bradshaw et al., 2006a).

To date, two of the eleven R genes from S. demissum, R1and R3a, have been cloned. R1was the first LB resistance genecloned, using a combination of the positional cloning techniqueand the candidate gene approach (Ballvora et al., 2002). R1 hasa coiled-coil (CC) domain, a putative nucleotide binding (NBS)domain and a leucine rich repeat (LRR) domain. Using fine map-ping and accurate disease screening, it was determined that R3consisted of two tightly linked genes, R3a and R3b, which werelocated 0.4 cM apart on the distal end of chromosome 11 (Huanget al., 2004). R3a confers foliage resistance whereas R3b pre-sumably confers foliar and tuber resistance (Park et al., 2005d).Employing a comparative genomics approach using tomato asa model Solanaceous plant, R3a was cloned and determined tohave CC, NBS and LRR motifs (Huang et al., 2005).

An unnamed R gene was reported in S. berthaultii, whichwas mapped to potato chromosome 10, 4.8 cM from TG63 andin a region similar to that of the tomato LB resistance gene Ph-2(Ewing et al., 2000). In the diploid species S. mochiquense, anLB resistance gene (Rpi-moc1) was mapped to the distal partof the long arm of chromosome 9, in a similar region as thetomato LB resistance gene Ph-3 (Smilde et al., 2005). In S.phureja, a major locus, Rpi-phu1, conferring leaf and tuber re-sistance to LB was mapped to chromosome 9, 6.4 cM from thepotato genomic clone GP94 (Sliwka et al., 2006). Similarly, inS. pinnatisectum an LB resistance gene, Rpi1, was mapped tochromosome 7, 5.2 cM from the tomato genomic clone TG20A,9.4 cM from the potato cDNA clone CP56 and within 10 cMof resistance loci I1 (conferring resistance to fusarium wilt)and Gro1 (conferring resistance to potato cyst nematode) (Kuhlet al., 2001). It should be noted that Rpi1 may correspond tothe resistance gene R9 from S. demissum as the P. infestans iso-late used in the LB evaluations possessed avr9, the avirulencelocus that corresponds to R9 (Kuhl et al., 2001). To initiatemap-based cloning of Rpi1, two S. pinnatisectum BAC librarieswere constructed and screened with TG20A and CP56 (Chenet al., 2004). This led to the identification of two and four BACs,respectively.

Unlike the race-specific resistance observed in S. demissum,S. berthaultii, S. mochiquense, S. phureja, and S. pinnatisectum,LB resistance in diploid species S. bulbocastanum generallyslows disease progress against a broad spectrum of P. infestansisolates (Naess et al., 2000; Song et al., 2003; van der Vossenet al., 2005). To date, three to possibly four LB resistance locihave been identified in this species. The first gene, RB, wasmapped to potato chromosome 8 and subsequently cloned usinga map-based cloning approach (Naess et al., 2000; Song et al.,2003). Simultaneously, using the same source of resistance, theRB gene was mapped and cloned by another group who named itRpi-blb1(van der Vossen et al., 2003). Subsequently, to assist intransferring RB to cultivated potato, a linked PCR-based markerwas developed (Colton et al., 2006). However, S. bulbocastanum

cannot be directly crossed with S. tuberosum, making breedingfor RB (Rpi-blb1) tedious and time-consuming. Recently, RBhomologues were discovered in S. stoloniferum, a species thatcan be crossed with S. tuberosum, though with low efficiency,providing the possibility that the resistance may be more easilytransferred to the cultivated potato (Wang et al., 2008).

A second source of LB resistance in S. bulbocastanum led tothe identification and mapping of a new resistance gene (Rpi-blb2) on potato chromosome 6 (van der Vossen et al., 2005).Interestingly, this gene was found in the same chromosomal re-gion as the tomato resistance genes Mi-1 (conferring resistanceto root-nematode, potato aphid and sweet potato whitefly), Ol-4(conferring resistance to tomato powdery mildew), Ol-6, Cf-2(conferring resistance to tomato leaf mould) and Cf-5. Rpi-blb2was also fine-mapped and cloned using a map-based cloningapproach, and it was determined that it showed 82% sequenceidentity at the amino acid level with the Mi-1 gene (van derVossen et al., 2005). A third source of LB resistance in S. bulbo-castanum led to the identification and fine mapping of Rpi-blb3to a 0.93 cM region of potato chromosome 4 (Park et al., 2005a).Using a comparative genomics approach, it was determined thatRpi-blb3 lies in an LB resistance gene cluster that also containsR2, R2-like and Rpi-abpt. R2-like does not come from the samegenetic background as R2, but the two are phenotypically indis-tinguishable. However, R2-like has been fine mapped to a 0.4 cMregion of chromosome 4 (Park et al., 2005b). The Rpi-abpt gene,which confers foliar LB resistance, was originally identified ina potato clone known as ABPT, which is a quadruple hybrid in-cluding genetic contributions from S. acaule, S. bulbocastanum,S. phureja and S. tuberosum (Hermsen and Ramanna, 1973).However, based on the analysis of the phenotypic response ofthe four parents to P. infestans and the molecular marker datafrom the four parents, it is thought that Rpi-abpt was derivedfrom S. bulbocastanum (Park et al., 2005c). Rpi-abpt has beenmapped to a 0.5 cM region of potato chromosome 4.

In addition to the major LB resistance genes in potato, a num-ber of studies have identified QTLs conferring field/quantitativeresistance in crosses within and between potato species. In moststudies, associations were observed between quantitative LB re-sistance and late maturity (Oberhagemann et al., 1999; Ewinget al., 2000; Visker et al., 2005; Simko et al., 2006) with few ex-ceptions (Visker et al., 2004). The first QTL analysis for potatoLB resistance was conducted by Leonards-Schippers et al.(1994), whom identified 11 resistance QTLs on 9 potato chromo-somes in crosses within S. tuberosum (Leonards-Schippers et al.,1994). Interestingly, some of these QTLs were race-specific orwere located near the race-specific R genes, a phenomenon alsoreported by other researchers (Oberhagemann et al., 1999). Ithas been hypothesized that R genes demonstrating a hypersen-sitive response may represent an extreme form of an allele forquantitative resistance (Gebhardt, 1994).

Several other studies have also identified and analyzed QTLsfor LB resistance in crosses within S. tuberosum. Collins et al.(1999) examined QTLs for foliar and tuber LB resistance and

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their relationships with plant maturity (Collins et al., 1999). Thisis believed to be the first study that examined the association be-tween quantitative LB resistance and maturity. Significant QTLswere identified on chromosomes 3, 5, 6, and 9 for foliar resis-tance, on chromosomes 2, 5, 9, and 12 for tuber resistance, andon chromosomes 5, 6, 7 and 8 for plant maturity. Interestingly,the LB QTL on chromosome 5, which exhibited the strongesteffect on foliar LB resistance, also had the greatest associationwith lateness in maturity (Collins et al., 1999). A more recentstudy further investigated QTLs for foliar and tuber LB resis-tance as well as QTLs affecting plant maturity in potato (Sliwkaet al., 2007). While 3 major QTLs were identified for LB resis-tance on chromosomes 3 (based on whole tuber tests), 4 (basedon tuber slice tests) and 10 (based on foliar tests), a QTL wasidentified on chromosome 5 for plant maturity, presumably thesame QTL as identified in the Collins et al. (1999) study. Thisstudy, however, also demonstrated the presence of major QTLsfor LB resistance independent of factors affecting plant maturity(Sliwka et al., 2007). This finding provides a positive outlookfor the identified QTLs from a breeding perspective, suggest-ing the possibility of developing potato cultivars with field LBresistance and early to mid-season maturity.

QTLs for LB resistance have also been studied in a cross be-tween S. phureja and di-haploid S. tuberosum (Ghislain et al.,2001; Bradshaw et al., 2006b). S. phureja was selected for sev-eral reasons, including the fact that it is a cultivated potato withdesirable agronomic and culinary quality and that it can easilybe used as a parent to make LB-resistant tetraploid potatoes. Inthe study conducted by Ghislain et al., the strongest QTL wasidentified on chromosome 12 of S. phureja, explaining ∼43% ofthe phenotypic variance (PVE), in addition to a smaller QTL onchromosome 7 (Ghislain et al., 2001). QTLs for LB resistancewere also detected on S. tuberosum chromosomes 3, 5 and 8. Asa follow-up to this study, markers that corresponded to defenserelated genes were developed to determine potential associationsbetween defense-related genes and the QTLs for LB resistance(Trognitz et al., 2002). This study demonstrated that markersdeveloped from genes related to the phenylpropanoid pathway,pathogenesis (PR), cytochrome p450 and WRKY transcriptionfactors were associated with LB resistance QTLs on chromo-somes 3 and 12. In the study conducted by Bradshaw et al., alarge QTL was identified accounting for 78% and 51% of thevariation in foliage resistance in the greenhouse and field exper-iments, respectively, and 27% of the variation in tuber resistance(Bradshaw et al., 2006b). This QTL, located on chromosome 4,is in the vicinity of Rpi-blb3, Rpi-abpt, R2 and R2-like. A BAClibrary of the resistant parent has since been constructed andscreened with potentially linked resistance gene analogues as astep toward cloning the LB resistance gene(s) responsible forthe QTL (Hein et al., 2008).

QTLs for LB resistance have also been identified in potatowild species in crosses with the cultivated potato. In S.berthaultii, for example, five QTLs conferring foliar resistancewere identified on chromosomes 1, 3, 7, 8 and 11 (Ewing et al.,

2000). QTLs on chromosomes 1, 3, 8 and 11 were in closeproximity to previously detected QTLs for plant maturity, againsuggesting association of field LB resistance with late matu-rity. In S. microdontum, 3 major QTLs for foliar LB resistancewere identified on chromosomes 4, 5 and 10, of which thoseon chromosomes 4 and 10 were consistent across populations(Sandbrink et al., 2000). In a later study, a major QTL (PVE >

60%) for LB resistance, and not associated with late maturity,was identified in S. microdontum (Bisognin et al., 2005). ThisQTL, however, remained unmapped as it did not show linkageassociation with any genetic markers with known position onpotato map. In S. paucissectum, major QTLs for foliar LB resis-tance were detected on chromosomes 10, 11 and 12, of whichonly the QTL on chromosome 11 was detected across all ex-periments (Villamon et al., 2005). This QTL was located on thesame chromosome 11 arm as an LB QTL in S. tuberosum, butthe opposite arm as the major LB resistance genes R3, R6 andR7. Also, a few QTLs for foliar LB resistance were detected onchromosomes 6, 8 and 9 in S. vernei, (Sorensen et al., 2006).

QTLs for LB resistance have also been identified in potatowild species in crosses with S. phureja. For example, in a diploidpopulation derived from a cross between S. stenotomum and S.phureja, 3 major QTLs for foliar LB resistance were reportedon chromosomes 3, 5 and 11, explaining 23, 17 and 10% of thetotal phenotypic variation, respectively (Costanzo et al., 2005).The QTLs on chromosome 3 and 11 corresponded to LB QTLsreported in other populations, while the QTL on chromosome 5was new. In a later study, it was reported that foliar resistance inthis population was associated with late maturity (Simko et al.,2006). However, in the same population, 4 new QTLs were alsodetected for tuber LB resistance, located on chromosomes 2, 6,8 and 10. Of these, the QTL on chromosome 10 accounted formore than 60% of the total phenotypic variance for tuber LBresponse (Simko et al., 2006).

Similar to many other complex disease resistance traits, QTLsidentified for LB resistance in potato often vary with the speciesand pathogen isolates used, methods of disease evaluation, map-ping analyses and environmental conditions of the experiments.However, in potato one major QTL on chromosome 5 in thevicinity of R1 has been reported across such variables in a num-ber of studies (Simko, 2002; and references therein) suggestingits potential significance. However, this QTL has also been re-ported to be associated with lateness in maturity (Visker et al.,2005). It is not currently known whether at this location there isa single gene with pleiotropic effects on both LB resistance andplant maturity or if there are tightly linked independent genesaffecting the two traits. Resolving this issue would determinethe practical significance and utility of this QTL.

In summary, considerable efforts have been made to identifygenetic sources of resistance, map and clone resistance genesor QTLs, and transfer resistance to modern potato cultivars. Al-though frequent transfer of race-specific major resistance genesto potato cultivars has significantly contributed to a better con-trol of LB and a considerable reduction in fungicide use, such

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resistance is often not durable and not effective across environ-ments. Pyramiding of major resistance genes has been provento improve the utility of this kind of resistance. Simultaneously,efforts in identifying QTLs conferring field resistance have beenrewarding and have led to identification of numerous contribut-ing genomic locations. A noticeable drawback, however, hasbeen an undesirable association of field resistance with plantphysiological maturity. Despite this frequent negative associa-tion, a few LB resistance QTLs have been identified which seemto be independent of plant maturity. Such QTLs may be useful indeveloping early to mid-season potato cultivars with sufficientLB resistance. However, the recent identification and character-ization of major LB resistance genes in S. bulbocastanum thatsignificantly reduce disease progression when potato plants areexposed to different isolates of P. infestans is very promising. Itis expected that the knowledge of different LB resistance genesand QTLs in potato will translate into development of new cul-tivars with improved and durable resistance and a reduction inthe use of fungicides for disease control.

2. Late Blight Resistance in TomatoFollowing an LB outbreak in the U.S. in 1946, which af-

fected potatoes and tomatoes, a substantial amount of researchwas initiated to locate sources of genetic resistance in tomato.Such research led to the discovery of resistant accessions withinwild tomato species, in particular S. pimpinellifolium (Galleglyand Marvel, 1955). Subsequent research resulted in the identi-fication and mapping of a few race-specific major resistancegenes in this species, as discussed below. In addition, in atleast one S. pimpinellifolium accession, L3707 (PI365951), race-nonspecific LB resistance has been reported (Irzhansky and Co-hen, 2006), though to date no resistance gene or QTL has beenidentified or mapped in this accession. Inheritance studies indi-cated that race-nonspecific LB resistance in this accession wascontrolled by two genes which interacted in an epistatic man-ner. Further studies of this accession indicated that despite fo-liage LB resistance, L3707 segregated for fruit blight resistance(Irzhansky and Cohen, 2006).

The first reported tomato LB resistance gene, Ph-1, a com-pletely dominant gene conferring resistance to P. infestans raceT-0, was originally located in S. pimpinellifolium accessionsknown as West Virginia 19 and 731 (Bonde and Murphy, 1952;Gallegly and Marvel, 1955; Peirce, 1971). In 1962, an LB resis-tant cultivar, Rockingham, which contained Ph-1, was released(Rich et al., 1962). This cultivar was subsequently used to mapPh-1 to the distal end of chromosome 7 using morphologicalmarkers (Peirce, 1971). Furthermore, Ph-1 was incorporated inthe old processing tomato cv. ‘Nova’ and the old fresh markettomato cv. ‘New Yorker’. However, as T-0 is no longer the pre-dominant race and as Ph-1 has been overcome by new races ofP. infestans, this resistance gene is no longer considered a usefulsource of LB resistance in tomato.

A second tomato LB resistance gene, Ph-2, was subsequentlyidentified in an S. pimpinellifolium accession known as West

Virginia 700 (Gallegly and Marvel, 1955). This incompletely-dominant resistance is also narrowly focused and provides onlya reduction in the rate of disease development rather than block-ing the disease. Furthermore, Ph-2 often fails in the presenceof more aggressive P. infestans isolates (Goodwin et al., 1995;Black et al., 1996; M.R. Foolad, pers. obser.). Characterizationof this resistance has been hampered because its expression ispartially dependent upon environmental conditions, plant phys-iological age, the organ assessed and the pathogen isolate used(Moreau et al., 1998). The resistance conferred by West Virginia700 was originally reported to slow down LB progress throughthe action of multiple genes. However, in 1998, the resistancewas mapped to a single location on chromosome 10, betweenmarkers CP105 and TG233, and was named Ph-2 (Moreau et al.,1998). Using RFLP markers, Ph-2 was mapped within an 8.4cM interval on the long arm of chromosome 10. There has notbeen any further effort to fine map or clone Ph-2 (N. Grim-sley, CNRS-INRA, pers. commun.). However, Ph-2 has beenincorporated into a number of named fresh-market and process-ing varieties, including Legend, Centennial, Macline, Pieraline,Herline, Fline, Flora Dade, Heinz 1706, Campbell 28 and Eu-ropeel (Gallegly, 1960; Laterrot, 1994).

Observations in Taiwan, Nepal, Indonesia and the Philippinesthat new P. infestans isolates overcame Ph-1 and Ph-2 promptedfurther screening of S. pimpinellifolium accessions in searchof new sources of LB resistance. This led to the discovery of astrong LB resistance gene, Ph-3, in S. pimpinellifolium accessionL3708 (a.k.a. LA1269 and PI365957) by L. Black at the AsianVegetable Research and Development Center in Taiwan (AV-DRC, 1994). Ph-3, a partially dominant gene, confers resistanceto a wide range of P. infestans isolates that overcome Ph-1 andPh-2. A bulked segregant analysis (BSA) using AFLP markerswas employed to identify markers associated with Ph-3 (Chun-wongse et al., 2002). Furthermore, using the AFLP markers andthe tomato introgression lines derived from a cross between S.lycopersicum cv. M82 and S.pennellii Correl [formerly L. pen-nellii (Correl)] accession LA716 (Eshed and Zamir, 1995), Ph-3was mapped to the long arm of chromosome 9, close to RFLPmarker TG591A (Chunwongse et al., 2002), and a co-dominantPCR marker has been developed for it (MA Mutschler, pers.commun.). Recently, several public tomato breeding programsin the U.S., including North Carolina State University (RG Gard-ner, pers. commun.), Cornell University (MA Mutschler, pers.commun.; Kim and Mutschler, 2005) and the Pennsylvania StateUniversity (M.R. Foolad, unpub. data), have developed fresh-market and/ or processing tomato lines possessing Ph-3. Kimand Mutschler (2003) developed processing tomato inbred lineswith Ph-3 resistance from L3708 and determined that LB re-sistance in L3708 was controlled by more than just Ph-3 locusand suggested that breeding lines developed at AVDRC werelacking at least one of those additional hypostatic genes (Kimand Mutschler, 2005; Lee et al., 2006). The authors further con-cluded that inbred lines or hybrids which contain only Ph-3gene in either homozygous or hetertozygous conditions would

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not be commercially highly desirable as they would not exhibitstrong resistance against aggressive isolates such as US-7 andUS-17. The presence of the yet undetermined additional hy-postatic gene(s) in homozygous or heterozygous condition isnecessary to provide full resistance (Kim and Mutschler, 2005).However, despite the superiority of the Ph-3 complex, in com-parison with Ph-1 and Ph-2, there have been P. infestans isolateswhich have overcome this resistance (Chunwongse et al., 2002;Scott and Gardener 2007; RG Gardner, pers. commun.). This andthe occurrence of newer, more aggressive recombinant isolatesof P. infestans, necessitated a search for identifying additionalsources of LB resistance in tomato.

A comprehensive effort was recently initiated at thePennsylvania State University with the goals of identifying newsources of LB resistance as well as identifying, mapping andpyramiding LB resistance genes in commercial lines and culti-vars of tomato. Screening of ∼300 accessions of S. pimpinel-lifolium accessions resulted in the identification of a few newsources of LB resistance (Foolad et al., 2006). One accession,PSLP153, which exhibited strong resistance to at least 7 isolatesof P. infestans, was chosen for the identification and mappingof resistance gene(s). A selective genotyping approach, employ-ing F2 and F3 populations of a cross between PSLP153 and asusceptible tomato breeding line and using various molecularmarkers including RFLPs, SSRs, ESTs, CAPS and AFLPs, ledto the identification and mapping of a new resistance gene, ten-tatively named Ph-5, on the long arm of tomato chromosome1 (M.R. Foolad et al., unpublished data). Efforts are currentlyunderway to further delineate the region of chromosome 1 thatcontains Ph-5 by adding more markers to this region. Ph-5 willbe fine-mapped by developing NILs in order to clone it using apositional cloning approach. However, Ph-5 is being transferredto fresh-market and processing tomato lines by a combination ofMAS and traditional breeding protocols (M.R. Foolad et al., un-published data). Furthermore, efforts are underway at Penn Stateto pyramid Ph-2, Ph-3 and Ph-5 in new tomato breeding lines.

Thus far no major LB resistance gene has been identified inother wild species of tomato. However, several QTLs conferringrace-nonspecific resistance have been reported in tomato wildspecies S. habrochaites (Robert et al., 2001; Brouwer et al.,2004), S. pennellii and S. pimpinellifolium (Frary et al., 1998).In S. habrochaites accession LA2099, QTLs conferring LB re-sistance were identified on all 12 tomato chromosomes (Brouweret al., 2004). The locations of these QTLs were compared withQTLs for LB resistance in potato using common RFLP markers.It was determined that the QTLs on chromosomes 3 and 4 cor-responded to LB resistance QTLs in potato. Subsequently, threeNILs were developed to fine-map three of the QTLs and to in-troduce them into tomato breeding lines (Brouwer and St.Clair,2004). Each NIL contained one resistance QTL on an intro-gressed interval of 6.9, 8.8 or 15.1 cM. However, severe linkagedrag problems prevented the NILs being useful for breedingpurposes (Brouwer and St.Clair, 2004). Further inspection ofthe NILs determined that they also contained genes/QTLs for

other characteristics such as plant shape, canopy density, ma-turity, fruit yield or fruit size in the same introgressed regions.Interestingly, similar to reports in potato, QTLs for tomato LB re-sistance were also associated with plant maturity and/or growthhabit (Brouwer et al., 2004).

Most recently, a QTL contributing to LB resistance was iden-tified on chromosome 6 of S. pennellii accession LA716 using anF2 population of a cross between this accession and a cultivatedtomato, and the QTL was further confirmed in S. lycopersicum ×S. pennellii introgression lines (Smart et al., 2007). It should benoted that in this study resistance was defined not as the absenceof the disease, but as a relative measure of the disease comparedto highly susceptible plants. However, the value of moderate orhorizontal resistance to LB is questionable due to the short lifecycle of the disease, heavy production of airborne spores whichcan travel long distances and quickly spread the disease, and thevery aggressive nature of the disease.

In summary, much less work has been conducted on genet-ics/genomics of LB resistance in tomato compared to potato.This has been in part due to a greater importance of LB in potatothan in tomato, at least prior to 1990s when many of the potatoisolates were not pathogenic to tomato. However, because ofthe recent increased occurrence of LB in tomato worldwide,greater efforts are being devoted to the identification, charac-terization and introgression of LB resistance genes in tomato.New breeding lines and cultivars containing Ph2 and Ph3 arebeing developed and additional efforts are underway to combinethese genes with the newly identified LB resistance gene, Ph-5.It is expected that more LB resistance genes will be identifiedin tomato, similar to that in potato, and that many of the futurereleases will have different combinations of genes and/or QTLsfor LB resistance.

III. EARLY BLIGHT

A. BackgroundEarly blight (EB), caused by the fungus Alternaria solani

Jones and Grout, is one of the most common and destructive dis-eases of the cultivated tomato in areas with heavy dew, frequentrainfall, and high relative humidity (Barksdale and Stoner, 1977;Nash and Gardner, 1988a). It can also be important in semi-aridareas when nightly dew is frequent (Rotem and Reichert, 1964).Tomato EB is distributed worldwide, however, high epidemicincidence occurs in many regions of North, Central and SouthAmerica, South Asia and Africa. EB is the most important foliardisease of tomatoes and potatoes in the U.S., damaging cropsfrom New England and the Mid Atlantic to the southeast, centraland midwestern states (Jones, 1991). The necrotrophic lifestyleof the pathogen can lead to complete defoliation of tomato plants(Lawrence et al., 2000) and result in yield reductions of morethan 79% in countries such as Canada, India, Nigeria, and theU.S. (Basu, 1974; Datar and Mayee, 1985; Sherf and MacNab,1986; Chaerani and Voorrips, 2006). EB is also an importantdisease of other members of Solanaceae, including potatoes,

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eggplant, horse nettle, and black nightshade. These host speciesmay serve as a reservoir for the fungus and can be a source ofinoculum for tomato crops.

Alternaria solani was believed to be the causal agent of EBin both tomato and potato. However, due to substantial variabil-ity of the fungus (see below), the distinction between causalagents of EB in tomato and potato has been controversial. In astudy of pathogenic specialization on tomato and potato, Sim-mons (2000) reported two culturally and morphologically dis-tinct species of the pathogen, A. tomatophila E.G. Simmonsand A. solani, as causal agents of EB in tomato and potato, re-spectively (Zitter and Drennan, 2005). Conversely, accordingto an AFLP analysis of 112 isolates collected from around theworld, no distinction was made between the isolates that attackedtomato and potato (Perez Martınez et al., 2004). Therefore, thelatter authors suggested that the name A. solani be applied to thecausal agent of EB in both potato and tomato until the detailedmicroscopic and cultural examinations proposed by Simmons(2000) were performed. In the present article, to be consistentwith much of the literature, we identify A. solani as the causalagent of EB in both tomato and potato.

B. Disease SymptomsEB affects all above ground parts of the tomato plant, and

three distinct phases of the disease can be distinguished depend-ing on the symptoms: collar rot in seedlings (stem lesions orcankers), leaf blight (commonly referred to as early blight) andfruit rot (sunken lesions at the stem end of the fruit) (Barksdale,1969; Sherf and MacNab, 1986; Foolad et al., 2002b). Collarrot, which is considered a seed-bed disease that is transportedto the field on tomato transplants, is characterized by forma-tion of dark canker-like lesions at or near ground level on thestem. Eventually, the lesions girdle the stem, forming “collars,”and damage the vascular system (Horsfall and Huberger, 1942).Many diseased plants break at the point of infection on the stem,resulting in poor stands, and those which survive generally haveimpaired growth and low fruit yield. Collar rot has serious im-plications to tomato growers both as a disease and as a sourceof inoculum for an EB epidemic. The leaf blight phase normallyappears during the adult phase of the tomato plant, though itcan also be seen at earlier stages. It first appears as dark, smalland coalescing concentric lesions (target-like appearance), usu-ally on lower older leaves and progresses upward as the plantreaches maturity (Rotem, 1994). The tissue surrounding the le-sions turns yellow, senesces and the cells die through a nonhostspecific action of the pathogen, which produces toxic secondarymetabolites such as alternaric acid and zinniol (Lawrence et al.,2000), as discussed below. The leaves eventually either dry upor fall off, leading to complete defoliation of the plant towardsthe end of the season (Barksdale and Stoner, 1977). Defoliationreduces the photosynthetic rate and increases the respiration rateof healthy tissue, thereby disrupting the equilibrium between thesupply and demand for nutrients. Defoliated plants often result

in sun-scalded fruit due to the lack of foliage to protect it from thesun. Leaf blight is thus the most important phase of the disease.In tomato, as the disease progress on the leaves, dark sunkenspots also develop on the stems, causing stem lesions. Necroticlesions may also appear in floral parts and fruits, which occur inmid to late season and lead to significantly reduced fruit yieldand quality (Strandberg, 1992; Rotem, 1994).

C. The PathogenAlternaria solani is one of the best-known and economically

most important member of its genus and is a true member ofthe Kingdom Fungi in the Phylum Ascomycota (Order Pleospo-rales, Class Dothideomycetes) (Rotem, 1994). The genus nameactually refers to its asexual morphology, which is classifiedin the form-class, but its molecular phylogenetic placement inthe Ascomycota is well-established (Schoch et al., 2006; Tayloret al., 2007).

Historically, the genus Alternaria was first described by Neesin 1817 (cited in Neergaard, 1945) in the description of A.tenuis. Elliot (1917) tentatively reduced the genus Alternariato seven species, but his classification was not widely accepted.Angell (1929) and Wiltshire (1933) divided Alternaria into twotypes, characterized by their spore morphology: species withshort-beaked asexual spores (conidia), which were named ‘A.tenuis’, and species with conidial chains and long-beaked spores,which were named ‘A. porri’. This classification differentiatedAlternaria species so effectively that it formed the basis for Neer-gaard’s book (Neergaard, 1945). Sorauer (1896) had not madeany distinction between A. solani and A. tenuis and referred toboth as A. solani (cited in Neergaard, 1945). The dispute overnomenclature continued until Jones and Grout (1897) distin-guished between A. solani and A. tenuis based on isolation andinfection experiments. This distinction defined the authority forA. solani as Jones and Grout.

1. Disease CycleA. solani overwinters and survives as conidia or mycelia on

buried host debris and potato tubers, particularly in fields withpoor cultural practices such as continuous cropping of tomatoesor potatoes. Conidia may serve as a primary source of inocu-lum early in the next season until the secondary disease cyclestarts. Conidia have thick walls, which make them resistant tolow temperatures and other adverse environmental conditions.Therefore, cultural methods for controlling EB incorporate prac-tices of lengthy non-host crop rotation, debris sanitation and theutilization of aseptic seeds and transplants (discussed below).It is generally accepted that sporulation (production of coni-dia) is a two-phase process (Rotem, 1994). In the initial phase,conidiophores are produced. This phase is induced by near-ultraviolet wavelength radiation (310–400 nm) and enhanced bythe presence of light (Witsch and Wagner, 1955; Rotem, 1994).The second phase favorably occurs in the absence of light, inwhich conidia are formed on conidiophores. In vivo and in the

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presence of favorable conditions, conidia germinate. Tempera-ture and humidity affect production of conidiophores, sporula-tion and germination. Infection is favored by a relative humidityof >80% and moderate temperatures (optimal ∼27◦C). Underfavorable conditions, Alternaria spores are capable of germinat-ing on leaves of host (resistant or susceptible) as well as nonhostplants (Rotem, 1994). Germination usually does not require nu-trients, however, certain conditions such as leaching materialsfrom the invaded host tissue may facilitate the germination pro-cess. After germination, germ tubes emerge from spores andspread on the leaf surface. Infection occurs by direct hyphalpenetration of the foliar surface through stomata, dead epider-mal cells, or wounds caused by insects or sand particles carriedby strong winds (Jones, 1991; Strandberg, 1992). The germ tubedevelops from the inner layers of the spore wall within approx-imately 1 to 3 hours. Penetration begins 2 to 3 hours after germtube and appressoria development. After penetration through aninfection peg, the pathogen ruptures the epidermis and developsintercellularly in the mesophyll tissue. Cells adjacent to the in-fection site undergo hypertrophy. Cell content reduction occursin the distal cells. In later stages, the mesophyll shrinks and cellsdeform in the infection site, which causes cell death and symp-toms characteristic of the disease (Rotem, 1994). In advancedstages of EB, secondary infection starts by proliferation of newconidia on infected tissue.

2. Variability of Alternaria Isolates and Causes of VariabilityAlthough Alternaria species are known to reproduce asexu-

ally, they are highly variable in morphology, toxin production,and genetic composition both in vivo and in vitro (Henning andAlexander, 1959; Rotem, 1994; Weir et al., 1998; Akamatsuet al., 1999; Simmons, 2000; Van der Waals et al., 2004). Therehave been various hypotheses describing the high level of geneticvariation in Alternaria, including occurrence of heterokaryosis(Stall and Alexander, 1957; Stall, 1958) and natural mutation(Petrunak and Christ, 1992; Leung et al., 1993; Van der Waalset al., 2004). Heterokaryosis and asexual recombination, theso-called parasexual life cycle, were suggested to be more im-portant than natural mutation in causing genetic variation inimperfect fungi such as A. alternata. However, it is also knownthat heterokaryon incompatibility tends to block parasexualityin nature. Here we discuss the extents of variability at differentlevels.

a. Variability in Morphology. Different species in thegenus Alternaria are highly variable in their hyphal and conidialmorphology. For instance, the presence of beaked conidia vs.nonbeaked conidia can distinguish between distinct species butnot between closely-related/similar species (Rotem, 1994). Thelength of conidiophores and the dimensions of the spore body,including the beak, are other characteristics that can be usedin species differentiation. The vegetative compatibility (VC)test is another assay of measuring variability among isolates.In this test, two isolates already grown in separate Petri-dishesare placed adjacent to each other in one Petri-dish and their re-

actions are recorded after 1 to 2 weeks. A distinct line formsbetween the two isolates if they are different or incompatible,whereas an undistinguishable border forms between the two ifthey are the same or compatible isolates (Van der Waals et al.,2004).

In early studies, phenotypic variability was used for nomen-clature and classification of species within Alternaria. Subse-quently it was determined that morphological characteristicsalone were not reliable tools in differentiating among species asthey were influenced by many environmental factors, includingthe substrate, light intensity and the temperature under which thefungus grew (Vakalounakis and Christias, 1985). For instance,when the same isolate of A. macrospora was grown on differentorgans of cotton, the total length of spores varied from 101 µm to165 µm (Ling and Yang, 1941; Rotem, 1994). This kind of varia-tion led to the belief that host specialization was a more reliablemeasure of differentiation among Alternaria species (Rotem,1994). However, the diagnostic value of morphological charac-teristics should not be undermined. For example, in cases wherean isolate of Alternaria can infect more than just the commonhost, host specialization per se may not be a useful diagnosticmeasure (Rotem, 1994).

b. Variability at the Molecular Level. Determination ofgenetic variation at the molecular level and establishment ofmolecular phylogeny and taxonomy became feasible with theadvent of molecular markers in the 1980s. During the past twodecades, genetic variability among isolates of A. solani and otherAlternaria species has been extensively studied using differentmolecular markers, including isozymes and RAPDs (Petrunakand Christ, 1992; Cooke et al., 1998; Sharma and Tewari, 1998;Weir et al., 1998; Roberts et al., 2000; Morris et al., 2000; Varmaet al., 2006), RFLPs (Akamatsu et al., 1999), AFLPs (Bocket al., 2002; Perez Martınez et al., 2004), random amplifiedmicrosatellites (RAMS) (Van der Waals et al., 2004), nuclearinternal transcribed spacers (ITS), mitochondrial small subunits(mtSSU) and ribosomal DNAs (Pryor and Gilbertson, 2000).According to these studies, extensive genetic variation existsamong isolates of Alternaria spp. throughout the world, althoughregional populations are often genetically homogeneous (PerezMartınez et al., 2004; Van der Waals et al., 2004; Varma et al.,2006). The latter observation suggests that the dispersal rangeof Alternaria species may be confined within a limited geo-graphical range. Despite these studies, the molecular phylogenyand taxonomy of the genusAlternaria is far from being wellunderstood.

c. Variability in Toxin Production. Inter- and intraspecificvariation among Alternaria species is not limited to their mor-phology or DNA sequences. They are also highly variable intoxin production, especially those that produce host-specific tox-ins (HSTs), such as certain isolates of A. alternata. To date, atleast 10 phytopathogenic Alternaria species have been identified(Akamatsu et al., 1999). Seven of these cannot be differentiatedfrom saprophytic A. alternata based on morphological charac-teristics but they can be distinguished based on their unique

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pathogenicity (Akamatsu et al., 1999), which is conditioned byspecific toxins (Nishimura and Kohmoto, 1983; Kohmoto andOtani, 1991). Despite this variability, the concept of classifica-tion of Alternaria species based on their pathotype has not beenwidely accepted among plant pathologists (Yu, 1992; Simmons,2000; Simmons, 2003).

3. Physiological RacesPhysiological races are defined as specialized pathogen races

for pathogenicity on different genotypes of a host (Agrios, 2005).Such host genotypes are known as differential lines (Chaeraniand Voorrips, 2006). Although some reports claim to identifyphysiological races of A. solani, to date there is no confirmedevidence of the presence of such races. Bond (1929) first de-scribed the presence of physiological races of A. solani basedon different characteristics of the pathogen, including sporesize, pigment production, capacity for sporulation, intensity ofmycelial growth and tendency to saltate. Neergaard (1945) re-viewed Bond’s work and proposed more differential diagnosticscriteria. Henning and Alexander (1959) reported the existenceof physiological races based on different responses of tomatoplant to pure single-conidia derived cultures of A. solani. Suchobservations, however, could not be verified as after repetitionof the experiments different reactions of A. solani pure culturesto the same tomato genotype were observed. The authors at-tributed this to variation within isolates or within host lines.However, the lack of any known physiological races in A. solanisuggests that the host recognition may occur in a nonspecificmanner, thereby complicating pathological and genetic studiesof EB, as discussed below (Rotem, 1994; Lawrence et al., 1996;Lawrence et al., 2000).

4. Mechanisms of Pathogenicitya. Enzymatic Reactions. The pathogenicity mechanism

of A. solani has not been clearly determined despite recent ad-vances in molecular biology and molecular plant pathology.However, it is known that upon successful infection, conidiagerminate in both resistant and susceptible plants, resulting inthe production of germ tubes, which primarily penetrate into theleaf epidermis and grow in the intercellular spaces in a biotrophicmanner (Rotem, 1994). In this process, enzymatic action is themost likely means of pathogen penetration and movement. Highactivity of cellulase in culture filtrate of A. solani has been ob-served (Mehta et al., 1974), however, variation in cellulase activ-ity in vivo and in vitro has also been reported (Wasfy et al., 1977;Rotem, 1994). The limited knowledge available does not allowdrawing any unequivocal conclusion regarding the significanceof enzymatic reactions in A. solani pathogenicity.

b. Toxin Production. Toxins are low molecular weightcompounds that may cause histological and physiologicalchanges in host organisms. Toxins produced by plant pathogenicfungi are called phytotoxins, and may be host-specific (selec-tive) or nonspecific (nonselective). Host-specific toxins (HST)are usually toxic to a particular plant species or cultivar that

serves as the host, and are not usually toxic to other (nonhost)species. Conversly, nonselective toxins usually have a broaderrange of plant species that they can affect. In either case, toxinproduction is a part of the pathogenicity process. Toxins can haveconfounding effects on other pathogenicity processes, includingenzymatic reactions and production of specific recognition fac-tors (elicitors) (Rotem, 1994). Thus, interactions among thesefactors can complicate studies of the mechanisms of pathogenic-ity (Rotem, 1994). Much effort has been devoted to determinemechanisms of action of HSTs as well as non-HSTs in A. solani.It is speculated that unlike non-HSTs, HSTs completely destroythe membrane structure of the host (Park and Ikeda, 2008). Fur-thermore, partial dysfunction of organelles such as mitochondriaand chloroplasts by HSTs provides opportunity for the pathogento penetrate the host membrane. In below, we discuss studiesthat investigated production of HSTs and non-HSTs and theirimpacts on pathogenicity process in the genus Alternaria.

Nishimura and Kohmoto (1983) listed a number of HSTsproduced by A. alternata and other members of the same group,including A. mali, A. citri, A. kikuchiana, A. longipes, and A.alternata f. sp lycopersici. Previous studies had suggested dif-ferent HSTs produced by A. solani compared to other members.For example, Matern et al. (1978) had identified two lipid-likesubstances in A. solani cultures that were acting synergisticallyto produce symptoms of the disease. It was later determined thatthese substances were different from the commonly known com-pound, alternaric acid, which is not yet recognized as an HST(Langsdorf et al., 1990; Park and Ikeda, 2008) (see below). Al-ternaria solani also produces complex reduced-type polyketidescalled solanapyrones. Oikawa et al. (1998) identified compo-nents of phytotoxin solanapyrones A to E from A. solani culturefiltrates. The effects of formyl analogues of solanapyrones A andD were tested on germination of lettuce seeds, which showedthat they had inhibitory effect on roots and hypocotyls forma-tion at 250 ppm. Subsequently, an attempt was made to clonethe gene for solanapyrone A synthase in A. solani (Fujii et al.,2005). This research led to the identification of another gene(alt5) for polyketide synthase (PKSN), which was determinedto be responsible for production of a solanapyrone compoundin Aspergillus oryzae transformants. To date, however, there hasbeen no report of cloning of any of the solanapyrone synthasegenes in A. solani.

In addition to HSTs, there are several other compounds thathave been isolated from Alternaria culture filtrates that seem tobe common among all Alternaria sp. Two major non-HSTs, zin-niol and alternaric acid, have been recovered from A. solani cul-ture filtrates (Cotty et al., 1983; Maiero et al., 1991; Haraguchiet al., 1996). Zinniol induces stem wilting and leaf necrosis onzinnias but its effect on tomato is unknown. Altersolanol A isthe second major component of all tetrahydroanthraquinones ofA. solani (Okamura et al., 1993; Haraguchi et al., 1996). Thiscompound has antimicrobial, antiprotozoal and cytotoxic activ-ities. However, in tomato, potato, and pea it results in tissuenecrosis (Holenstein and Stoessl, 1983; Haraguchi et al., 1996).

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The presence of a number of other metabolites and nonhost-specific compounds such as Alternariol, Macrosporin, Tentoxinand Tenuzonic acid in Alternaria culture filtrates has been re-ported (Nishimura and Kohmoto, 1983).

Alternaric acid has been recovered from A. solani filtrate(Brian et al., 1952; Langsdorf et al., 1990). Alternaric acid pro-duces symptoms characteristics of EB on tomatoes, with leaflesions, chlorosis and necrosis (Maiero et al., 1991). Attempts toextract alternaric acid from A. solani culture filtrate have failed.However, it has been shown that genotypes that have resistanceto both EB and collar rot are tolerant to culture filtrate containingalternaric acid and zinniol, whereas genotypes that are resistantonly to EB exhibit susceptibility to culture filtrates containingthese two compounds (Maiero et al., 1991). No clear relationshipbetween the amount of alternaric acid production in isolates andsusceptibility of tomato cultivars has been determined, as somehighly virulent isolates produce small amounts of alternaric acid(Brian et al., 1952). It has been concluded that toxin susceptibil-ity is not a reliable marker when screening for resistance to A.

solani (Maiero et al., 1991). It is obvious that A. solani and otherAlternaria species can synthesize a number of toxins; however,isolation, purification, and characterization of these compoundshave remained a challenging task for chemists, biochemists, andplant pathologists.

D. Mechanisms of Host ResistanceIn recent years, several major resistance genes have been

cloned and characterized from various plant species, includingtomato, Arabidopsis and rice (Martin et al., 1993; Lin et al.,2007; Van-Damme et al., 2008). Successful cloning of diseaseresistance genes has dramatically advanced our understanding ofthe molecular basis of resistance and defense response in plants.However, only a few studies have examined the molecular basisof resistance to A. solani or the possible pathways involved inthis process.

The defense mechanisms used by plants in response to dis-eases caused by Alternaria sp. are likely similar to those usedin response to other phytopathogens. However, a hypersensitiveresponse to Alternaria is unlikely due to the necrotrophic natureof the pathogen (Rotem, 1994). In general, plants have evolvedmultiple mechanisms of resistance/defense to avoid diseases (Guand Martin, 1998; Eckardt, 2004; Park and Ikeda, 2008). Pro-duction of cuticular cell layers and an epicuticular wax layer isconsidered the first barrier/mechanism against pathogens. It isspeculated that the presence of wax increases the hydrophobic-ity of the leaf surface, which reduces the retention of waterborneinoculum. As a result, providing a dry leaf surface limits the ger-mination of A. solani conidia and the formation of germ tubes,thereby decreasing disease incidence. In addition, reflection ofcertain wavelengths of solar radiation from the cuticular surfaceacts as a barrier against pathogen attacks. The effects of waxyleaves have been reported in certain plant species including Bras-sicas (Conn and Tweari, 1989), but not in tomatoes or potatoes.

The second defense mechanism against diseases is microflora ofthe leaf surface. Several studies have demonstrated that certainmicroorganisms present on the leaf surface interfere with growthof pathogenic fungal hyphae (Norse, 1972; Tsuneda and Sko-ropad, 1977; Tsuneda and Skoropad, 1978; Hebbar et al., 1991).For example, it was demonstrated that spraying of a saprophyticisolate of A. alternata on tobacco plants that were infected by ahighly pathogenic isolate of A. alternata reduced disease inci-dence by 65% (Spurr and Main, 1974).

Secretion of phytoalexins is the third plant defense mecha-nism against pathogen attacks. Phytoalexins are low molecularweight compounds that are produced when a particular set ofgenes are rapidly activated in plants upon pathogen attack. Thisprocess is one of the fastest activation-dependent gene responsesin plants. Thomma et al. (1999) showed that deficiency in phy-toalexin production caused enhanced susceptibility of Arabidop-sis thaliana to Alternaria brassicicola. Several lines of evidencesuggest that resistance to A. solani most likely does not dependon detoxification of alternaric acid (Lawrence et al., 1996). How-ever, there might be a mechanism by which prior to and/or uponpathogen infection, the plant creates an antifungal environment.

Another plant defense mechanism is enzymatic reactions inresistant lines. For example, the role of hydrolytic enzymes inthe breakdown of fungal cell wall polysaccharides has beenwell demonstrated (Meins et al., 1992; Simmons, 1994). Currentknowledge of enzymatic reactions in resistant plant genotypesin response to A. solani is limited to results from a few studies.For example, it has been shown that constitutive production ofhydrolytic enzymes such as acidic or basic chitinase and β-1,3-glucanase are involved in conferring resistance to EB in tomatoresistant lines (Jongedijk et al., 1955; Lawrence et al., 2000).Also, Western blot analysis demonstrated that upon challengingtomato plants with A. tomatophila, the four isozymes of chiti-nase (26, 27, 30 and 32 kDa) were induced in both susceptibleand resistant lines (Lawrence et al., 1996). However, resistantlines (NCEBR-1 and NCEBR-2) had significantly greater activ-ity of the 30 kDa chitinase. In a more recent study, Lawrenceet al. (2000) determined that during A. solani infection, a highly-resistant breeding line (NC24-E) rapidly accumulated mRNAtranscripts coding for multiple pathogenesis-related (PR) genes,including antifungal isozymes of chitinase and β-1,3-glucanaseisozymes. It should be noted that such isozymes have not shownany antifungal activity in vitro. However, the observation ofelevated levels of both acidic (extracellular) and basic (intra-cellular) isozymes of PR proteins in tomatoes infected by A.solani suggested that multiple defense pathways such as sali-cylic acid (SA), ethylene and/or jasmonate–dependent pathwayswere also involved (Lawrence et al., 2000). This conclusion wasbased on previous studies where the effects of SA and ethylenewere examined in response to A. solani. For instance, applica-tion of exogenous SA on tomato roots caused enhanced expres-sion of the PR-1 gene (Spletzer and Enyedi, 1999) and sprayingtomato leaves with arachidonic acid increased the level of aPR-1 like protein (Coquoz et al., 1995). Furthermore, elevated

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expressions of ST-ACS4 and ST-ACS5 genes, members of the 1-aminocyclopropane-1-carboxylic acid gene that are precursorsin ethylene synthesis pathway, were observed in potato in re-sponse to A. solani (Schlagnhaufer et al., 1997). A transgenicapproach also has been employed to determine the role of β-1,3-glucanase and similar isozymes that enhance tomato EB resis-tance. Such studies have concluded that only a small percentageof the transgenic plants exhibited enhanced EB resistance (Cam-mue et al., 1992; Schaefer et al., 2005), suggesting the role ofother more important factors in EB resistance.

Another system in plant defense against pathogen attacks isinvolvement of metabolites and metabolic processes (Abu-Nadaet al., 2007). However, few studies have been conducted to fur-ther elucidate the role of metabolites in tomato EB resistance.For instance, in one study, tomatine, a common tomato glycoal-kaloid, was shown to be highly associated with A. solani resis-tance (Zhuchenko et al., 1975). In another study, total tanninsand phenols were suggested as possible metabolites for resis-tance to EB (Bhatia et al., 1972). However, it is expected thatrecent technological advancements in metabolic profiling usinggas chromatography and mass spectrometry (GC/MS) will helpbetter understand the functions of metobolites and underlyinggenes involved in tomato EB resistance.

In conclusion, rather limited knowledge is available with re-spect to the mechanism of EB resistance in tomato or potato.This is in part due to the complexity of the disease and the plantresponse, as well as the limited efforts devoted to characterizingthe mechanism(s). However, while there is a need for furtherresearch in this area, such shortage of the knowledge should notbe a major limiting factor to breeders’ efforts to identify andtransfer genes and QTLs for EB resistance in tomato.

E. Disease Screening MethodsIdentification of a reliable and fast disease screening method

has been a major obstacle in breeding tomatoes (and potatoes) forEB resistance. While research has shown a relatively good cor-relation between field and greenhouse screenings, the agreementbetween field or greenhouse screening and detached-leafletsevaluation has been controversial (Locke, 1948; Foolad et al.,2000). Here we discuss different methods that have been em-ployed for screening tomatoes for EB resistance.

1. Inoculum PreparationAlterinaria solani can grow on different culture media, in-

cluding potato-dextrose agar (PDA) (Locke, 1948), V-8 juiceagar (Vakalounakis, 1991), lima bean agar (Maiero et al., 1989),and Sporulation medium (Shahin and Shepard, 1979). AlthoughA. solani can grow on different media, specific conditions canincrease efficiency of mycelia growth and sporulation. For in-stance, incubation of fungus culture at 21◦C to 23◦C under acool-white fluorescent diurnal light with a 12-h photoperiod in-creases the efficiency of sporulation (Barksdale, 1969). In ad-dition, in vitro sporulation can be induced by providing stress

conditions such as wounding or transferring mycelia to minimalmedia (Shahin and Shepard, 1979). However, when proper con-ditions are provided, after 10 to 14 days conidia can be collectedby flooding the plates with ddH2O (containing 0.01% Tween20) and brushing the agar surface with a paintbrush (Fooladet al., 2000). The spore density in the suspension culture canbe counted using a hemacytometer. Normally, a spore densitybetween 10,000 and 40,000 conidia/ml is used for inoculation.The method of inoculation depends on the screening method,which may include field, greenhouse or growth chamber (us-ing detached leaflets) screening, as described elsewhere (Fooladet al., 2000).

2. Field ScreeningField evaluation has been the most utilized method of screen-

ing tomatoes for EB resistance (Barksdale and Stoner, 1977;Gardner, 1984; Nash and Gardner, 1988a). The advantages offield screening include the ability to grow large populations,evaluating plants under natural conditions, and recording dis-ease progress throughout the entire life cycle of the plant. Thedisadvantages include preparation of large amount of inocu-lum (in cases where natural inoculum is not present), potentialnon-uniformity of inoculation and disease pressure, the need forgrowing disease spreader rows, and the intensive labor requiredfor evaluation of a large area. Additional potential problemsinclude confounding effects of other foliar diseases of tomatopresent in the field, the possibility of having environmental con-ditions not conducive to the disease, and variation in climaticconditions from year to year. Furthermore, in most areas proneto EB development, field experiments could be conducted onlyonce a year if all other conditions are optimal, limiting the breed-ing progress.

Field evaluation is usually conducted throughout the plant’sgrowing season, starting with observation of the first diseasesymptoms, usually 2 to 3 weeks after initial inoculation, andending with a recording of final percent defoliation at the end ofthe season. Disease severity is typically expressed as percent de-foliation (Horsfall and Barratt, 1945) and the data expressed overtime is used to determine the AUDPC. Although AUDPC andfinal percent defoliation are the most common criteria used forevaluation of EB resistance, other indices used include percentof disease index (PDI) and cumulative disease index (CDI) foreither stem or foliage infections (Thirthamallappa et al., 2000;Chaerani et al., 2007). In general, however, field evaluation ishighly useful as the data can be used to compare across plantgenotypes at various time intervals during the season. This kindof information can be particularly useful when breeding EB re-sistant tomatoes for targeted environments.

3. Greenhouse ScreeningGreenhouse (GH) evaluation, using seedlings or cuttings, has

been used for screening tomatoes for EB resistance with vari-able results (Andrus et al., 1942a; Barksdale, 1971; Nash andGardner, 1988a; Gardner, 1990; Maiero et al., 1990b; Poysa and

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Tu, 1996; Foolad et al., 2000). For instance, when lines withmoderate levels of resistance were compared with susceptiblelines, symptoms were not sufficiently different in most green-house tests to discriminate among genotypes (Barksdale, 1969;Nash and Gardner, 1988b). Other studies, involving F1 and back-cross progeny of a cross between a resistant S. habrochaitesaccession and a susceptible S. lycopersicum line indicated dif-ferential resistance among genotypes (Gardner, 1984). It ap-pears that unless the variation in disease response in susceptibleand resistance lines is large, the differences may not be eas-ily distinguishable in GH screening (Gardner, 1990). Thus, GHscreening may be useful to screen large numbers of accessionsin preliminary screenings to eliminate the very susceptible lines.Other factors affecting success of GH screening include plantage, inoculum quality and quantity, inoculation technique, andpre- and postinoculation environmental conditions (Strandberg,1988; Evans et al., 1992; Vloutoglou and Kalogerakis, 2000).Plant age is particularly important as susceptibility to EB usu-ally increases with maturity. Evaluation for leaf blight resistanceduring the seedling stage may not be useful as it may not reflectthe true response of a mature plant to EB infection. However,GH evaluation may be particularly useful for screening for stemlesion resistance, as describe elsewhere (Gardner, 1990). Properconditions for inoculation and successful GH screenings havealso been described elsewhere (Barksdale, 1969; Foolad et al.,2000). A few studies have investigated the relationship betweenfield and GH screenings with a general conclusion of the pres-ence of a good correlation between the two screening methods(for a review see Foolad et al., 2000). The observation of a goodcorrelation between GH and field evaluations indicates that ini-tial EB screenings may be conducted in the GH, assuming thatproper conditions are provided. Furthermore, we have experi-enced that tomato screening for EB resistance in the GH cansuccessfully be conducted during the late seedling and earlyflowering stages, when plants are 7 to 8 weeks old, and thatit is not necessary to wait until fruiting (Foolad et al., 2000).Thus, GH screening can significantly speed up the EB resis-tance breeding process by allowing multiple cycles of selectionin one year. Moreover, GH evaluation should be particularlyuseful in regions where field screening is ineffective due to un-suitable environmental conditions or when other diseases whichcause plant defoliation are also present (e.g., septoria leaf spot).

4. Growth Chamber Screening Using Detached LeafletsThe detached-leaflet method was first used by Douglas (1922)

to compare the response of tomato varieties to inoculation withan undetermined species of Alternaria. Subsequently, the useof detached leaflets for comparing the virulence of strains of A.solani and evaluating the resistance of tomato genotypes wasreported to be either unsuccessful (Wellman, 1943) or success-ful (Locke, 1948). A recent study determined the most effi-cient screening procedure for EB resistance in tomato, where29 tomato genotypes from different species were evaluated forresistance in replicated field and greenhouse trials and in growth

chamber using detached-leaflets (Foolad et al., 2000). While thisstudy indicated a good correspondence between the field andgreenhouse evaluations, there was no agreement between resis-tance at the detached-leaflet level and that in the field or in thegreenhouse. The authors dismissed the adequacy of detached-leaflet assay for screening tomatoes for EB resistance and sug-gested the utility of GH screening as a faster method than thefield screening.

F. Nongenetic Measures of Disease ControlThere is no commercial cultivar of tomato with sufficient

level of resistance to EB (discussed below). The most commonmeasures of disease control are cultural practices, including sani-tation, crop rotation, elimination of volunteer tomato and potatoplants in and around the field, maintenance of host vigor viaadequate nitrogen and phosphorus fertilization, reduction of fo-liar wetness through soil-directed irrigation systems, and heavyuse of fungicides (Agrios, 1988; Maiero et al., 1990a; Rotem,1994; Kucharek, 2000; Foolad et al., 2002a; Foolad et al., 2002b;Narayanasamy, 2002; Chaerani and Voorrips, 2006). Fungicideapplications may begin 1 to 2 days after transplanting and con-tinue on a 5 to 7-day schedule thereafter. However, in areaswith high humidity and frequent rainfalls these control mea-sures may have limited success due to high disease pressure. Inaddition, fungicide applications increase production costs andare environmentally unsafe. For example, in some areas withhigh humidity up to 15–25 fungicide applications per growingseason may be needed to obtain sufficient protection against EB(Sherf and MacNab, 1986). Also, frequent fungicide applica-tions promote development of new fungal biotypes, which maybe fungicide resistant. Furthermore, a growing number of chem-icals are being withdrawn from agricultural use due to concernsover their safety, presenting the tomato industry with a majorchallenge. Therefore, an integrated pest management strategy,focusing on exploitation of genetic resistance along with preven-tive cultural practices, must be adopted for achieving a long-termand sustainable control of EB.

G. Genetic Measures of Disease Control1. Genetic Sources of Resistance

Sources of genetic resistance to EB in tomato have been iden-tified mainly within the related wild species S. habrochaites, S.pimpinellifolium, and S. peruvianum but also infrequently in thecultivated species S. lycopersicum, as discussed below.

a. Solanum lycopersicum (formerly Lycopersicon es-culentum). Complete resistance to EB in the cultivatedspecies of tomato is rare. Brock (1950) examined reactionsof 109 tomato varieties against 5 isolates of A. solani undergreenhouse conditions and none exhibited resistance. In 1967,USDA researcher R. E. Webb observed field resistance intomato breeding lines 67B833 and 68B134, which later werereleased to growers and plant breeders and used to developother resistant breeding lines (Barksdale, 1971; Barksdale

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and Stoner, 1973; USDA, 2007). Barksdale (1969) evaluatedseveral breeding lines and accessions of S. lycopersicumand found a resistant accession (PI138630), which was laterused for EB resistant breeding. This led to the developmentand release of resistant breeding lines 71B2 and Campbell1943 (C1943) (Barksdale and Stoner, 1977). Although 71B2displayed good resistance to leaf blight, it was susceptibleto collar rot. In contrast, C1943, a genotype otherwise withunknown background from the Campbell Institute for Agri-cultural Research, exhibited a high level of resistance to stemlesion (collar rot) and a moderate resistance to leaf blight.These two lines were subsequently used in other programs forEB resistance breeding. For example, C1943 was used as asource of EB resistance in developing breeding lines NC63EB,NC870, NCEBR-2, NCEBR-3 and NCEBR-4 (Gardner, 1988)(http: / /www.ces.ncsu.edu/fletcher/programs/tomato/releases/pollenlines.html). Subsequently, 71B2 was also used as asource of resistance in developing tomato breeding linesNCEBR-5 and NCEBR-6 (Gardner, 2000). The new resistantbreeding lines developed by RG Gardner have been used fordeveloping hybrid cultivars with improved EB resistance, asdiscussed below. Only a few other studies have found anyuseful source of EB resistance within the cultivated species oftomato. Poysa and Tu (1996) identified 11 moderately resistantS. lycopersicum accessions, but it is not known whether anyof them was used for EB resistance breeding. Andrus et al.(1942a) identified a number of varieties and S. lycopersicum PIsas sources of resistance to collar rot. These included varieties‘Devon Surprize,’ ‘Red Cherry,’ ‘Red Pear 414,’ ‘Red Pear 415,’‘Targinnie Red,’ ‘Vetomold,’ ‘Montgomery,’ ‘Norduke,’ and‘Marglobe,’ as well as USDA Plant Introductions PI127814,PI128602, PI126452, PI118324 and PI95558. In an extensivegermplasm screening experiment, including cultivated andwild species of tomato, Martin and Hepperly (1987) found oneaccession (PI406758) of S. lycopersicum f. sp. cerasiformehighly resistant and 6 accessions with moderate resistance toEB. However, there has not been any report of the use of any ofthese accessions in breeding for EB resistance in tomato.

b. Solanum habrochaites (formerly Lycopersicon hirsu-tum). A greater number of EB resistant accessions has beenidentified in S. habrochaites than in any other tomato species.An evaluation of 488 accessions of different tomato Solanumspecies under field conditions in Ohio and Indiana led to theidentification of ten S. habrochaites accessions with resistanceto EB and Septoria leaf spot (caused by Septoria lycoper-sici) (Alexander et al., 1942). Andrus et al. (1942b) reporteda high degree of EB resistance in F1 progenies of crosses be-tween S. habrochaites and S. lycopersicum. Locke (1949) re-ported an accession (PI127827) of S. habrochaites with highresistance to EB. Another highly resistant S. habrochaites ac-cession (PI126445) was identified by Alexander and Hoover(1955), which was subsequently utilized in many tomato ge-netics and breeding programs (see below). Martin and Hepperly(1987) reported presence of 7 highly resistant S. habrochaites ac-

cessions, including PI390513, PI390514, PI390516, PI390658,PI390660, PI390662 and PI390663.It was subsequently demon-strated that hybridization between these accessions and the cul-tivated tomato resulted in F1hybrids with moderate to high levelsof EB resistance. Screening of ∼200 tomato genotypes by Poysaand Tu (1996) resulted in the identification of a few highly resis-tant S. habrochaites accessions, including LA2100, LA2650 andPE36. More recently, screening of 29 tomato genotypes from dif-ferent species identified several new S. habrochaites accessionswith resistance to EB (Foolad et al., 2000). Obviously, the greatgenetic diversity within S. habrochaites has presented opportu-nities to identify sources of EB resistance for use in genetics andbreeding studies, as discussed below.

c. Solanum pimpinellifolium (formerly Lycopersiconpimpinellifolium). After S. habrochaites, the best sourceof EB resistance in tomato is the red-fruited wild speciesS. pimpinellifolium. This species is the most closely relatedwild species of tomato, and the only one from which naturalintrogression into the cultivated type has been observed (Rick,1982; Miller and Tanksley, 1990). In addition, extensive geneticintrogression from this species into the cultigen has been madethrough plant breeding for various traits, including diseaseresistance and high fruit quality. Solanum pimpinellifoliumaccessions are highly self-compatible and bi-directionally crosscompatible with the cultivated tomato. Because of the closephylogenetic relationship, there is little or no difficulty ininitial crosses between S. lycopersicum and S. pimpinellifoliumor in subsequent generations of pre-breeding and breedingactivities. In addition, there are fewer undesirable horticulturaland agronomic characteristics in S. pimpinellifolium, whichmake it more desirable to exploit than the distantly related wildspecies S. habrochaites or S. peruvianum. Because of theseimportant considerations, accessions of S. pimpinellifolium areoften highly favorable for use in tomato breeding programs andfor basic studies of biologically important traits.

Reynard and Andrus (1945) reported that S. pimpinellifoliumaccessions PI179532 and PI127833 had good levels of resis-tance to collar rot but were susceptible to foliar phase of theEB disease. Martin and Hepperly (1987) described 11 highlyand 6 moderately resistant accessions of S. pimpinellifolium ina field screening, and were examined as sources of EB resis-tance in crosses with the cultivated tomato. Kalloo and Banerjee(1993) reported an accession of S. pimpinellifolium (A1921)with unknown origin as being highly resistant to EB underfield conditions. Perhaps the largest germplasm screening ofS. pimpinellifolium in search of EB resistance was done re-cently at the Pennsylvania State University Tomato BreedingProgram (M.R. Foolad, unpublished data). Approximately 270S. pimpinellifolium accessions were evaluated for resistance un-der field, greenhouse and growth chamber (detached leaflets)conditions. Several accessions were identified with strong resis-tance to EB, and one of them, named PSLP125, was chosen andused for extensive genetics and breeding research, as discussedbelow.

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d. Solanum peruvianum (formerly Lycopersicon peru-vianum). A number of studies have identified accessionswithin S. peruvianum with resistance to EB. Unlike S. pimpinel-lifolium, S. peruvianum is a highly variable species in part dueto its outcrossing nature enforced by the presence of strong self-incompatibility within the species (Taylor, 1986). Often thereis also extensive genetic variation within each accession. Fur-thermore, the use of this species in breeding programs is notwithout inherent difficulties. The presence of reproductive bar-riers in the initial cross and early backcross generations, reducedviability and sterility of the hybrid, segregation distortion andreduced recombination in the segregating generations, often in-terferes with the use of this species as a gene resource. Also,because of considerable genetic variation within each accession,no particular accession can be identified as resistant or suscep-tible. For example, Barksdale (1969) described S. peruvianumaccession PI129152 and S. peruvianum var. humifusum acces-sion PI127829 as resistant to EB. However, due to the presence ofheterogeneity within these accessions, he suggested that selec-tion of the most resistant plants within each accession should bea more useful approach for utilizing these germplasms. Poysaand Tu (1996) identified six “resistant” S. peruvianum acces-sions, including PE33, LA1292, LA1365, LA1910, LA1983 andPI270435, however, none has been utilized for EB resistancebreeding in tomato.

2. Inheritance StudiesMany of the studies of EB inheritance have focused on the

leaf blight phase of the disease with little attention to collar rotor fruit rot. The few studies that did investigate the collar rot(stem lesion) or fruit rot phase of the disease generally con-cluded that resistance was possibly independent of that for theleaf blight phase (Reynard and Andrus, 1945; Maiero et al.,1990b). In general, EB (leaf blight) resistance in tomato doesnot follow the gene-for-gene model of vertical/qualitative re-sistance proposed by Flor (1971), where the specificity of aresistant host against its avirulent pathogen is determined by theinteraction of their respective R and avr genes. Also, no hy-persensitive response (HR) has been reported for EB resistancein tomato or in other Solanaceae species. In contrast, EB hasbeen characterized as a complex quantitative trait, controlledby the additive and nonadditive interaction effects of multiplegenes, which are highly influenced by various environment fac-tors (Nash and Gardner, 1988b). In some tomato lines, the inher-itance of EB resistance was reported as being quantitative andrecessive (Barksdale and Stoner, 1977; Maiero et al., 1990a),whereas in other lines as quantitative and partially dominant,with epistasis involved (Martin and Hepperly, 1987; Gardner,1988; Nash and Gardner, 1988b). The presence of modifyingfactors with both dominant and recessive effects has also beensuggested (Reynard and Andrus, 1945; Martin and Hepperly,1987). The heritability (h2) of EB resistance has been reportedto be generally low to moderate. For example, Nash and Gardner(1988a) reported a narrow sense h2 estimate of 0.26–0.38 based

on parent-offspring regression analysis. Application of the samemethod using different interspecific (BC1 and BC1S1) and in-traspecific (F2 and F3) populations of tomato led to h2 estimatesof 0.65–0.75 for EB resistance (Foolad and Lin, 2001; Fooladet al., 2002a). Most recently, a RIL population of tomato basedon a S. lycopersicum × S. pimpinellifolium cross was evaluatedfor EB resistance in different generations (F7 to F10) and yearsand h2 for EB resistance was estimated. The estimates based onpair-wise correlation analyses varied from 0.53 to 0.65 and basedon a restricted maximum likelihood (REML) analysis, combinedwith mixed models with replications over the environments, was0.56 (Ashrafi, 2007). The low to moderate nature of h2 for EBresistance in tomato suggests that improvement of this trait intomato via traditional phenotypic selection is possible but it maybe slow.

Tomato EB resistance is also influenced by plant physiolog-ical age, growth habit, foliage morphology, earliness in matu-rity, fruit load, and nutritional status. For example, under fieldconditions, EB infection in tomato plants usually occurs afterflowering and fruit set, though under optimal conditions infec-tion may also occur at earlier stages. Generally older, senesc-ing leaves are more susceptible to EB than young, immatureleaves (Barksdale, 1971; Martin and Hepperly, 1987; Nash andGardner, 1988b; Maiero et al., 1990a). Studies of sporulation inAlternaria concluded that photosynthesis and high sugar con-tent in younger leaves play an inhibitory role in the productionof spores at early stages of pathogenesis. Early blight infec-tion is also positively correlated with earliness-in-maturity andnegatively correlated with fruit load (Nash and Gardner, 1988a;Foolad et al., 2000; Scott and Gardener, 2007). Late-maturingand/or low-yielding plants appear to be more resistant, thoughthey may not possess genetic resistance. Plants with indetermi-nate growth habit or self-incompatibility (or sterility) may alsoappear resistant, as they may outgrow the disease or because ofreduced stress resulting from lower fruit to foliage ratio. In con-trast, plants with early maturity, small and determinate growthhabit, or those that are nutritionally stressed exhibit a higher sus-ceptibility to the disease. Furthermore, plants with curled foliageare generally more susceptible to EB, while those with potatoleaf foliage are less affected compared to those with normal, ser-rated leaves (Scott and Gardener, 2007). These factors contributeto the difficulty in breeding for EB resistance, in particular whenusing traditional phenotypic selection. To circumvent some ofthese difficulties, during the past two decades efforts have beenmade to facilitate selection and breeding for EB resistance byidentification of genetic markers which may be associated withtrue resistance. Once such associations are identified, the geneticmarkers can be used as indirect selection criteria for breedingfor EB resistance, as discussed below.

3. Genetic MappingTo develop a better understanding of the genetic controls

of EB resistance and to facilitate marker-assisted breedingfor improved resistance, a number of studies have identified

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96 M. R. FOOLAD ET AL.

QTLs conferring EB resistance in tomato. These studies usedsegregating populations derived from interspecific crosses be-tween the cultivated tomato and related wild species, mainly S.habrochaites and S. pimpinellifolium, as briefly discussed below.

a. S. lycopersicum × S. habrochaites. The first QTL map-ping study of EB resistance in tomato employed backcross pop-ulations of a cross between a susceptible tomato breeding line(NC84173) and a highly-resistant S. habrochaites accession(PI126445) (Foolad et al., 2002b). NC84173 is an advancedtomato breeding line with a determinate (spsp) growth habitand midseason maturity, and PI126445 is a self-incompatibleaccession with an indeterminate (sp+sp+) growth habit, a vig-orous vine and late maturity (Gardner, 1984; Nash and Gard-ner, 1988b; Foolad et al., 2000). All of the latter characteristicsaffect plant response to EB, as described earlier. However, inthe QTL mapping study (Foolad et al., 2002b), to avoid con-founding effects of such factors on disease evaluation, all plantsthat were self-incompatible, late maturing, indeterminate, or lowyielding were eliminated from the BC1 mapping population.A total of 145 BC1 plants, which were self-compatible withhigh yield, early-to-midseason maturity, and determinate growthhabit, were chosen for EB evaluation and QTL mapping. TheBC1 plants were also self-pollinated to produce BC1S1 progeny.The progeny population, consisting of 145 BC1S1 families, wasgrown and evaluated for EB symptoms in replicated field trials intwo subsequent years. The final percent defoliation and/or AU-DPC were determined in all three years of screening and usedfor QTL mapping. Altogether, about 10 QTLs for EB resistancewere identified, with individual effects ranging from 8.4% to25.9% and combined effects of >57% of the total phenotypicvariation (PVE). All QTLs had positive alleles from the disease-resistant wild parent, PI126445. QTL locations were consistentacross generations and years (Fig. 1), suggesting their stabil-ity and potential utility for improving tomato EB resistance viaMAS. Chromosomal locations of some resistance gene analogs(RGAs) coincided with the locations of some QTLs, suggest-ing their possible involvement with EB resistance (Foolad et al.,2002b).

In a separate study, a “selective genotyping” approach wasused to identify and confirm QTLs for EB resistance in PI126445(Zhang et al., 2003). In this study, in a population of 820 BC1

plants of a cross between NC84173 and resistance accessionPI126445, the most resistant (5.6%) and most susceptible (3.7%)plants were selected and subjected to marker genotyping. A trait-based marker analysis detected seven QTLs for EB resistanceon chromosomes 3, 4, 5, 6, 8, 10 and 11, as shown in Fig. 1.Of these, all but the QTL on chromosome 3 were contributedfrom the resistant parent PI126445. Four of these QTLs werethe same as those identified in the previous study (Foolad et al.,2002b). The high level of correspondence between the resultsof two studies indicated the reliability of the detected QTLs andtheir potential utility for improving tomato EB resistance viaMAS breeding.

b. S. lycopersicum × S. pimpinellifolium. As mentionedearlier, through an extensive screening of wild species S.pimpinellfolium, an accession (PSLP125) was identified withgood resistance to EB (M.R. Foolad et al., unpublished data). Toidentify and map QTLs for EB resistance in this accession, filialpopulations were developed from crosses between PSLP125 andtomato breeding line NCEBR-1. NCEBR-1 has some level of fo-liar resistance to EB introgressed from S. habrochaites accessionPI126445 (Gardner, 1988). Initially, F2, F3 and F4 populationsof the cross between NCEBR-1 and PSLP125 were evaluatedfor EB resistance under field conditions. A genetic linkage mapwas developed based on the F2 population and used for QTLmapping in the F2, F3 and F4 generations (M.R. Foolad et al.,unpublished data). A total of ∼10 QTLs for EB resistance wereidentified on chromosomes 2, 3, 4, 5, 6, 7, 9 and 12 (Fig. 2),with individual effects ranging from 7.6% to 13.4% and com-bined effects of ∼ 44% of the total phenotypic variation. Ofthese, 7 QTLs were contributed from the S. pimpinellifoliumparent (PSLP125) and three from the S. lycopersicum parent(NCEBR-1). Subsequently, to confirm these QTLs, a selectivegenotyping approach, using a very large F2population of thesame cross, was employed for QTL mapping. Nine QTLs forEB resistance were detected on chromosomes 1, 2, 3, 4, 5, 6 and11 (M.R. Foolad et al., unpublished data), of which 6 were thesame as those identified in the unselected F2, F3 and F4 popu-lations (Fig. 2). Once again, the high level of correspondencebetween the results of these two studies suggested the reliabilityof the detected QTLs and their potential utility for improvingtomato EB resistance via MAS breeding.

Most recently, a RIL population of the same S. lycopersicum× S. pimpinellifolium cross was developed (M.R. Foolad et al.,unpublished data). Initially, an F7 generation of this populationwas used to develop a linkage map using ∼282 molecular mark-ers (Ashrafi, 2007). Subsequently, F7, F8, F9 and F10 generationsof this population were grown and evaluated for EB resistanceunder field conditions and were used for QTL mapping. SeveralQTLs were found on different chromosomes, of which two onchromosomes 5 and 6 were stable across 3 to 4 generations andyears and two on chromosomes 2 and 9 were consistent in atleast two generations. The individual effects of these QTLs var-ied from 7% to 30%, with a total phenotypic effect of ∼40%.There was also a good level of consistency between QTLs iden-tified in the initial (F2-F4) study and in the RILs (F7-F10). Forexample, four QTLs were stable in the 7 years (7 generations),suggesting the reliability of the identified QTLs and the powerof QTL mapping in identifying important QTLs. The resultsof these mapping studies indicate the potential utility of theidentified QTLs for improving tomato EB resistance via MASbreeding.

The overall results of the above studies indicated that whilesimilar QTLs were identified in different generations of thesame cross, generally different QTLs were identified in pop-ulations derived from different interspecific crosses. The results

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Downloaded By: [CDL Journals Account] At: 18:23 16 December 2008

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98

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BLIGHT RESISTANCE IN TOMATO 99

suggested stability of QTLs across environments and genera-tions but QTL variation in different genetic backgrounds. Itis expected that pyramiding of resistance QTLs from differ-ent sources would result in tomato genotypes with strong anddurable resistance to EB. Further inspections of the above resultshas led to the identification and selection of six QTLs with stableand independent effects for use in MAS to improve tomato EBresistance (M.R. Foolad, unpublished data).

c. S. lycopersicum × S. peruvianum. Chaerani et al.(2007) evaluated F2 and F3 populations of a cross between EB-resistant S. peruvianum accession LA2157 and a susceptible S.lycopersicum genotype and identified a total of 6 QTLs for EBresistance in the two generations. However, because they useddifferent environments and disease scoring parameters in theF2 and F3 generations, QTLs were not consistent across gen-erations. Nevertheless, in F3 a few QTLs were identified whichappeared to have contributed to resistance to both foliar and stemlesion phases of the disease. If confirmed, such QTLs would bevery useful for MAS breeding of tomatoes with EB resistance.

4. Breeding for Early Blight ResistancePathotype non-specific resistance is usually controlled by

multiple genes (or QTLs), and thus plants carrying such traitsare usually resistant to multiple races of the pathogen. Thistype of resistance is generally more durable than single-generesistance. However, phenotypic evaluation and selection forpathotype nonspecific resistance are often difficult because somequantitative measures of the disease severity, disease epidemic,or specific disease parameter(s) are necessary for selecting re-sistant plants. In addition, phenotypic evaluation for such traitsis often influenced by uncontrollable environmental factors aswell as plant physiological characteristics such as maturity andplant type and size. This complexity has certainly contributed tothe limited success in developing EB-resistant tomato cultivarsthrough traditional breeding.

Research to develop EB-resistant tomato cultivars using tradi-tional protocols of plant genetics and breeding began more thanhalf-a-century ago and has continued since. Despite the com-plex nature of the disease and the host resistance, to date severalbreeding lines and cultivars with measurable levels of resistancehave been developed. Early resistance breeding focused mainlyon the collar rot phase of the disease due to the severity of thisphase in field grown transplants before the advent of effectivefungicides (Scott and Gardener, 2007). For example, ‘Campbell1943’ (also known as C1943) was developed by G.B. Reynard(of Campbell Institute for Agricultural Research) with collar rotresistance and moderate resistance to leaf blight (Andrus et al.,1942a; Reynard and Andrus, 1945; Barksdale, 1969). In 1970,the USDA tomato-research program in Beltsville, MD, releasedtwo breeding lines, 67B833 and 68B134, with some level of re-sistance to EB; a more advanced processing breeding line, 71B2,with higher levels of leaf blight resistance but susceptibility tostem and fruit lesion phases, was released later (Barksdale andStoner, 1973; Barksdale and Stoner, 1977). Maiero et al. (1990a)

measured general and specific combining ability of several resis-tant lines, including USDA line 87B187. This line was originallydeveloped fromS. habrochaites accession PI390662 (Barksdale,1969). Another S. habrochaites accession, B6013, was used todevelop three EB resistant lines, H-7, H-22 and H25, under fieldconditions in a backcross-breeding program (Kalloo and Baner-jee, 1993). Although the authors observed association betweenEB resistance and lateness in maturity and small fruit size inB6013, they reported that the derived breeding lines had shortermaturity time (69 to 75 days) and larger fruit size than B6013.Kalloo and Banerjee (1993) also identified an accession of S.pimpinellifolium (A1921) with unknown origin as highly resis-tant to EB under field conditions. Hybridization of this accessionwith cv. ‘HS-101’ followed by a few generations of backcrossbreeding resulted in the development of BC4F4 progeny exhibit-ing as much resistance as the original S. pimpinellifolium acces-sion (Kalloo and Banerjee 1993).

R.G. Gardner at the North Carolina State University initi-ated breeding for EB resistance in tomato in the late 1970s. In1988, two moderately-resistant tomato breeding lines, NCEBR-1 and NCEBR-2 were released (Gardner, 1988; Nash and Gard-ner, 1988a). NCEBR-1 derives foliar resistance from the S.habrochaites accession PI126445 but is susceptible to collarrot, and NCEBR-2 derives moderate foliar and collar resis-tance from C1943. Subsequently, several other fresh-markettomato breeding lines with EB resistance were released bythe same program, which were named NCEBR-3 to NCEBR-8 (http://www.ces.ncsu.edu/fletcher/programs/tomato/releases/index.html). Briefly, NCEBR-3 and NCEBR-4 derive EB re-sistance from both C1943 and NCEBR-1 (Gardner and Shoe-maker, 1999). NCERB-5 is an F8 selection with breeding linesPiedmond and EB-resistant 71B2 in its background (Gardner,2000). NCEBR-6 has a more complex pedigree, but with EBresistance also derived from 71B2 (Gardner, 2000). NCEBR-7is moderately resistant to EB and has NCRBR-5 and NCRBR-6in its background ( Gardner, 2000; Gardner, 2006). NCEBR-8is a fresh-market plum (Roma type) tomato, which is moder-ately resistant to EB and has Florida 7547 and NCEBR-6 in itsbackground.

A few hybrid cultivars with EB resistance have also beenreleased by the same program at the North Carolina State Uni-versity. For example, a cross between NCEBR-3 and NCEBR-4resulted in the development of Mountain Supreme, which hasEB resistance (from C1943) and can extend the fungicide sprayinterval from 5 to 10 days (Gardner and Shoemaker, 1999). Across between NCEBR-5 and NCEBR-6 resulted in the com-mercial cultivar ‘Plum Dandy,’ which has moderate EB resis-tance (Gardner, 2000). NCEBR-7 was developed from ‘PlumDandy’ and was crossed with NCEBR-8 to produce the com-mercial hybrid ‘Plum Crimson,’ with EB resistance comparableto ‘Plum Dandy’ but also with high fruit lycopene content (Gard-ner, 2006). Ingeneral, the NC lines and hybrids can tolerate anextended fungicide spray interval and contribute to a signifi-cant reduction in chemical inputs for EB control. It should be

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100 M. R. FOOLAD ET AL.

noted, however, that none of the currently available resistantbreeding lines or cultivars exhibit a resistance level comparableto the original parental line of the wild species S. habrochaitesor the F1 progeny generation. It appears that some of the re-sistance components were lost through successive generationsof phenotypic selection and backcrossing to the susceptible S.lycopersicum lines (Gardner, 1988; Gardner, 1990).The lowerlevel of resistance in the advanced breeding lines could also bedue to additional stresses imposed by producing determinateplants with earlier maturity and higher fruit set compared to theoriginal wild accession (Scott and Gardener 2007).

Breeding efforts at the Pennsylvania State University to de-velop tomatoes with improved EB resistance were initiated dur-ing the late 1990s. Much of these efforts have been focused onthe identification and transfer of EB resistance from the tomatowild species S. pimpinellifolium. As discussed earlier, some newsources of EB resistance were identified, which have been char-acterized and utilized for breeding purposes. A major goal hasbeen to develop tomato breeding lines and cultivars with im-proved EB resistance and acceptable yield and maturity time.Using traditional breeding protocols of phenotypic selection andbreeding, fresh market and processing tomatoes with improvedEB resistance, derived from S. pimpinellifolium have been de-veloped. These materials are in the process of final evaluationand release (M.R. Foolad, unpublished data). In addition, as in-dicated earlier, QTLs for EB resistance have been identified inaccessions of S. habrochaites and S. pimpinellifolium. A num-ber of these QTLs have been verified, which can be used forimproving tomato EB resistance via MAS breeding.

IV. CONCLUSION AND FUTURE PROSPECTS

A. Late BlightMore than 150 years ago, LB made its mark as a destruc-

tive disease of potato. Although the disease was subsequentlycontrolled relatively well through the use of fungicides and semi-resistant cultivars, LB re-emerged in 1970s as an important dis-ease of both potato and tomato. This has been in part due tothe occurrence of sexual reproduction in populations of P. in-festans and generation of new and more aggressive isolates ofthe pathogen, many of which exhibit resistance to known sys-temic fungicides. New isolates have also appeared that are morepathogenic to tomato than potato. The identification, character-ization and introgression of new LB resistance genes and QTLsin potato within the past two decades have significantly con-tributed to the effective control of the disease. Although manyof the single-gene resistance traits have been overcome by thepathogen, pyramiding of multiple resistance genes as well asidentification and incorporation of genes conferring resistanceagainst a broad spectrum of P. infestans isolates have increasedthe prospect for successful control of the disease via geneticmeans. In tomato, recent breeding efforts have focused on in-corporation of the known resistance genes, Ph-2 and Ph-3, andidentification, characterization and incorporation of newer resis-

tance genes, such as Ph-5. It is expected that pyramiding of suchgenes will contribute to development of cultivars with strongerand more durable resistance. Efforts in identification and in-trogression of QTLs conferring horizontal resistance in tomatohave encountered limited success due to problems associatedwith linkage drag. This has been mainly due to the presence ofQTLs in distantly related, green-fruited wild species of tomato,including S. habrochaites and S. peruvianum. Future efforts toidentify new and more desirable sources of resistance and newresistance genes and QTLs are imperative. Fine mapping of suchgenes or QTLs can facilitate their clean transfer to the cultigenvia MAS with little or no linkage drag problems.

B. Early BlightEarly blight is known as a destructive disease of tomato in

many production areas in the U.S. and elsewhere in the world.Unlike many other diseases of tomato, the genetic resistance toEB is quite complex. It does not follow the gene-for-gene in-teraction model. No major resistance gene has been identifiedin tomato and there is no pathogen race specificity. In addition,many factors such as plant physiological maturity, plant typeand growth habit, earliness in maturity and fruit load affect theplants’ response to the disease. Sources of genetic resistance toA. solani have been identified within the related wild speciesof tomato and have been utilized in traditional breeding pro-grams. Such efforts have resulted in lines or cultivars that ex-hibit moderate resistance to the disease or possess undesirablehorticultural characteristics such as late maturity or low-yieldability. The limited success of traditional breeding for improvedEB resistance has been due to many factors, including the com-plexity of the disease and polygenic nature of the resistance,complex interactions between resistance and other physiologi-cal and morphological characteristics, difficulties in maintain-ing resistance through successive generations of backcrossing,pathogen nonspecificity, lack of sufficient correlation betweenfield and greenhouse screenings, lack of ability to directly iden-tify and select for resistance genes, and confounding effectsof other defoliating diseases during phenotypic screening. Atpresent, disease control measures mainly include crop rotationand routine application of fungicides. However, recently newsources of EB resistance have been identified within the closely-related wild species of tomato S. pimpinellifolium, which havebeen utilized extensively for identifying resistance QTLs anddeveloping tomatoes with improved resistance. The preliminaryresults from these studies have been encouraging, suggestingthe possibility of developing commercially acceptable cultivarswith good level of EB resistance. However, it seems that de-velopment of high-yielding tomato cultivars with early maturityand high resistance to EB remains a challenging task.

ACKNOWLEDGMENTSWe would like to thank Liping Zhang, David O. Nino-Liu

and Arun Sharma, former members of the tomato genetics and

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BLIGHT RESISTANCE IN TOMATO 101

breeding program at Penn State, for their contributions to iden-tifying QTLs for early blight resistance. This review article con-tains information gathered from numerous published and unpub-lished resources, and thus we would like to extend our appreci-ations to all authors of the references used in this manuscript.

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