Next Article in Journal
Construction of Watermelon Mutant Library Based on 60Co γ-ray Irradiation and EMS Treatment for Germplasm Innovation
Next Article in Special Issue
Comparative Analysis Revealed Intrageneric and Intraspecific Genomic Variation in Chloroplast Genomes of Actinidia spp. (Actinidiaceae, Viridiplantae)
Previous Article in Journal
Exogenous Application of Coconut Water to Promote Growth and Increase the Yield, Bioactive Compounds, and Antioxidant Activity for Hericium erinaceus Cultivation
Previous Article in Special Issue
Comparative Transcriptome Analyses Reveal Different Regulatory Mechanisms in Ecological Adaptation between Chrysanthemum vestitum and Chrysanthemum mongolicum
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Horticulturae
Volume 9
Issue 10
10.3390/horticulturae9101132
horticulturae-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Recent Advances and Challenges in Management of Colletotrichum orbiculare, the Causal Agent of Watermelon Anthracnose

by
Takshay Patel
1,
Lina M. Quesada-Ocampo
2,
Todd C. Wehner
1,*,
Bed Prakash Bhatta
3,4,
Edgar Correa
5 and
Subas Malla
3,4,*
1
Department of Horticultural Science, North Carolina State University, Raleigh, NC 27695, USA
2
Department of Entomology and Plant Pathology, North Carolina State University, Raleigh, NC 27695, USA
3
Department of Horticultural Sciences, Texas A&M University, College Station, TX 77843, USA
4
Texas A&M AgriLife Research and Extension Center, Uvalde, TX 78801, USA
5
Department of Soil and Crop Sciences, Texas A&M University, College Station, TX 77843, USA
*
Authors to whom correspondence should be addressed.
Horticulturae 2023, 9(10), 1132; https://doi.org/10.3390/horticulturae9101132
Submission received: 22 September 2023 / Revised: 7 October 2023 / Accepted: 10 October 2023 / Published: 13 October 2023
(This article belongs to the Special Issue Horticultural Crops Genetics and Genomics)
Browse Figure
Review Reports Versions Notes

Abstract

:
The fungus Colletotrichum orbiculare causes watermelon anthracnose and is an important pathogen of watermelon in the United States, causing a significant impact on yield and quality of the produce. The application of fungicides as preventative and post-occurrence control measures is currently being deployed by growers. Further study of the genetic and molecular basis of anthracnose resistance will help in guiding future watermelon breeding strategies. Several conserved virulence factors (effectors) in C. orbiculare have been reported to interact with the host, at times impairing the host immune machinery. A single dominant gene conferring race 1 anthracnose resistance was reported independently on two watermelon germplasm. The recent advances in genomics, transcriptomics, proteomics, and metabolomics could facilitate a better understanding of the interaction between C. orbiculare effectors and host resistance genes in the already sequenced watermelon genome. In this review, we encompass and discuss (i) the history of watermelon anthracnose, taxonomy, morphology, and diversity in races of C. orbiculare; (ii) the epidemiology of the anthracnose disease and host resistance; (iii) the genetics behind the pathogenesis; and (iv) the current advances in breeding and molecular efforts to elucidate anthracnose resistance.

1. Introduction

Watermelon is an important crop grown worldwide with 102 Megatons produced in 2021 [1]. In the United States (U.S.), the crop had a total economic value of USD 748 million in 2022 [2]. Watermelons are grown in most U.S. states, but the majority of the production occurs in Florida, Georgia, California, and Texas because of higher temperatures and a longer growing season [3]. The introduction of seedless cultivars has increased the per capita consumption of watermelon by 37% since 1980 [3]. As the watermelon industry grew, challenges related to fruit quality, yield, and production methods emerged. As is the case with many crops, disease and pest management is a significant limitation to watermelon production. Diseases are a major priority for watermelon producers since new races of pathogens continue to break down host resistance and develop insensitivity to fungicides. The major diseases of watermelons are Fusarium wilt, anthracnose, gummy stem blight, powdery mildew, Phytophthora, downy mildew, bacterial fruit blotch, and viruses. While Fusarium wilt and Phytophthora are considered the most devastating diseases to watermelon when fields become infested, foliar diseases such as anthracnose, gummy stem blight, downy mildew, and powdery mildew affect the crop on a yearly basis, forcing growers to make significant investments in cultural practices and crop protection to manage these diseases [4].

2. History of Watermelon Anthracnose

Anthracnose has been a major problem in watermelon production worldwide at least since the 19th century, and it occurs wherever cucurbits are grown in a humid climate. Passerini in Italy first observed anthracnose on calabash/bottle gourd in 1867 [5]. In 1875, Passerini reported anthracnose on watermelon and cantaloupe, which is the first known scientific report of anthracnose on watermelon. More reports came from Europe during the late 19th century. In the U.S., Dr. Eckfeldt (Philadelphia) and Prof. A. B. Seymour (Wisconsin) noted anthracnose on gourds and watermelons, respectively, in 1885. In 1889, Galloway reported melon anthracnose in New Jersey, Virginia, and North Carolina. Substantial losses of cantaloupes, cucumbers, and watermelons due to anthracnose epidemics started in 1904 in Nebraska, Indiana, New Jersey, West Virginia, North Carolina, South Carolina, Wisconsin, and Ohio. Yield losses of up to 30% in watermelons [6] and 60% in cucurbits [7] have been reported to be caused by anthracnose. Anthracnose became a major plant disease during the late 19th century, and by the early 20th century, many U.S. states started focusing on anthracnose as an important watermelon pathogen [8].

3. The Pathogen

3.1. The Causal Agent: Colletotrichum Orbiculare

The genus Colletotrichum is economically and scientifically important because it contains many plant-pathogenic fungi infecting a wide range of crops; those crops include row crops, fruits, flowers, and vegetables. Almost every domesticated crop is a host of a species belonging to the genus Colletotrichum [9]. In many hosts, Colletotrichum spp. infect all the aerial parts of the plant, including stems, leaves, fruits, and flowers. Colletotrichum spp. are also significant postharvest pathogens since spores from foliar infections during field growth can cause an infection that progresses during transport or on the market shelves, resulting in complete loss of the crop [10]. When the plant pathogens were defined based on host specificity, Colletotrichum consisted of almost 700 species. Later, von Arx reclassified those species in 1957 to 11 taxa based on morphological traits [11,12]. Colletotrichum has been used as a model system for biochemical, physiological, and genetic studies. The concept of pathogen races was initially recognized in Colletotrichum lindemuthianum [13]. Many of the studies that laid the foundation for the concept of systemic acquired resistance were performed using the cucumber model of Colletotrichum orbiculare [14].
Colletotrichum orbiculare ((Berk. & Mont.) Arx) is an important pathogen of cucurbits including cucumber, muskmelon, watermelon, squash, gourd, pumpkin, cantaloupe, honeydew, and Luffa spp. [15]. C. orbiculare can also infect tobacco [16]. Anthracnose can cause severe damage, both in the field and postharvest, and is one of the top research priorities for watermelon in the U.S. [17]. In the field, C. orbiculare infects all the above-ground parts of plants, including leaves, stems, flowers, and fruits [3,5,8]. Infections on any of these plant parts have direct effects on yield. The pathogen also infects all growth stages of the plant, from the seedling stage to mature fruit-bearing plants. Defoliation will leave the plant with poor photosynthetic capacity, stunt fruit development, and expose the mature fruit to direct sunlight, leading to sunburn. Infection on both growing and mature plants leads to unmarketable produce.

3.2. Taxonomy

Colletotrichum orbiculare belongs to the kingdom Fungi, phylum Ascomycota, class Sordariomycetes, order Glomerellales, family Glomerellaceae, genus Colletotrichum [18,19].
Earlier when the taxonomic classification of plant pathogenic fungi was based on plant disease specificity, C. orbiculare was named and identified multiple times by different researchers around the world [12]. Cucurbit and bean anthracnose were assumed to be caused by the same fungus, which was named Colletotrichum lagenarium [8]. This assumption was discarded in a comparative study of anthracnose fungi, in which bean anthracnose was named Volutella citrulli. Based on modern molecular tests, C. orbiculare is recognized as a species complex, with C. lindemuthianum, C. malvarum, C. orbiculare, and C. trifolii as distinct species [20]. Currently, the isolates causing watermelon anthracnose are classified as a subspecies in the C. orbiculare species group. Researchers still differ in classifying this pathogen [18].

3.3. Fungal Morphology

The mycelium at first is colorless, thin-walled, septate, and uniformly cylindrical. Many of the cells later increase in diameter up to threefold, becoming thick-walled and dark brown in color [8,18]. On culture media, mycelium is first colorless and then pink and black at the end. Pink coloration is sometimes observed in host tissues, with blackening of mycelium being common in fruit lesions. Acervuli, anthracnose mycelium aggregates, branch and become intertwined to send out a layer of short colorless conidiophores [8]. From the tip of the conidiophores, spores bud off apically one at a time, piling up to form a pink slimy cluster on top of acervuli. Spores are surrounded by a sticky water-soluble matrix and are single-celled, clear, oblong or ovate, and vaguely pointed at one end [8]. Spore size varies from 13 to 19 µm by 4 to 6 µm, and masses are pink in color. Acervuli have two to three long setae scattered among the conidiophores, which are brown, thick-walled bristles 90 to 120 µm in length [8,18]. The number of setae in a single acervulus varies and can be up to 36. Spores form heaps as high as setae, with setae supporting the spore mass. Sclerotial bodies are typically observed more on media and fruit lesions and are formed due to the further development of the bases of stromata or acervuli, where the whole mass is enlarged and black in color [8]. On media, the spore mass may dry and remain as a part of sclerotial bodies, whereas in fruit lesions, the spores are washed away, and the black stroma that forms the black spots on fruits remains [8].
Germinating spores form an appressorium at the tip of each of the germ tubes, which are brown, melanized, thick-walled, and ovoid to spherical in shape. Appressoria are slightly tapered at one end and flattened on the side of contact with the host [8]. Melanization of appressoria is important for pathogenicity. Melanin-deficient mutants have reduced pathogenicity, and melanization is supposed to resist the high turgor pressure within the appressorium and direct the force on the leaf epidermis for penetration [21]. Melanization in fungal spores is also assumed to provide protection under adverse conditions like oxygen radicals, high temperatures, irradiation, or lysis by other microbes [22].

3.4. Life and Disease Cycle

Colletotrichum orbiculare is the asexual form of the cucurbit anthracnose pathogen and propagates through conidia. It normally exits in the asexual stage, and it rarely undergoes the sexual stage [18,23]. There has been no defined complete life cycle for C. orbiculare and only a few reports of the sexual stage of C. orbiculare. The sexual stage of C. orbiculare was reported as a species of Glomerella but was not classified [23]. Ascospores are produced in abundance when paired with other isolates of C. orbiculare, but few ascospores develop when isolates are selfed [23].
There is a close relation between C. orbiculare spread and wet weather conditions like rain, morning dew, and overhead irrigation. Conidia are mainly dispersed by rain splashing, but also by wind, instruments, and workers [4]. Spore heaps in acervuli are surrounded by a sticky water-soluble matrix [8,18], explaining the need for moisture for spread. The importance of moisture for C. orbiculare was observed and established in the early 20th century when anthracnose epidemics were new. A wetness period of 16 h or more shows maximum disease development [24]. Further, temperatures from 18 °C to 27 °C (65° to 80 °F) are ideal for the establishment and growth of C. orbiculare on watermelon [25]. C. orbiculare overwinters by surviving on the debris of infected plants. Cucurbit anthracnose was more severe on fields that had melons as previous crops [26]. Sheldon documented the spread of C. orbiculare by the transportation of diseased fruit and contaminated seeds [26]. Overall, C. orbiculare spreads by rain, irrigation, seeds, fruit, and overwintering and survives between seasons on infected plant debris, on volunteer plants, and in and on seeds from infected fruits [4].

3.5. Infection Process

Colletotrichum orbiculare is a hemibiotrophic fungus; during the initial stage of infection, it behaves as a biotrophic pathogen, keeping the host cells alive, and later it takes nutrients from dead host cells, switching to the necrotrophic stage [27]. C. orbiculare penetrates host leaves using two entry modes: turgor-mediated invasion (TMI) via melanized appressoria and hyphal tip-based entry (HTE). During TMI, C. orbiculare penetrates the adaxial epidermis [8]. After landing, the spores adhere to the plant surface and then germinate to produce germ tubes and further form melanized appressoria. The appressoria penetrate plant epidermal cells directly through the cuticle and cell wall but not from the stomata [28]. The epidermal cell wall below the appressorium swells, mostly due to cell-wall-degrading enzymes secreted by C. orbiculare [27,28]. After penetration, biotrophic intracellular hyphae develop inside the host cells, infecting via intracellular colonization at the cellular level. The intracellular hypha is surrounded by an intact host cell plasma membrane, growing within the plant cell lumen, i.e., between the plant plasma membrane and plant cell walls [29]. The infection then proceeds to the necrotrophic phase where the secondary necrotrophic hyphae arise from the intracellular hyphae, obtaining nutrients from dead host cells [27,28]. HTE works independently of the melanized appressoria and is a morphogenic response at wound sites [30]. The existence of these two invasive strategies implies a sensing system that induces the respective morphogenesis response on wound sites and intact leaf tissue for pathogenesis.
In watermelon fruits, the hyphae grow throughout the rind, and acervuli are formed after 4 to 5 days of infection. Conidiophores form conidia masses rupturing the rind epidermis. In resistant watermelon plants, the appressorium entry during foliar disease is the same as in a susceptible plant but the hyphae are only able to infect a few cells around the penetration site [28]. Plant cells around infected leaf sites elongate, divide, and form a raised compact mass to resist fungal growth [28], most likely through lignification. Fruits from resistant plants develop raised areas that are greener as compared to the surrounding rind and remain darker even when the remaining rind starts to bleach [28]. Like what is seen in leaves of resistant plants, C. orbiculare only infects one to two epidermal cells in the fruit rind after penetration [28].

3.6. Disease Symptoms

Colletotrichum orbiculare causes anthracnose in all cucurbits, and the symptoms on each of the species vary. All the above-ground parts of plants are susceptible to anthracnose. Photosynthetic cells are more susceptible than non-photosynthetic tissue [28]. Lesions gradually increase in size with abundant acervuli formation (acervuli are conidiophores producing mycelium aggregates). The descriptions of symptom were added later [4]. On watermelon leaves, anthracnose produces blackish-brown lesions (Figure 1A). Centers of older lesions on leaves fall out, giving it a ‘shot hole’ appearance. Petioles and stems show sunken and dark-color spindle-shaped lesions, which penetrate deeply and finally grid the stem (Figure 1D). Infected young fruits show aborted growth or are abnormal. Lesions on young fruit are small, black depressed spots. On mature fruits, lesions start as yellow translucent centered elevated pimples, which later turn into flat-topped, circular, water-soaked elevations (Figure 1B,C). Lesions on mature fruit further sink and show pink spore masses on a black or cream-colored background. The black lesions are the result of the black stroma left behind after the washing of spores, whereas the pink masses are like the spore masses found in culture media.

3.7. Pathogenesis Genes and Effectors

The average genome size of Colletotrichum species is 40 Mb, but C. orbiculare has a surprisingly large genome of 90 Mb [27]. C. orbiculare expresses a large arsenal of genes during the infection process, including 287 protease-encoding genes, 327 plant cell-wall-degrading enzymes, 700 small secreted proteins (SSPs), and many secondary metabolite backbone-forming proteins [27]. All of these are expressed at a higher level than in other species such as C. graminicola and C. higginsianum. SSPs and secondary metabolite synthesis genes are upregulated during the initial biotrophic stage of infection, whereas degrading enzymes are upregulated during the later necrotrophic stage of infection. The upregulation of SSP genes during early infection in C. orbiculare suggests their importance in maintaining biotrophy during infection. As C. orbiculare is a hemibiotroph, there is an orchestrated expression of the effector genes during the shift from biotrophy to necrotrophy [31].
Although through genomic and transcriptomic studies it is known that C. orbiculare expresses an arsenal of genes involved in pathogenesis, only a few effectors and pathogenic pathways are known yet. C. orbiculare produces several effectors, which include necrosis- and ethylene-inducing peptide1-like protein1 (NLP1) [32,33], DN3, necrosis-inducing secreted protein1 (NIS1) [34,35], MC69 [36], and suppression of immunity in Nicotiana benthamiana (SIB1, SIB2) [37]. NLP1 is expressed specifically in necrotrophic invasive hyphae [33], has cytotoxic activity, induces cell death, and triggers immune response [38]. The effector DN3 of C. orbiculare suppresses NLP1-triggered plant cell death [39], which was also previously observed in the C. higginsianum DN3 homolog [40]. NIS1 induces cell death and is expressed in bulbous biotrophic primary hyphae, but its activity reduces in necrotrophic hyphae. Homologs of NIS1 are present in C. higginsianum, C. graminicola, and Magnaporthe oryzae, suggesting it is a conserved sequence in Colletotrichum species. Although NIS1-knockout mutants are virulent on tobacco [35], the transgenic expression of NIS1 in Arabidopsis made the plant susceptible to C. orbiculare [41]. Intuitively, the expression of cell-death-inducing NIS1 during the biotrophic phase suggests it is a recognized avirulence (AVR) protein. Recently, it has been shown that NIS1 associates with the receptor-like kinases (RLKs) such as brassinosteroid insensitive 1 (BRI1)-associated kinase (BAK1) and receptor-like cytoplasmic kinases (RLCKs) such as botrytis-induced kinase1 (BIK1) in the host, inhibits pathogen-associated molecular pattern (PAMP)-induced reactive oxygen species (ROS) generation, and ultimately impairs pathogen-associated molecular pattern (PAMP)-triggered immunity (PTI) [42]. DN3 suppresses NIS1-triggered HR-like cell death [35]. MC69 is one of the characterized effectors in C. orbiculare. Although the MC69 mutants of C. orbiculare had normal colony morphology and conidiogenesis, they had reduced lesion development on cucumber and tobacco [36]. C. orbiculare expresses MC69 predominantly during the biotrophic phase of infection. A recent study has proposed two novel effectors of C. orbiculare (SIB1 and SIB2), where SIB1 was found to be expressed at both the primary and later stages of host invasion in tobacco and cucumber, respectively [37].
Few signaling infection-related morphogenesis and pathogenesis pathways have been identified in C. orbiculare. C. orbiculare has three cascades important for virulence: CMK1, MAF1, and RPK1. The CMK1 cascade is involved in conidial germination, infection, growth, and appressorium maturation. The MAF1 cascade is required for appressorium differentiation, whereas the RPK1 pathway is essential for vegetative growth and conidiation. The melanization of appressoria is important for the normal function of C. orbiculare [21], and three melanin biosynthesis enzyme genes, PKS1, SCD1, and THR1, and one regulatory gene, CMR1, have been characterized [43,44,45,46,47]. Mutants of melanin-related genes showed defects in the melanization of appressoria and penetration ability. Further, fatty acid oxidation of peroxisomes is also required for melanization and metabolic processes involved in turgor generation for penetration [41].
Plants have receptors that recognize PAMPs. An example of a PAMP is chitin, a major cell wall component in filamentous fungi. C. orbiculare has the SSD1 gene involved in cell wall integrity. Mutants of ssd1 were not virulent and induced host defense response along with papillae formation. The ssd1 mutants had increased induction of salicylic acid-induced protein kinase (SIPK) and wound-induced protein kinase (WIPK) activity as compared to the wild type. The effector secretion mechanism has also been studied. C. orbiculare shows a strong ring-like signal of the effectors around the primary biotrophic hyphae. The ring signal is present in the interfacial region between the host and C. orbiculare, and not inside C. orbiculare [31,33]. C. orbiculare continuously secretes effectors towards the interfacial region. Effectors are also secreted in a single-dot fashion at the bottom of the appressoria and at putative interfaces on the hyphal surface.

3.8. Pathogen Races

The first commercial anthracnose-resistant watermelon cultivars were released in 1949 and were widely used to manage the rampant anthracnose outbreaks at that time. Since then, C. orbiculare has emerged and overcome host resistance. In 1958, severe anthracnose symptoms on resistant watermelon cultivars were observed in North Carolina [48]. Pathogen isolates from the new symptoms were observed to be undistinguishable from earlier C. orbiculare isolates. Isolates were defined into races based on the differential host reactions on different cucurbit cultivars. Most isolates before 1954 were defined as race 1. Isolates found in 1954 and 1955 were defined as race 2 and were highly pathogenic on all cucurbit cultivars of that time. Some isolates were defined as race 3. The difference between race 1 and race 3 was pathogenicity on squash cultivars. The new isolates had no morphological and cultural differences and were considered as different races [48]. Race 4 [49] and races 5, 6, and 7 [50] were also identified later.
Overall, from 1954 to 1964, seven pathological races of C. orbiculare were identified based on differential host reactions. The seven races were based on virulence differences on different cucurbit species and cultivars. Race diversity was reevaluated, and the seven races of C. orbiculare were combined into three vegetative compatibility groups (VCGs) by using 92 isolates from the U.S. [15]. Based on vegetative compatibility (a phenomenon where fungi with certain genetic similarity can fuse together to form a single heterokaryon), 11 VCGs were formed among the 92 C. orbiculare isolates. Out of the 11 VCGs, only 3 were pathogenic on cucurbits, namely VCGs 1, 2, and 3. VCGs 1 and 3 showed virulence on similar hosts. The watermelon cultivar ‘Charleston Gray’ was resistant to both VCGs 1 and 3, but susceptible to VCG 2. Similar resistance was shown by cucumber cultivars ‘Poinsett 76’ and ‘Gy 14’. VCGs 1 and 3 were classified as race 1 virulence phenotype, and VCG 2 as race 2 virulence phenotype. A race 2B has also been somewhat characterized through vegetative compatibility and virulence [51]. Race 2B has been found on watermelon, bottle gourd, and muskmelon and belongs to VCG 2 [4].

4. Disease Management

Field management practices for anthracnose include planting disease-free seed material, deep plowing of crop residue immediately after harvest, crop rotation with non-cucurbit crops for a minimum of 1 year (2 to 3 years is optimum), avoiding usage of farm machinery among fields when the foliage is wet, fungicide applications with effective active ingredients such as quinone outside inhibitors (QoIs), and resistant plant cultivars [4]. To reduce fruit damage by anthracnose, growers are recommended to avoid mechanical injury to fruits, inspect for infected fruits during harvest and discard them, disinfect the fruit surface with chlorinated water, and refrigerate the fruit after harvest to prevent or delay anthracnose development postharvest [4].

4.1. Host Resistance

Host resistance screening was conducted either on cotyledons [15] or the two- to four-true-leaf stage [52,53,54,55,56,57]. The disease inoculum used for the seedling screening varied from 2.5 × 103 [57] to 5 × 105 [54] conidial spores/mL. The incubation time also varied from 3 to 14 days post-inoculation (DPI). A recent study reported the optimum inoculum concentration and DPI for screening two- to four-leaf-stage seedlings for race 2 anthracnose resistance [53]. The authors of this study found that 1 × 105 spores/mL and a single disease rating of percent leaf lesion at 9 DPI were optimal.
In 1937, Layton started breeding for anthracnose resistance and identified sources of resistance to develop commercial cultivars. Five cultivars from Africa with high resistance to anthracnose were identified, out of which three had edible fruit and desired horticultural traits [5]. The three cultivars were named Africa 8, 9, and 13, and were further used as parents. Homozygous anthracnose-resistant selections from Africa 8, 9, and 13 were crossed with commercial cultivars Iowa Belle and Iowa King and a few other cultivars [5]. The commercial Iowa cultivars were wilt-resistant, large-fruited, and crisp-fleshed. The first widely accepted anthracnose-resistant watermelon cultivars were ‘Congo’ (1949), ‘Fairfax’ (1953), and ‘Charleston Gray’ (1955), released by Andrus [57,58]. Charleston Gray, Congo, and Fairfax are resistant to races 1 and 3 but susceptible to race 2 [48]. Cultivars resistant to race 1 were also resistant to race 3 [57,59].
Resistance to race 2 was first found in a citron, W695, which was also resistant to races 1 and 3 [57]. PI 326515 was the first PI reported to have resistance only to race 2 [56]. More resistance sources to race 2 including PI 189225, 271775, 271778, and 512385 were identified [52,55]. Resistance to anthracnose race 2 was also identified in Citrullus colocynthis, designated as R309 [60]. Interestingly, two studies found that resistance in Citrullus colocynthis, R309, did not follow the single gene inheritance and was suggested to be multigenic [60]. These studies suggested that a dominant single gene confers major resistance, but there are other genes contributing to the phenotype. R309 has been the only source of multigenic resistance; no other multigenic resistance sources have been reported.
The first inheritance work on anthracnose resistance was performed in 1937 [5]. Resistance to race 1 is dominant over susceptibility and segregates as a single gene. Resistance to races 1 and 3 is controlled by the same gene, Ar-1 [57]. Inheritance of race 2 resistance is like race 1 resistance, dominant and segregating as a single gene [56]. In Korea, Jang et al. used a biparental population, ‘DrHS7250’ (female parent, resistant breeding line) and ‘Oto9491’ (male parent, susceptible breeding line), to identify a C. orbiculare race 1-resistance quantitative trait locus (QTL) on chromosome 8 and further conducted transcriptomics via RNAseq on the parents to identify a coiled-coil (CC)–nucleotide-binding site (NBS)–leucine-rich repeat (LRR) gene in the QTL region that conferred resistance to the disease [61]. They hypothesized that residue 18 of a conserved motif, IxxLPxSxxxLYNLQTLxL, could govern resistance in ‘DrHs7250’. An independent study conducted in the U.S. using a biparental mapping population, ‘Charleston Gray’ (female parent, resistant) and ‘New Hampshire Midget’ (male parent, susceptible), found a major C. orbiculare race 1-resistance QTL in the same region on chromosome 8 [62]. A PACE SNP marker designed from the SNP marker CL 14-27-9, identified earlier [61], was also the diagnostic marker for the QTL (LOD = 14.06) in the study. Even though the resistance source for the breeding line ‘DrHS7250’ was not reported, it seemed that both ‘DrHS7250’ and ‘Charleston Gray’ may have the same resistance gene for C. orbiculare race 1. Both studies in different watermelon resistance sources validated that the race 1 anthracnose resistance is governed by a single dominant gene. Even today, anthracnose is a problem and a major research priority in watermelon [17]. Most of the current commercial cultivars with anthracnose resistance were developed by private industry (Table 1). These commercial cultivars have intermediate to high levels of resistance to anthracnose race 1, and some descriptions do not specify the race. Many hybrid watermelon cultivars are resistant to races 1 and 2B and susceptible to race 2 [4]. The SNP marker CL 14-27-9 could be utilized as a diagnostic marker to develop race 1-resistant cultivars via marker-assisted selection in watermelon breeding programs.

4.2. Crop Protection

Growers often use fungicides to manage watermelon anthracnose throughout the growing season. Fungicides can be applied preventatively if cost-effective, or application should be started with the occurrence of the symptoms in a 5-to-10-day interval. If disease severity is high or environmental conditions are conducive to disease (wet weather), growers will use the shorter application interval. Effective fungicide active ingredients for managing watermelon anthracnose include compounds in group 11: trifloxystrobin, azoxystrobin, pyraclostrobin, fluoxastrobin; group 7: boscalid, fluxapyroxad; group 3: difenoconazole; group M05: chlorothalonil; and group M03: mancozeb [63,64]. Group 11 fungicides correspond to quinone outside inhibitors (QoIs), group 7 fungicides correspond to succinate dehydrogenase inhibitors (SDHIs), group 3 fungicides correspond to demethylation inhibitors (DMIs), and M05 and M03 have multi-site contact activity. Products commonly recommended for watermelon anthracnose control include ‘Kocide 3000’ (Copper Hydroxide), ‘Pristine’ (pyraclostrobin, boscalid), ‘Cabrio’ (pyraclostrobin), ‘Quadris Top’ (azoxystrobin, difenoconazole), ‘Bravo WeatherStik’ (chlorothalonil), and ‘TopGuard EQ’ (azoxystrobin) [63,65,66,67].

5. Prospects and Challenges

Although anthracnose has been an important watermelon disease for around a century, there are still many unanswered questions regarding pathogen biology and disease management. Anthracnose races have been identified for over 60 years now, but the genetic bases of those races remain unknown, forcing researchers to rely on differential phenotypic responses for race identification. In genomic studies like that of [27] the genome of C. orbiculare was sequenced, comparative studies were conducted, and transcriptomic analysis unraveling candidate effectors involved during different stages of infection was performed. Recently, the core effector NIS1 was shown to attack the conserved immune kinases in the host system and disrupt the first layer of plant immunity (PTI) [42]. We are yet to understand how plant resistance genes interact with the C. orbiculare effectors through the second layer of defense, i.e., effector-triggered immunity (ETI).
Many genomic resources have been developed for cucurbit crops and watermelon in particular [68,69,70]. Nonetheless, genetic determinants of anthracnose resistance have not been clearly identified. Cucurbit breeders still rely on genetic maps, loci, and QTLs to describe genetic resistance for conventional breeding, but specific resistance genes and the mechanism of anthracnose resistance have not been characterized. The whole genome sequence available for watermelon and advancements in genomics, transcriptomics, proteomics, and metabolomics offer the opportunity to identify genes responsible for the anthracnose-resistant phenotype; however, these approaches are sensitive to noise from phenotyping and genotyping and do not always result in a clear candidate gene. The occurrence of races in C. orbiculare is an indication that pyramiding resistance will be required to ensure the durability of the trait and minimize the risk of new isolates overcoming individual resistance genes as has occurred with other cucurbit diseases [48,71]. Establishing the resistance gene repertoire in watermelon and characterizing the interactions of such proteins with pathogen proteins that result in a resistant phenotype will be needed to achieve durable anthracnose resistance in watermelon. Likewise, continued identification of new and improved resistance sources will remain a priority for breeding anthracnose resistance.

Author Contributions

Conceptualization, T.P., T.C.W., L.M.Q.-O. and S.M.; writing—original draft preparation, T.P., B.P.B. and E.C.; writing—review and editing, T.C.W., L.M.Q.-O. and S.M.; supervision, T.C.W. and S.M.; funding acquisition, S.M. All authors have read and agreed to the published version of the manuscript.

Funding

This research was supported by USDA Hatch Project TEX09665, Texas A&M AgriLife Vegetable Seed Grant, Texas A&M University Excellence Fellowship, and Texas A&M AgriLife Research Strategic Initiative Assistantship.

Data Availability Statement

No new data were created or analyzed in this study. Data sharing is not applicable to this article.

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

References

  1. FAO. Agricultural Production Statistics 2000–2021; FAOSTAT Analytical Brief Series No. 60; FAO: Rome, Italy, 2022. [Google Scholar] [CrossRef]
  2. USDA-NASS. Vegetables 2022 Summary (February 2023). National Agricultural Statistics Service (United States Department of Agriculture). Available online: https://downloads.usda.library.cornell.edu/usda-esmis/files/02870v86p/hq37x121v/4b29ck28c/vegean23.pdf (accessed on 10 May 2023).
  3. Wehner, T.C. Watermelon. In Vegetables I: Asteraceae, Brassicaceae, Chenopodiaceae, and Cucurbitaceae; Prohens, J., Nuez, F., Eds.; Part of Book Series: Handbook of Plant Breeding; Springer: New York, NY, USA, 2008; pp. 381–418. [Google Scholar]
  4. Keinath, A.P. Anthracnose. In Compendium of Cucurbit Diseases and Pests, 2nd ed.; Keinath, A.P., Wintermantel, W.M., Zitter, T.A., Eds.; The American Phytopathological Society: St. Paul, MN, USA, 2017; pp. 54–55. [Google Scholar]
  5. Layton, D.V. The Parasitism of Colletotrichum lagenarium (Pass.) Ell. and Halst.; Research Bulletin 223; Agricultural Experiment Station, Iowa State College of Agriculture and Mechanic Arts: Ames, IA, USA, 1937; pp. 37–67. [Google Scholar]
  6. Parris, G. Watermelon Breeding. Econ. Bot. 1949, 3, 193–212. [Google Scholar] [CrossRef]
  7. Thompson, D.; Jenkins, S. Influence of cultivar resistance, initial disease, environment, and fungicide concentration and timing on anthracnose development and yield loss in pickling cucumbers. Phytopathology 1985, 75, 1422–1427. [Google Scholar] [CrossRef]
  8. Gardner, M.W. Anthracnose of Cucurbits; U.S. Department of Agriculture Bulletin; U.S. Department of Agriculture: Washington, DC, USA, 1918; pp. 1–68.
  9. Dean, R.; Van Kan, J.A.; Pretorius, Z.A.; Hammond-Kosack, K.E.; Di Pietro, A.; Spanu, P.D.; Rudd, J.J.; Dickman, M.; Kahmann, R.; Ellis, J. The Top 10 fungal pathogens in molecular plant pathology. Mol. Plant Pathol. 2012, 13, 414–430. [Google Scholar] [CrossRef]
  10. Prusky, D. Pathogen quiescence in postharvest diseases. Annu. Rev. Phytopathol. 1996, 34, 413–434. [Google Scholar] [CrossRef] [PubMed]
  11. von Arx, J.A. Die arten der gattung Colletotrichum Cda. Phytopathol. Z. 1957, 29, 413–468. [Google Scholar]
  12. von Arx, J.A. A revision of the fungi classified as Gloeosporium. Bibl. Mycol. 1970, 24, 1–203. [Google Scholar]
  13. Barrus, M.F. Variation of varieties of beans in their susceptibility to anthracnose. Phytopathology 1911, 1, 190–195. [Google Scholar]
  14. Durrant, W.E.; Dong, X. Systemic acquired resistance. Annu. Rev. Phytopathol. 2004, 42, 185–209. [Google Scholar] [CrossRef]
  15. Wasilwa, L.; Correll, J.; Morelock, T.; McNew, R. Reexamination of races of the cucurbit anthracnose pathogen Colletotrichum orbiculare. Phytopathology 1993, 83, 1190–1198. [Google Scholar] [CrossRef]
  16. Shen, S.; Goodwin, P.; Hsiang, T. Infection of Nicotiana species by the anthracnose fungus, Colletotrichum orbiculare. Eur. J. Plant Pathol. 2001, 107, 767–773. [Google Scholar] [CrossRef]
  17. Kousik, C.S.; Brusca, J.; Turechek, W.W. Diseases and disease management strategies take top research priority in the Watermelon Research and Development Group members survey (2014 to 2015). Plant Health Prog. 2016, 17, 53–58. [Google Scholar] [CrossRef]
  18. Damm, U.; Cannon, P.F.; Liu, F.; Barreto, R.W.; Guatimosim, E.; Crous, P.W. The Colletotrichum orbiculare species complex: Important pathogens of field crops and weeds. Fungal Divers. 2013, 61, 29–59. [Google Scholar] [CrossRef]
  19. Kirk, P.M. Catalogue of Life. Available online: https://www.catalogueoflife.org/data/taxon/X437 (accessed on 12 October 2023).
  20. Liu, B.; Wasilwa, L.; Morelock, T.; O’Neill, N.; Correll, J. Comparison of Colletotrichum orbiculare and several allied Colletotrichum spp. for mtDNA RFLPs, intron RFLP and sequence variation, vegetative compatibility, and host specificity. Phytopathology 2007, 97, 1305–1314. [Google Scholar] [CrossRef]
  21. Kubo, Y.; Furusawa, I. Melanin biosysnthesis: Prerequisite for successful invasion of the plant host by appressoria of Colletotrichum and Pyricularia. In The Fungal Spore and Disease Initiation in Plants and Animals; Cole, G.T., Hoch, H.C., Eds.; Springer: New York, NY, USA, 1991; pp. 205–218. [Google Scholar]
  22. Bell, A.A.; Wheeler, M.H. Biosynthesis and functions of fungal melanins. Annu. Rev. Phytopathol. 1986, 24, 411–451. [Google Scholar] [CrossRef]
  23. Jenkins, S.J.; Winstead, N. Observations on the sexual stage of Colletotrichum orbiculare. Science 1961, 133, 581–582. [Google Scholar] [CrossRef]
  24. Thompson, D.; Jenkins, S. Effect of temperature, moisture, and cucumber cultivar resistance on lesion size increase and conidial production by Colletotrichum lagenarium. Phytopathology 1985, 75, 828–832. [Google Scholar] [CrossRef]
  25. Monroe, J.; Santini, J.; Latin, R. A model defining the relationship between temperature and leaf wetness duration, and infection of watermelon by Colletotrichum orbiculare. Plant Dis. 1997, 81, 739–742. [Google Scholar] [CrossRef]
  26. Sheldon, J.L. Diseases of Melons and Cucumbers during 1903 and 1904; West Virginia Agricultural and Forestry Experiment Station Bulletin 94; West Virginia University Agricultural Experiment Station: Morgantown, WV, USA, 1904; pp. 119–138. [Google Scholar]
  27. Gan, P.; Ikeda, K.; Irieda, H.; Narusaka, M.; O’Connell, R.J.; Narusaka, Y.; Takano, Y.; Kubo, Y.; Shirasu, K. Comparative genomic and transcriptomic analyses reveal the hemibiotrophic stage shift of Colletotrichum fungi. New Phytol. 2013, 197, 1236–1249. [Google Scholar] [CrossRef]
  28. Anderson, J.; Walker, J. Histology of watermelon anthracnose. Phytopathology 1962, 52, 650–653. [Google Scholar]
  29. Perfect, S.E.; Hughes, H.B.; O’Connell, R.J.; Green, J.R. Colletotrichum: A model genus for studies on pathology and fungal–plant interactions. Fungal Genet. Biol. 1999, 27, 186–198. [Google Scholar] [CrossRef] [PubMed]
  30. Hiruma, K.; Onozawa-Komori, M.; Takahashi, F.; Asakura, M.; Bednarek, P.; Okuno, T.; Schulze-Lefert, P.; Takano, Y. Entry mode–dependent function of an indole glucosinolate pathway in Arabidopsis for nonhost resistance against anthracnose pathogens. Plant Cell 2010, 22, 2429–2443. [Google Scholar] [CrossRef] [PubMed]
  31. Irieda, H.; Takano, Y. Identification and characterization of virulence-related effectors in the cucumber anthracnose fungus Colletotrichum orbiculare. Physiol. Mol. Plant Pathol. 2016, 95, 87–92. [Google Scholar] [CrossRef]
  32. Gijzen, M.; Nürnberger, T. Nep1-like proteins from plant pathogens: Recruitment and diversification of the NPP1 domain across taxa. Phytochemistry 2006, 67, 1800–1807. [Google Scholar] [CrossRef] [PubMed]
  33. Irieda, H.; Maeda, H.; Akiyama, K.; Hagiwara, A.; Saitoh, H.; Uemura, A.; Terauchi, R.; Takano, Y. Colletotrichum orbiculare secretes virulence effectors to a biotrophic interface at the primary hyphal neck via exocytosis coupled with SEC22-mediated traffic. Plant Cell 2014, 26, 2265–2281. [Google Scholar] [CrossRef]
  34. Stephenson, S.-A.; Hatfield, J.; Rusu, A.G.; Maclean, D.J.; Manners, J.M. CgDN3: An essential pathogenicity gene of Colletotrichum gloeosporioides necessary to avert a hypersensitive-like response in the host Stylosanthes guianensis. Mol. Plant Microbe Interact. 2000, 13, 929–941. [Google Scholar] [CrossRef] [PubMed]
  35. Yoshino, K.; Irieda, H.; Sugimoto, F.; Yoshioka, H.; Okuno, T.; Takano, Y. Cell death of Nicotiana benthamiana is induced by secreted protein NIS1 of Colletotrichum orbiculare and is suppressed by a homologue of CgDN3. Mol. Plant Microbe Interact. 2012, 25, 625–636. [Google Scholar] [CrossRef]
  36. Saitoh, H.; Fujisawa, S.; Mitsuoka, C.; Ito, A.; Hirabuchi, A.; Ikeda, K.; Irieda, H.; Yoshino, K.; Yoshida, K.; Matsumura, H. Large-scale gene disruption in Magnaporthe oryzae identifies MC69, a secreted protein required for infection by monocot and dicot fungal pathogens. PLoS Pathog. 2012, 8, e1002711. [Google Scholar] [CrossRef]
  37. Zhang, R.; Isozumi, N.; Mori, M.; Okuta, R.; Singkaravanit-Ogawa, S.; Imamura, T.; Kurita, J.-I.; Gan, P.; Shirasu, K.; Ohki, S. Fungal effector SIB1 of Colletotrichum orbiculare has unique structural features and can suppress plant immunity in Nicotiana benthamiana. J. Biol. Chem. 2021, 297, 101370. [Google Scholar] [CrossRef]
  38. Azmi, N.S.A.; Singkaravanit-Ogawa, S.; Ikeda, K.; Kitakura, S.; Inoue, Y.; Narusaka, Y.; Shirasu, K.; Kaido, M.; Mise, K.; Takano, Y. Inappropriate expression of an NLP effector in Colletotrichum orbiculare impairs infection on cucurbitaceae cultivars via plant recognition of the C-terminal region. Mol. Plant Microbe Interact. 2018, 31, 101–111. [Google Scholar] [CrossRef]
  39. Isozumi, N.; Inoue, Y.; Imamura, T.; Mori, M.; Takano, Y.; Ohki, S. Ca2+-dependent interaction between calmodulin and CoDN3, an effector of Colletotrichum orbiculare. Biochem. Biophys. Res. Commun. 2019, 514, 803–808. [Google Scholar] [CrossRef]
  40. Kleemann, J.; Rincon-Rivera, L.J.; Takahara, H.; Neumann, U.; van Themaat, E.V.L.; van der Does, H.C.; Hacquard, S.; Stüber, K.; Will, I.; Schmalenbach, W. Sequential delivery of host-induced virulence effectors by appressoria and intracellular hyphae of the phytopathogen Colletotrichum higginsianum. PLoS Pathog. 2012, 8, e1002643. [Google Scholar] [CrossRef]
  41. Kubo, Y.; Takano, Y. Dynamics of infection-related morphogenesis and pathogenesis in Colletotrichum orbiculare. J. Gen. Plant Pathol. 2013, 79, 233–242. [Google Scholar] [CrossRef]
  42. Irieda, H.; Inoue, Y.; Mori, M.; Yamada, K.; Oshikawa, Y.; Saitoh, H.; Uemura, A.; Terauchi, R.; Kitakura, S.; Kosaka, A. Conserved fungal effector suppresses PAMP-triggered immunity by targeting plant immune kinases. Proc. Natl. Acad. Sci. USA 2019, 116, 496–505. [Google Scholar] [CrossRef]
  43. Kubo, Y.; Takano, Y.; Endo, N.; Yasuda, N.; Tajima, S.; Furusawa, I. Cloning and structural analysis of the melanin biosynthesis gene SCD1 encoding scytalone dehydratase in Colletotrichum lagenarium. Appl. Environ. Microbiol. 1996, 62, 4340–4344. [Google Scholar] [CrossRef] [PubMed]
  44. Perpetua, N.S.; Kubo, Y.; Yasuda, N.; Takano, Y.; Furusawa, I. Cloning and characterization of a melanin biosynthetic THR1 reductase gene essential for appressorial penetration of Colletotrichum lagenarium. Mol. Plant Microbe Interact. 1996, 9, 323–329. [Google Scholar] [CrossRef] [PubMed]
  45. Takano, Y.; Kubo, Y.; Shimizu, K.; Mise, K.; Okuno, T.; Furusawa, I. Structural analysis of PKS1, a polyketide synthase gene involved in melanin biosynthesis in Colletotrichum lagenarium. Mol. Gen. Genet. 1995, 249, 162–167. [Google Scholar] [CrossRef] [PubMed]
  46. Tsuji, G.; Kenmochi, Y.; Takano, Y.; Sweigard, J.; Farrall, L.; Furusawa, I.; Horino, O.; Kubo, Y. Novel fungal transcriptional activators, Cmr1p of Colletotrichum lagenarium and Pig1p of Magnaporthe grisea, contain Cys2His2 zinc finger and Zn (II) 2Cys6 binuclear cluster DNA-binding motifs and regulate transcription of melanin biosynthesis genes in a developmentally specific manner. Mol. Microbiol. 2000, 38, 940–954. [Google Scholar] [PubMed]
  47. Tsuji, G.; Sugahara, T.; Fujii, I.; Yuichiro, M.; Ebizuka, Y.; Shiraishi, T.; Yasuyuki, K. Evidence for involvement of two naphthol reductases in the first reduction step of melanin biosynthesis pathway of Colletotrichum lagenarium. Mycol. Res. 2003, 107, 854–860. [Google Scholar] [CrossRef]
  48. Goode, M.J. Physiological specialization in Colletotrichum lagenarium. Phytopathology 1958, 48, 79–83. [Google Scholar]
  49. Dutta, S.; Hall, C.; Heyne, E. Observations on the physiological races of Colletotrichum lagenarium. Bot. Gaz. 1960, 121, 163–166. [Google Scholar] [CrossRef]
  50. Jenkins, S.F.; Winstead, N.N.; McCombs, C.L. Pathogenic comparisons of three new and four previously described races of Glomerella cingulata var. orbiculare. Plant Dis. Rep. 1964, 48, 619–622. [Google Scholar]
  51. Wasilwa, L.; Correll, J.; Morelock, T. Further characterization of Colletotrichum orbiculare for vegetative compatibility and virulence. Phytopathology 1996, 86, S62. [Google Scholar]
  52. Boyhan, G.; Norton, J.; Abrahams, B.; Wen, H. A new source of resistance to anthracnose (Race 2) in watermelon. HortScience 1994, 29, 111–112. [Google Scholar] [CrossRef]
  53. Correa, E.; Crosby, K.; Malla, S. Optimizing a seedling screening method for anthracnose resistance in watermelon. Plant Health Prog. 2021, 22, 536–543. [Google Scholar] [CrossRef]
  54. Keinath, A.P. Identification of races of Colletotrichum orbiculare on muskmelon in South Carolina. Plant Health Prog. 2015, 16, 88–89. [Google Scholar] [CrossRef]
  55. Sowell, G., Jr.; Rhodes, B.; Norton, J. New sources of resistance to watermelon anthracnose [Colletotrichum langenarium]. J. Am. Soc. Hortic. Sci. 1980, 105, 197–199. [Google Scholar] [CrossRef]
  56. Suvanprakorn, K.; Norton, J. Inheritance of resistance to race 2 anthracnose [caused by Coletotrichum lagenarium] in watermelon. J. Am. Soc. Hortic. Sci. 1980, 106, 862–865. [Google Scholar] [CrossRef]
  57. Winstead, N.; Goode, M.; Barham, W. Resistance in watermelon to Colletotrichum lagenarium races 1, 2, and 3. Plant Dis. Rep. 1959, 43, 570–577. [Google Scholar]
  58. Andrus, C.F. New watermelon varieties: Bring new life to that industry. Seed World 1955, 4, 36–40. [Google Scholar]
  59. Jenkins, S.F.; Winstead, N. Glomerella magna, cause of a new anthracnose of cucurbits. Phytopathology 1964, 54, 452–454. [Google Scholar]
  60. Love, S.; Rhodes, B. Single gene control of anthracnose resistance in Citrullus? Cucurbit Genet. Coop. Rep. 1988, 11, 64–67. [Google Scholar]
  61. Jang, Y.J.; Seo, M.; Hersh, C.P.; Rhee, S.-J.; Kim, Y.; Lee, G.P. An evolutionarily conserved non-synonymous SNP in a leucine-rich repeat domain determines anthracnose resistance in watermelon. Theor. Appl. Genet. 2019, 132, 473–488. [Google Scholar] [CrossRef]
  62. Bhatta, B.P.; Patel, T.; Correa, E.; Wehner, T.C.; Crosby, K.M.; Thomson, M.J.; Metz, R.; Wang, S.; Brun, M.; Johnson, C.D.; et al. Dissection of race 1 anthracnose resistance in a watermelon (Citrullus lanatus var. lanatus) biparental mapping population. Euphytica 2022, 218, 157. [Google Scholar] [CrossRef]
  63. FRAC. FRAC Code List ©*2018: Fungicides Sorted by Mode of Action (Including FRAC Code Numbering); Fungicide Resistance Action Committee: Brussels, Belgium, 2018. [Google Scholar]
  64. Adams, M.L.; Noël, N.A.; Collins, H.; Quesada-Ocampo, L.M. Evaluation of fungicides for control of anthracnose on cucumber, Clinton 2017. Plant Dis. Manag. Rep. 2018, 12, V099. [Google Scholar]
  65. Egel, D.S.; Marchino, C. Evaluation of systemic fungicide timing for the control of anthracnose on watermelon, 2017. Plant Dis. Manag. Rep. 2018, 12, V049. [Google Scholar]
  66. Everts, K.L.; Korir, R.C. Evaluation of fungicides for management of foliar diseases on watermelon, 2016. Plant Dis. Manag. Rep. 2017, 11, V022. [Google Scholar]
  67. Everts, K.L.; Korir, R.C. Evaluation of fungicide programs for management of foliar diseases on watermelon, 2017. Plant Dis. Manag. Rep. 2018, 12, V039. [Google Scholar]
  68. Huang, S.; Li, R.; Zhang, Z.; Li, L.; Gu, X.; Fan, W.; Lucas, W.J.; Wang, X.; Xie, B.; Ni, P. The genome of the cucumber, Cucumis sativus L. Nat. Genet. 2009, 41, 1275–1281. [Google Scholar] [CrossRef]
  69. Garcia-Mas, J.; Benjak, A.; Sanseverino, W.; Bourgeois, M.; Mir, G.; González, V.M.; Hénaff, E.; Câmara, F.; Cozzuto, L.; Lowy, E. The genome of melon (Cucumis melo L.). Proc. Natl. Acad. Sci. USA 2012, 109, 11872–11877. [Google Scholar] [CrossRef]
  70. Guo, S.; Zhang, J.; Sun, H.; Salse, J.; Lucas, W.J.; Zhang, H.; Zheng, Y.; Mao, L.; Ren, Y.; Wang, Z. The draft genome of watermelon (Citrullus lanatus) and resequencing of 20 diverse accessions. Nat. Genet. 2013, 45, 51–58. [Google Scholar] [CrossRef] [PubMed]
  71. Holmes, G.J.; Ojiambo, P.S.; Hausbeck, M.K.; Quesada-Ocampo, L.; Keinath, A.P. Resurgence of cucurbit downy mildew in the United States: A watershed event for research and extension. Plant Dis. 2015, 99, 428–441. [Google Scholar] [CrossRef] [PubMed]
Horticulturae 09 01132 g001
Figure 1. Anthracnose symptoms on watermelon. (A) Leaf; (B,C) fruit; (D) stem; (E–G) foliage.
Figure 1. Anthracnose symptoms on watermelon. (A) Leaf; (B,C) fruit; (D) stem; (E–G) foliage.
Horticulturae 09 01132 g001
Table 1. Anthracnose-resistant cultivars of watermelon.
Table 1. Anthracnose-resistant cultivars of watermelon.
CultivarLevel of ResistanceRaceCompany
SSX8585High1Sakata, Yokohama, Japan
ValentinoHigh1Sakata, Yokohama, Japan
BelmontIntermediate1Sakata, Yokohama, Japan
Sweet TreasureIntermediate1Sakata, Yokohama, Japan
FascinationIntermediate1Syngenta, Basel, Switzerland
MelodyIntermediate1Syngenta, Basel, Switzerland
ExcursionIntermediate1Syngenta, Basel, Switzerland
CaptivationIntermediate1Syngenta, Basel, Switzerland
CooperstownHigh1Seminis, St. Louis, MO, USA
MajesticHigh? †Seminis, St. Louis, MO, USA
Road TripHigh?Seminis, St. Louis, MO, USA
Santa MatildeHigh1Seminis, St. Louis, MO, USA
HMX 1925Intermediate1HM Clause, Davis, CA, USA
Maistros F1High1HM Clause, Davis, CA, USA
AccompliceHigh1HM Clause, Davis, CA, USA
MillenniumHigh1HM Clause, Davis, CA, USA
† Resistance to anthracnose race was not specified in the varietal description by the company.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Patel, T.; Quesada-Ocampo, L.M.; Wehner, T.C.; Bhatta, B.P.; Correa, E.; Malla, S. Recent Advances and Challenges in Management of Colletotrichum orbiculare, the Causal Agent of Watermelon Anthracnose. Horticulturae 2023, 9, 1132. https://doi.org/10.3390/horticulturae9101132

AMA Style

Patel T, Quesada-Ocampo LM, Wehner TC, Bhatta BP, Correa E, Malla S. Recent Advances and Challenges in Management of Colletotrichum orbiculare, the Causal Agent of Watermelon Anthracnose. Horticulturae. 2023; 9(10):1132. https://doi.org/10.3390/horticulturae9101132

Chicago/Turabian Style

Patel, Takshay, Lina M. Quesada-Ocampo, Todd C. Wehner, Bed Prakash Bhatta, Edgar Correa, and Subas Malla. 2023. "Recent Advances and Challenges in Management of Colletotrichum orbiculare, the Causal Agent of Watermelon Anthracnose" Horticulturae 9, no. 10: 1132. https://doi.org/10.3390/horticulturae9101132

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop