Identificación de Hospederos Alternos de Spongospora subterranea f. sp. subetrranea en Colombia por Bioensayos

Ivon Magaly Arcila Aristizabal¹; Elena Paola González Jaimes²; Catalina María Zuluaga Amaya³ and José Miguel Cotes Torres⁴

¹ Agronomist. Master's student, Universidad Nacional de Colombia - Sede Medellín - Facultad de Ciencias Agrarias - Departamento de Ciencias Agronómicas - Grupo de Investigación de Mejoramiento y Producción de Especies Andinas y Tropicales. Calle 59A No 63-20, Medellín, Colombia. <imarcila@unal.edu.co>
² Assistant Professor. Politécnico Colombiano Jaime Isaza Cadavid - Facultad de Ciencias Agrarias - Laboratorio de Sanidad Vegetal. Carrera 48 No. 7-151, Medellín, Colombia. <epgonzalez@elpoli.edu.co>
³ Professional Professor. Politécnico Colombiano Jaime Isaza Cadavid - Facultad de Ciencias Agrarias - Laboratorio de Sanidad Vegetal. Carrera 48 No 7-151, Medellín, Colombia. <catazuluaga81@gmail.com>
⁴ Associate Professor. Universidad Nacional de Colombia - Sede Medellín- Facultad de Ciencias Agrarias - Departamento de Ciencias Agronómicas - Grupo de Investigación de Mejoramiento y Producción de Especies Andinas y Tropicales. Calle 59A No 63 - 20, Medellín, Colombia. <jmcotes@unal.edu.co>

Abstract. Potato powdery scab, caused by Spongospora subterranea f. sp. subetrranea, is a disease that limits worldwide potato crop production. Incidence of the disease has been increasing in Colombia, thereby affecting tubers production, so far effective control methods have yet to be developed. The aim of this research was to establish the host range plants for Spongospora subterranea f. sp. subterranea by artificial inoculations. Thus, 33 species were inoculated with 1×10⁶ sporosori.mL^-1 solution for 12 days. The plants were then planted in field, evaluations were performed at 15, 30, 60, 90 and 120 days after inoculation, 10 plants of each species were selected for every evaluation, the roots were stained and the pathogenic structures were identified by microscopy. Morphological examination enabled the identification of trap plants species, which presented zoosporangia only; Type I hosts, which present sporosori only; and Type II hosts, which presented both sporosori and zoosporangia. Some of these hosts belonged to the Alliaceae, Apiaceae, Brassicaceae, Poaceae, Polygonaceae and Solanaceae families, thus extending the species range that can be reported as hosts of Spongospora subterranea f. sp. subterranea in Colombia.

Key words: Powdery scab, sporosori, zoosporangium, Solanum tuberosum, trap plants.

Resumen. El agente causal de la sarna polvosa de la papa, Spongospora subterranea f. sp. subetrranea es una de las enfermedades más limitantes de la producción del cultivo a nivel mundial. En Colombia, viene incrementando su incidencia al punto de disminuir la producción, sin llegar a encontrar métodos de control efectivos. En este estudio, se buscó establecer un rango de hospederos del patógeno a partir de inoculaciones por bioensayos. Así, plantas de 33 especies se inocularon en solución a una concentración de 1x10⁶ quistosoros mL^-1 durante 12 días; posteriormente, las plantas fueron sembradas en campo, realizando evaluaciones a los 15, 30, 60, 90 y 120 días después de inoculadas. En cada evaluación se seleccionaron diez plantas de cada especie, con el fin de hacer tinción de raíces e identificar al microscopio estructuras asociadas al patógeno. A partir de la identificación morfológica fue posible clasificar las especies como plantas trampa quienes sólo presentaron zoosporangios, hospedero Tipo I con presencia sólo de quistosoros y hospederos Tipo II quienes presentaron estructuras de zoosporangios y quistosoros. Algunos de éstos hospederos pertenecen a las familias Alliaceae, Apiaceae, Brassicaceae, Poaceae, Polygonaceae, Solanaceae, ampliándose así el rango de especies que pueden ser reportadas como hospederas de Spongospora subterranea f. sp. subterranea en Colombia.

Palabras clave: Sarna polvosa, quistosoros, zoosporangios, Solanum tuberosum, plantas trampa.

Plasmodiophorids are an order of zoospores eukaryotic characterized by their cruciform cellular divisions, biflagellate zoospores and the formation of environmentally resistant spores. They have a complex extrusome called the Rohr and Stachel, whose function is to infect host cells (Braselton, 1995; Webster and Webber, 2007). Today, there are 10 genera and 35 recognized species of Plasmodiophorids, some of which are important phytopathogens, such as Spongospora subterranea f. sp. subetrranea (Wallr.) Lagerh (the causal organism of potato powdery scab) and Plasmodiophora brassicae Woronin (the causal organism of clubroot) (Bulman et al., 2001). Additionally, some Plasmodiophorids species are vectors for virus-causing diseases, such as the potato mop top virus PMTV (Jones and Harrison, 1969). The PMTV virus can remain several years on tubers used as seeds, decreasing by factors as low precipitation or non S. subetrranea f. sp. subetrranea infected fields or infected with no virulent sporosori. However, there is a very close relation among S. subetrranea f. sp. subetrranea and PMTV (Carnegie et al., 2012).

The S. subetrranea f. sp. subetrranea plasmodium is characterized by its resistant spores aggregation, which can remain in the soil for several years and survive environmental stress (Harrison et al., 1997). These structures comprise 3 layers, which are known as W1, W2, and W3 and which provide greater resistance and survival over time (Merz, 1997).

Zoospore release (primary zoospores) is stimulated by water and roots exudates (Harrison et al., 1997; Merz, 2008; Montero-Astúa and Rivera, 2005). Prior to infection, the spore encysts, releases its flagella and deposits its cellular contents being a mononuclear plasmodium the first post-infection structure. Following several divisions of this plasmodium, a zoosporangium is formed. At this stage, it may release a secondary zoospore that can infect other host cells or multiply until it forms resistance structures. It remains unclear whether S. subetrranea f. sp. subetrranea has a sexual phase; however, this sexual phase is assumed based on studies performed using P. brassicae. If the sexual phase in S. subetrranea f. sp. subetrranea is present, genetic variability would be expected within populations (Harrison et al., 1997; Jaramillo and Botero, 2007). A new Type III genetic group was discovered recently in Colombia (Osorio et al., 2012) which is different from the previously reported Type I and Type II groups (Bulman and Marshall, 1998; Qu and Christ, 2004; Merz, 2008); thus, a sexual phase may be assumed.

Until now, many studies have been focused on the control of potato powdery scab such as the use of antagonistic microorganisms by Hoyos et al. (2008), who proposed Trichoderma asperellum Samuels, Lieckf. & Nirenberg as a bio-controlling microorganism for potato powder scab. Nevertheless, Gilchrist et al. (2009) evaluating the effect of T. asperellum strains on the incidence and severity of powdery scab, did not report any significant differences between plants treated with the microorganism compared with untreated plants. Similarly, Restrepo et al. (2009) evaluating T. harzianum Rifai, Mycorrhizae, Pseudomonas fluorescents Migula 1895 and pine shavings, found a positive effects on the incidence and severity reduction of the disease with all treatments. Other studies have focused on the application of commercial chemical products such as fluazinam, mancozeb, dichlorophen-Na, dichlofluanid, or a mixture of fluazinam + mancozeb before planting (Falloon et al., 1996), azoxystrobin, propamocarb, difenoconazole, pentachloronitrobenzene (PCNB) and plant extracts of Lippia origanoides Kunth and Calotropis procera (Ait.), which shows that those application reduced the incidence of powdery scab in harvested tubers. The plant extracts were the most effective against disease incidence and severity (Bittara et al., 2009). Larking and Griffin (2007) reported 15 to 40% reduction in powdery scab following prior application of green manure produced from Brassicas. This reduction was attributed to the sulfur compounds such as glucosinolates in these species. At the global level, the management of this disease is framed by the search for plant material showing resistance to the pathogen. Nevertheless, various management methods are necessary to control powdery scab (Falloon, 2008).

Reports of S. subetrranea f. sp. subetrranea infecting non-solanaceous plants have suggested that these alternative hosts could provide a means for the pathogen to survive and multiply over various periods of time (Falloon, 2008). Thus, one of the methods for detecting S. subetrranea f. sp. subetrranea includes the inoculation of plants using bioassays observing pathogenic structures inside the roots. Using this method, Merz (1989) detected S. subetrranea f. sp. subetrranea and quantified soil sporosori using a 3-stage protocol, the first of which is a pre-inoculation period in which the sporosori are suspended in a nutrient solution over 3 days to allow zoospores release. This stage is follow by a trapping stage where plants are inoculated over 2 days to assure roots infection and then a period of disease development where the plants are again suspended in nutrient solution over 7 days. Using this method, early detection of the disease and also the identification of potato varieties resistant to S. subetrranea f. sp. subetrranea was possible with the eventual goal of developing genetic improvement programs (Merz et al., 2004). In studies of early detection of S. subetrranea f. sp. subetrranea, the trapping period has been modified from 1 day (Nakayama et al., 2007) to 14 days (Qu and Christ, 2006) of inoculation, followed by observation of S. subetrranea f. sp. subetrranea structures in the roots of the inoculated plants.

Jones and Harrison (1972) reported species from the Chenopodiaceae, Aizoaceae, Brassicaceae, Caryophyllaceae, Gramineae and Leguminosae families as hosts of S. subetrranea f. sp. subetrranea. These authors observed only the zoosporangial stage of S. subetrranea f. sp. subetrranea at 38 and 45 days after inoculation on Chenopodium album, Matricaria matricarioides, Solanum nigrum, Urtica dioica, Poa annua, Lapsana communis and Rumex acetosella with sporosori drawn from tubers in nutrient solution.

Andersen et al. (2002) evaluated 17 weeds associated with potato crops in Denmark and reported Artemisia vulgaris L., Chamomilla suaveolens (Pursh) Rydb., Chenopodium album L., Galium aparine L., Geranium pusitlum L., Matricaria inodora L., Polygonum avicular L., P. convolvulus, L., R. acetosella L., Solanum nigrum L., Sonchus arvensis L., Urtica urens L. and Viola tricolor L. as zoosporangial hosts of S. subetrranea f. sp. subetrranea. Qu and Christ (2006) evaluated common crops and weeds in the United States and found that tomato (Lycopersicon esculentum L.), yellow mustard (Brassica campestris L.) and oat (Avena sativa L.) showed sporosori whereas Amaranthus retroflexus L., Ambrosia artemisiifolia L. Raphanus sativus L. Brassica napus L. Chenopodium album L., Cyperus esculentus L., Trifolium pratense L., Phleum pratense L., Secale cereale L., Fagopyrum esculentum and Datura stramonium L. were zoosporangial hosts indicating that these species may act as trapping plants. Trapping plants prevent the completion of the pathogen's life cycle, thus reducing the inoculum in the soil. These plants could contribute to integrated management of powdery scab if sown before the potato crop. White (1954) reported that Datura stramonium L. grown in infected soils reduced the severity of powdery scab by 79% after one potato harvest. Murakami et al. (2000) reported that Raphanus sativus L. var. longipinnatus reduced inoculum of P. brassicae by 71% in pots that were inoculated after sowing a susceptible plant compared with a test pot (R. sativus not sown) indicating that this plant may be used as a trapping plants for the management of cabbage clubroot.

More recently, Nitzan et al. (2009) reported Solanum sarrachoides Sendtn., which is an annual weed of irrigated crops in the US, as a host of S. subetrranea f. sp. subetrranea which had the ability to form galls increasing the soils inoculum. Similarly, in New Zealand, Shah et al. (2010) showed S. sarrachoides and Solanum nigrum L. as hosts of S. subetrranea f. sp. subetrranea and Meloidogyne fallax. Karssen concluded that these species most likely allowed the proliferation of zoospores from zoosporangia and the development of sporosori, thereby assisting the survival of the pathogen over many years.

This study attempted to use inoculations in bioassays to identify host plants of S. subetrranea f. sp. subetrranea that may be used within an integrated management program for this disease.

MATERIALS AND METHODS

Localization and inoculated species. The study was performed in the Paysandú Agrarian Station at the Universidad Nacional de Colombia, which is located in the Santa Elena Township, Medellín City, and it is at an altitude of 2,500 masl with average annual temperature of 14 °C. 33 species were evaluated in total, of which 22 are cultivatable, 11 are weeds associated with potato planting and two are commercial potato varieties (Table 1).

Inoculation by bioassays. Using the Alzate et al. (2008) method, sporosori extracted from naturally infected soil were used, as the infested dry soil was passed through a series of mesh screens of 90 and 25 mm pore size. Samples were then counted using a Neubauer chamber and the concentration was adjusted to 1×10⁶ sporosori.mL^-1. Plants were propagated in peat, and inoculated when they showed two pairs of leaves. Plants remaining for 12 days in 150 mL of water in which the sporosori had been diluted. To guarantee sufficient development of the plants, after 8 days, Hoagland (Millner and Kitt, 1992) nutrient solution was added to return to the initial volume. The control plants (non-inoculated) were submerged in water with nutrient solution. As a control variable, two types of Solanum tuberosum spp. andigena were used. Once the inoculation period had passed, the plants were sown in 1 m2 plots. 50 plants of each species were inoculated and a similar number were used as controls.

Evaluations. Evaluations were performed at 15, 30, 60, 90 and 120 days post-inoculation depending on the life cycle of each species. 10 inoculated and 10 control plants per species were used for each evaluation. The roots of each plant were washed using tap water and then examined for the presence of galls. To observe pathogenic structures under a microscope, a portion of root was stained using 0.5% trypan blue as follows. The root was placed in a 10-mL Falcon tube and trypan blue was applied directly to the root until it was covered completely. For evaluation, five roots per sample were placed on a slide and observed using a Nikon™ E-200 light microscope at a total magnification of 100× and 400×.

Data Analysis. For data analysis, the time of the appearance of the first observed microscopic structures was used. The incidence of zoosporangia and sporosori was estimated for the evaluated plants using the following equation:

Where "n" is the number of plants showing zoosporangia and sporosori on their roots, and "X" is the total number of plants. A confidence interval of 95% was also calculated using the equation (Clopper and Pearson, 1934):

Where the critical distribution value F0.025 is calculated using the following degrees of freedom: V1=2(n-x+1), V2 =2x, . Analysis was performed using the R program environment for data analysis (The R Development Core Team, 2012) and the Binomial package (Dorai-Raj, 2009).

RESULTS AND DISCUSSION

Periods of detection of S. subetrranea f. sp. subetrranea zoosporangia and sporosori in host-plant roots. P. segetum and S. nigrum species that were classified as trapping plants showed zoosporangia at 15 and 60 days, respectively. C. sativum, Pennisetum sp., R. glaucus, S. quitoense, T. officinale, A. graveolens, D. carota, S. oleraceus and R. sativus, classified as host plants Type I and II, showed resistance structures of S. subetrranea f. sp. subetrranea at 15 days. These results show that S. subetrranea f. sp. subetrranea colonizes and produces resistance structures within 15 days, demonstrating the susceptibility of various harvest species and weeds in Antioquia. Merz et al. (2004) observed zoosporangia at 3-4 days and the formation of mature galls 4 months after potato plants inoculation. In C. sativus, P. vulgaris, R. crispus, C. betacea, A. cepa, P. crispum, P. sativum, B. oleraceae, Z. mays, P. nepalense, S. lycopersicum and P. peruvianum, the infection was presented as a late infection and was observed after 30 days of inoculation. These results confirm the broad range of hosts for this Plasmodiophoromycete as reported previously by Jones and Harrison (1969; 1972) and Andersen et al. (2002) Table 2. S. subetrranea f. sp. subetrranea was not observed in non-inoculated plants, confirming the validity of the bioassays used.

Besides presenting with both zoosporangia and sporosori, D. carota and S. tuberosum (Puracé and Diacol Capiro varieties) also showed galls at 90 and 120 days (Figure 1), symptom that was not observed among the other studied species. The absence of gall formation in the other plants, despite the bioassay inoculation, has previously been reported by Qu and Christ (2006) and Van de Graaf et al. (2007), where many species of plant were inoculated with different sporosori concentrations, including Estima potato variety. These plants became infected but did not develop galls in their roots.

Pathogenic structures were not observed in A. sativum, H. radicata, T. repens, B. napus, B. campestris and B. vulgaris, making them candidates for rotation within a disease management strategy because they will not increase the soil inoculum. Nevertheless, Harrison and Jones (1970) and Van de Graaf et al. (2007) have reported B. vulgaris as a host of S. subetrranea f. sp. subetrranea. A difference in the varieties studied should have led the differences between these studies and the present results.

Some of the plants evaluated in this study have been studied in other countries. For example, in Pakistan, Iftikhar and Ahmad (2005) attempted to find alternate hosts for S. subetrranea f. sp. subetrranea, inoculated cultivated plants in nutrient solution, they reported that in Z. mays, P. sativum, D. stramonium, R. sativus and B. oleracea, the zoosporangial stage was only observed. However, in our study sporosori in these species were identified. In Colombia, Zuluaga et al. (2010) also reported Z. mays as forming pathogenic structures associated with S. subetrranea f. sp. subetrranea. However, Qu and Christ (2006), did not include Z. mays as a host but did include B. napus, which is a species that did not show pathogen-related structures in our study.

Betancur et al. (2011), studying naturally infested lots in Antioquia, reported that P. nepalense, P. clandestinum, R. crispus and Z. mays showed S. subetrranea f. sp. subetrranea-associated structures, corroborating the results of our study. These species belong to the Asteraceae, Polygonaceae and Poaceae families, which had already been reported as host families for S. subetrranea f. sp. subetrranea by Jones and Harrison (1969) and Harrison and Jones (1970). Some species from the Poaceae family, such as P. clandestinum (Betancur et al., 2011) and Z. mays (Iftikhar and Ahmad 2005; Zuluaga et al., 2010), have been reported as hosts for S. subetrranea f. sp. subetrranea and are also hosts for the Polymyxa genus, indicating a greater affinity of this group of Plasmodia for these species.

A. cepa, A. graveolens, C. sativum, C. sativus, P. crispum, P. vulgaris, P. peruviana, R. glaucus, S. quitoense and T. officinale have not been reported previously as hosts for S. subetrranea f. sp. subetrranea. Thus, this study is the first report of these species as hosts of. S. subetrranea. These species belong to the Alliaceae, Rosaceae and Cucurbitaceae families, which have not neither been previously reported hosting S. subetrranea f. sp. subetrranea.

With regard to the structures associated with S. subetrranea f. sp. subetrranea found in roots, the structures were morphologically different depending upon the species. These differences may be due to the affinity of the pathogen to each plant or to a reaction to the pathogen-host interaction. Indeed, these causes may underlie the time to detection of structures and zoosporangia in Z. mays, P. peruviana and S. tuberosum where cytoplasmic fragmentation showed a greater separation in root cells of S. tuberosum than in other species (Figure 2A, B, C). Sporosori were grooved, irregular and hollow with a spongy appearance due to the cavities in its exterior surface and the network of internal canals. They may be round, as observed in this study (Figure 2D, E, F), or oval, elongated or completely irregular as reported by Montero-Astúa and Rivera (2005). This diversity in sporosori shape and size was noted by Jaramillo et al. (2008) who reported differences in the size of sporosori from root galls compared with those from tuber pustules. Thus, differently shaped and sized structures may also occur in different host species.

Falloon et al. (2011), using light microscopy evaluated both diameter and zoospore number on each sporosori coming from 37 different inoculum country sources, founding that the average of zoospore diameter have a slight difference between the inoculum from Agraria (4 mm) and Estima (4.3 mm) varieties. However, the zoospore mean was 667 per sporosori varing from 155 to 1.526. The results of this study confirm that sporosori are a zoospore aggregation complex which are sponge-like with tunnel on the surface and the sporosori size is very different among the pathogen isolates from different countries and potato varieties. The result suggests that environmental conditions (more than host plant) affect the sporosori size of S subterranea f. sp. subetrranean.

Incidence of S. subetrranea f. sp. subetrranea sporosori and zoosporangia in host plant roots. Table 3 showed that significant differences were found (P<0.05) in the incidence of zoosporangia and sporosori among A. graveolens, C. sativus, D. carota, P. vulgaris, P. sativum, R. glaucus, R. crispus and C. betacea where incidences of 0.3000, 0.0286, 0.0600, 0.0400, 0.0600, 0.0286, 0.0313 and 0.0303, respectively, were also observed. D. stramonium, Pennisetum sp., P. vulgaris, R. glaucus, R. crispus and C. betacea showed incidences of 0.3478, 0.0208, 0.0400, 0.0286, 0.0313 and 0.0303, respectively.

A. cepa, A. graveolens, D. stramonium, S. lycopersicum, P. clandestinum, P. peruvianum, R. sativus, S. quitoense, S. oleraceus and Z. mays showed the highest incidence rates (Table 3) when using a high incidence-threshold (>10%) for plants infected with S. subetrranea f. sp. subetrranea.

Using this inoculation methodology, the incidence of S. subetrranea f. sp. subetrranea varies among species, reflecting the variability in resistance and susceptibility to infection as described by Mäkäräinen et al. (1994), who evaluated the disease incidence in wild and harvested solanaceous species inoculating with infected soil and zoospores in suspension. Among the species S. acaule, S. brevidens, S. tuberosum and S. sucrense, S. acaule was considered to be the most resistant because it showed a lower infection incidence in roots of tested plants.

Thus, the use of some species in rotation with potato crops, for example P. clandestinum (pennisetum grass), which is the primary rotational crop used in potato producing regions in Colombia, may increase the levels of S. subetrranea f. sp. subetrranea inoculum in the field because, even though there is no galls formation it is able to produce sporosori inside roots, enabling the pathogen to remain in the soil until later planting of an additional potato crop. Similarly, the weeds R. crispus and P. nepalense, always found in potato crops, repeatedly increase inoculum and improve the process of reinfection. In contrast, A. sativum and B. vulgaris did not show pathogenic structures, and these species could be used to rotate with potatoes within a program of integrated disease management with the goal of reducing silent multiplication of the pathogen.

When compared with Mäkäräinen et al. (1994) results, where inoculum from soil stored for 1 year led to a lower number of plants infected with S. subetrranea f. sp. subetrranea compared with a fresh inoculum, and Fornier et al. (1996), where exudates from different potato breeds stimulated the release of zoospores, it may be assumed that the survival of this pathogen over time is due to host plants which allow the repeated formation of resistance structures but not the persistence per se of the pathogen in the soil. Thus, the microorganism can infect different species and evade the plants mechanisms defense suggesting that species where resistance structures were found are a significant source of inoculum. This is especially true in the case of those plants associated with potato crops in Colombia, such as weeds or rotational crops. These crops may allow the maintenance, or even the slow increase of inoculum in soil while admit of pathogen resistance structures to remain viable and multiply for many years, thereby reinfecting new potato crops as reported by Harrison et al. (1997).

Following the proposal of Qu and Chist (2006), who classified as trapping plants, those plants inoculated with S. subetrranea f. sp. subetrranea showing only zoosporangial structures, i.e., plants that impeded follow up the life cycle of the pathogen and to form the resistance structures, thus in the current study P. segetum and S. nigrum may be classified as trapping plants. S. subetrranea f. sp. subetrranea may avoid plant basal resistance, enabling the entry of zoospore cellular material; however, it is possible that some resistance genes from S. nigrum and P. segetum trigger a hypersensitive response (Hammond-Kosack and Jones, 2000) impeding the sequel of the life cycle. With respect to the non-hosts, it is possible that these species show a basal resistance, cell wall rigidity and produced phytoalexins that control the infection and/or invasion of unadapted pathogens (Lipka et al., 2008). In plants reported as Type I and II hosts, it is possible that S. subetrranea f. sp. subetrranea evades host defense mechanisms and alters its metabolism thereby surviving and multiplying (Hammond-Kosack and Jones, 2000); however, more studies are required to understand this host-pathogen interactions.

Hernández et al. (2012) evaluated the infection intensity of S. subetrranea f. sp. subteranea by conventional (bioassays) and molecular (qPCR) methods in two potato varieties (Iwa and Agria) resistant and susceptible to the pathogen, respectively. The evaluated parameters were zoosporangia intensity in roots, galls number and S subterranea f. sp. subetrranea DNA quantity on roots by qPCR. It was found that inoculation did not affect the growing of the plants, but it reduces on 38% root dry matter of the susceptible variety. The intensity of zoosporangia development was similar in both varieties, galls was higher on the susceptible one and Spongospora subterranea f. sp. subetrranea DNA root quantity detected was twice larger on susceptible than on resistant variety. Authors suggest that zoosporangia stage on host roots is directly affected by host resistance factors more than sporosori stage, even on galls and tubers.

CONCLUSIONS

Bioassay inoculations allowed the identification of S. subetrranea f. sp. subetrranea associated structures, such as zoosporangia and sporosori, after 15 days in P. segetum, T. officinale, A. graveolens, D. carota, P. clandestinum, D. stramonium, C. sativum, Pennisetum sp., R. glaucus, S. quitoense, S. oleraceae, A. graveolens, D. carota and R. sativus. These data demonstrate the pathogen ability to induce early signs of infection and also the adaptation of this pathogen to different species, which may be a survival mechanism in the environment, thus confirming its broad hosts range.

Using bioassay inoculations, 3 types of hosts are reported based on pathogenic structures found in roots: 1) Trapping plants showed zoosporangia (P. segetum and Solanum sp.), 2) Type I hosts showed sporosori but not zoosporangia (C. betacea, S. quitoense, R. crispus, C. sativum, P. vulgaris, Pennisetum sp., C. sativus and R. glaucus) and 3) Type II hosts showed both structures (P. peruviana, P. crispum, D. carota, P. clandestinum, Z. mays, A. cepa, R. sativus, S. lycopersicum, P. sativum, P. nepalense, S. oleraceus, A. graveolens, B. oleraceae, T. officinale and D. stramonium). These results demonstrate that the pathogen can multiply and form resistance structures in Type I and II hosts. It does not have this multiplying ability in trapping plants, suggesting that these plants may help to reduce S. subetrranea f. sp. subetrranea inoculum in soil.

ACKNOWLEDGMENT

This research was supported by Ministerio de Agricultura y Desarrollo Rural (MADR), Asociación Hortifrutícola de Colombia (ASOHOFRUCOL), Politécnico Colombiano Jaime Isaza Cadavid, Universidad Nacional de Colombia Sede Medellín and Federación de Productores de Papa de Colombia (FEDEPAPA) into the project Uso de plantas trampa para la reducción de inóculo de Spongospora subterranea. The autors wish to thank members of research groups Sistemas Agrícolas Tropicales (COL0007149), and Mejoramiento y Producción de Especies Andinas y Tropicales (COL0039484) for their contribution to develop this research.

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