Isolation, identification, and pathogenicity of Steinernema carpocapsae and its bacterial symbiont in Cauca-Colombia

Publications

Share / Export Citation / Email / Print / Text size:

Journal of Nematology

Society of Nematologists

Subject: Life Sciences

GET ALERTS DONATE

ISSN: 0022-300X
eISSN: 2640-396X

DESCRIPTION

43
Reader(s)
139
Visit(s)
0
Comment(s)
0
Share(s)

SEARCH WITHIN CONTENT

FIND ARTICLE

Volume / Issue / page

Related articles

Isolation, identification, and pathogenicity of Steinernema carpocapsae and its bacterial symbiont in Cauca-Colombia

Esteban Neira-Monsalve / Natalia Carolina Wilches-Ramírez / Wilson Terán / María del Pilar Márquez / Ana Teresa Mosquera-Espinosa / Adriana Sáenz-Aponte *

Keywords : Biological control, Colombia, Epitrix cucumeris , Pandeleteius cinereus , Steinernema carpocapsae , Xenorhabus nematophila

Citation Information : Journal of Nematology. Volume 52, Pages 1-16, DOI: https://doi.org/10.21307/jofnem-2020-089

License : (CC-BY-4.0)

Received Date : 13-March-2020 / Published Online: 31-August-2020

ARTICLE

ABSTRACT

In Colombia, identification of entomopathogenic nematodes (EPN’s) native species is of great importance for pest management programs. The aim of this study was to isolate and identify EPNs and their bacterial symbiont in the department of Cauca-Colombia and then evaluate the susceptibility of two Hass avocado (Persea americana) pests to the EPNs isolated. EPNs were isolated from soil samples by the insect baiting technique. Their bacterial symbiont was isolated from hemolymph of infected Galleria mellonella larvae. Both organisms were molecularly identified. Morphological, and biochemical characterization was done for the bacteria. Susceptibility of Epitrix cucumeris and Pandeleteius cinereus adults was evaluated by individually exposing adults to 50 infective juveniles. EPNs were allegedly detected at two sampled sites (natural forest and coffee cultivation) in 5.8% of the samples analyzed. However, only natural forest EPN’s could be isolated and multiplied. The isolate was identified as Steinernema carpocapsae BPS and its bacterial symbiont as Xenorhabus nematophila BPS. Adults of both pests were susceptible to S. carpocapsae indicating this EPN potential for its management. The results of this study constitute the first record of S. carpocapsae in Colombia and the susceptibility of P. cinereus to this EPN.

Graphical ABSTRACT

Entomopathogenic nematodes (EPNs) of the Steinernematidae and Heterorhabditidae families are widely used as biological control agents that represent a promising alternative to replace pesticides (Labaude and Griffin, 2018), because of their ability to parasitize insects, being able to identify, locate, and infect a host and to kill it within 48 hr, as well as they are safe to vertebrates, plants, and other non-target organisms. Moreover, they can be applied by means of standard spraying equipment (Poinar, 1972; Hazir et al., 2003; Van Zyl and Malan, 2014), and can be propagated in mass either in vivo using host insects such as Galleria mellonella (Lepidoptera: Pyralidae) or in vitro in bioreactors using artificial culture media (Shapiro-Ilan et al., 2014).

EPNs life cycle has six states: egg, four juvenile stages, and adults: males, females, or hermaphrodites (Shapiro-Ilan et al., 2014). EPNs of the Steinernematidae family are characterized by symbiotic associations with bacteria of the genus Xenorhabdus, while EPNs of the Heterorhabditidae family have symbiotic associations with bacteria of the genus Photorhabdus. In these relationships, the nematode acts as a vector allowing the bacteria to reach and enter the host insect where they produce metabolites that kill it within 24 to 48 hr, degrade it and transform it into an optimal environment for the development of EPNs. Almost every stage of EPNs development takes place inside the insect. The exception is the infective juvenile (IJ) which is a modification of the J3 stage that does not feed or develop, responsible for actively mobilizing in the soil in search of potential hosts and for carrying the bacterial symbionts in the intestinal lumen (Heterorhabditis sp.), or in a specialized structure called the receptacle; a modification of the two most anterior intestinal cells (Steinernema sp.) (Griffin et al., 2005; Stock, 2015).

Because various environmental conditions can affect survival, reproductive potential, and virulence of EPNs, the use of native species for pest control is of great importance since they are better adapted to local environmental conditions than are foreign species (Campos-Herrera and Gutiérrez, 2009). Nevertheless, in a number of countries information on EPNs and their bacterial symbionts is scarce (De Brida et al., 2017). Colombia’s records of native EPNs, a tropical country with a large agricultural population and sector, do include Steinernema websteri, S. kraussei, S. colombiense, and Heterorhabditis bacteriophora (López-Núñez et al., 2007, 2008; Melo et al., 2009). The municipality of Toribío, in the Department of Cauca, Colombia, is a place where EPNs had never been registered before. It is characterized by small-scale family and community agriculture with varied subsistence and pasture crops. The aim of this study was to isolate and identify EPNs and their bacterial symbiont from crops and natural habitats at this municipality for later in vitro evaluation of susceptibility of two insect pests of economic interest for Hass avocado crops (Persea americana), the potato flea beetle (Epitrix cucumeris [Coleoptera: Chrysomelidae]), and a broad-nosed weevil (Pandeleteius cinereus [Coleoptera: Curculionidae]), to the EPNs isolated and the reference isolate Steinernema carpocapsae FA2015.

Materials and methods

Isolation and identification of entomopathogenic nematodes

Soil sampling

Soil samples were taken in the Toribío and Tacueyó reserves of the municipality of Toribío, in the Department of Cauca, Colombia, where EPNs have never been applied. These rural areas are located between 1,757 and 2,963 meters above sea level (m.a.s.l) where temperatures range between 13.9° and 20°C. Following the sampling method developed by Varón de Agudelo and Castillo (2001), 10 sampling sites in eight types of vegetation cover were selected: natural grassland (NG), pasture bordering annual crops (PBAC), natural forest (NF), first-growth American bamboo (Guadua sp.) (FGAB), coffee cultivation (Coffea arabica) (CC), horticultural cultivation (HC), strawberry cultivation (Fragaria sp.) (SC), and combined lulo (Solanum quitoense) and papaya cultivation (Carica sp.) (CLPC). At each site, two composite soil samples (10 subsamples, ~1 kg) were taken at depths between 20 and 30 cm, after removing soil cover to avoid cross contamination. Each independent composite sample was taken in an area measuring approximately 16 m2 with a zigzag distribution of 10 subsampling points. Samples were packed in plastic bags, labeled, and stored in a cooler (~15°C) (Orozco et al., 2014) in dark conditions for less than 24 hr until they arrived at the Biological Control laboratory of Pontificia Universidad Javeriana in Bogotá, Colombia to be immediately processed.

Isolation of EPNs

Composite soil samples taken from each site were mixed and stored at room temperature for 24 hr prior to testing for IJs using the insect baiting technique (Bedding and Akhurst, 1975). For this, six samples of 150 g of mixed soil were deposited in 200 cm3 plastic boxes. Then, five G. mellonella larvae and five Tenebrio molitor (Colepotera: Tenebrionidae) larvae were added per box at the soil surface. The boxes were labeled, inverted, and kept in darkness at 20°C. Four days later, the boxes were evaluated for dead larvae killed by EPNs. Sagging bodies and color changes were noted. Subsequently, dead larvae were placed individually in White traps (White, 1927) to obtain IJs. The IJs recovered were then multiplicated in G. mellonella larvae and maintained for later identification. To corroborate whether EPNs were absent or present in soil samples, samples were processed twice by repeating the process described above.

The IJs isolated were stored in polyurethane foams at 10°C in the Biological Control laboratory and deposited in the Entomology collection of the Javeriana Museum of Natural History of the Pontificia Universidad Javeriana in Bogotá, Colombia.

Abundance of EPNs at sampled sites (positive sites for EPNs/total sites) and the recovery frequency of EPNs (positive samples for EPNs/total samples analyzed) were determined according to the type of vegetation cover (Liu and Berry, 1995) and expressed as a percentage.

Molecular identification

DNA extraction

Genomic DNA was extracted individually from females following the protocol described by Çimen et al. (2016). After lysis of the nematodes, protein content was separated using NaCl at a final concentration of 1.7 M and centrifuging at 3,000 g for 15 min at room temperature. Finally, the supernatant was transferred to another tube for alcohol precipitation of DNA.

Amplification and sequencing of taxonomic markers

Sequences of the following three taxonomic markers commonly used for nematodes were amplified by PCR and used for molecular identification: a fragment of the 18S rRNA sequence using primers SSU18A-4F: 5′-GCTTGTCTCAAAGATTAAGCCATGCATG-3′ and SSU26Rplus4: 5′-AAGACATTCTTGGCAAATGCTTTCG-3′ (Morise et al., 2012); a fragment that contained the sequences ITS1, 5.8S, and ITS2 using primers 18S: 5′-TTGATTACGTCCCTGCCCTTT-3′ and 26S: 5′-TTTCACTCGCCGTTACTAAGG-3′ (Vrain et al., 1992); and a rRNA 28S fragment that contained the D2/D3 expansion sequence using D2F: 5′-CCTTAGTAACGGCGAGTGAAA-3′ (Nguyen et al., 2006) and 536: 5′-CAGCTATCCTGAGGAAAC-3′ as primers (Stock et al., 2001).

For fragment amplification, 50 µl of PCR mixture was prepared using 1X NH4 Buffer (Bioline, England), 3 mM MgCl2, 0.2 mM dNTPs, and 0.5 µg/µl Bovine Serum Albumin (New England Biolabs, United States), 0.5 µM forward primer, 0.5 µM reverse primer, Taq DNA polymerase 2U (Bioline, England), and 100 to 200 ng of template DNA (Hominick et al., 1997; Stock et al., 2001).

The PCR protocol for the 18S fragment consisted of one initial denaturation cycle at 94°C for 2 min followed by 35 cycles at 94°C for 10 sec, 55°C for 30 sec, 68°C for 1 min (Morise et al., 2012), and a final extension at 68°C for 7 min. For the ITS and D2/D3 fragments, the PCR protocol consisted of one cycle of initial denaturation at 94°C for 7 min followed by 35 cycles at 94°C for 1 min, 1 min at the annealing temperature (ITS: 50°C and D2/D3: 55°C), 72°C for 1 min and a final extension at 72°C for 7 min (Çimen et al., 2016).

Amplification of all PCR products and their respective negative amplification controls were verified by 1% (w/v) agarose gel electrophoresis in 1X TBE buffer stained with 0.5 X Hydragreen (ACTGene, United States). PCR products were purified for sequencing with Wizard SV Gel and PCR clean-up system (Promega, United States). Three purified amplicons obtained from three independent DNA extractions were sequenced for each molecular marker. Sequencing of each amplicon was performed in both directions.

Processing of the obtained sequences

The six sequences obtained for each marker were visualized, aligned, and edited manually using MEGA X software (Kumar et al., 2018) in order to obtain a consensus sequence. Its identity was initially verified by means of the Basic Local Alignment Search Tool (BLAST) (Altschul et al., 1990) on the basis of the non-redundant (nr) database of the NCBI. Consensus sequences of all markers were deposited in GenBank (NCBI) under the accession numbers listed in Table 1.

Table 1.

Accession numbers of sequences used for phylogenetic analysis of the EPN isolated and its bacterial symbiont.

10.21307_jofnem-2020-089-t001.jpg

Phylogenetic analysis

The consensus sequences of all markers were subsequently aligned with sequences of various species of EPNs deposited in GenBank (Table 1) using MUSCLE (Edgar, 2004) in MEGA X software (Kumar et al., 2018). The alignment obtained was edited so that all sequences used for analysis were of the same length and same genetic region.

Then, phylogenetic trees were independently constructed for each marker. The Tamura–Nei model of maximum likelihood estimation (Tamura and Nei, 1993) with gamma distribution and 1,000 bootstraps was used for the 18S fragment, and the General Time Reversible model of maximum likelihood estimation (Nei and Kumar, 2000) with gamma distribution and 1,000 bootstraps was used for the ITS fragment. Unweighted Pair Group Method with Arithmetic Mean (UPGMA) (Sneath and Sokal, 1973) with gamma distribution and 1,000 bootstraps was used for the D2/D3 fragment of rRNA 28S.

Because of the small number of sequences available for the 18S marker in GenBank, the additional phylogenetic analysis of concatenated sequences was made only with ITS (601 nts) and D2/D3 (754 nts) fragments. This phylogenetic tree was constructed using UPGMA (Sneath and Sokal, 1973) (Tamura–Nei model of maximum likelihood estimation (Tamura and Nei, 1993) with gamma distribution and 1,000 bootstraps).

Susceptibility of Epitrix cucumeris and Pandeleteius cinereus adults to EPNs

The susceptibility of E. cucumeris and P. cinereus adults (Hass avocado [P. americana] pests) to the EPNs isolated, and S. carpocapsae FA2015 as reference (commercially obtained), was evaluated and compared in vitro. Adults of both insect species were collected from a P. americana crop. In vitro bioassays were performed in multicell culture plates containing sterile Whatman No. 1 filter paper (GE Healthcare, United States). In total, 10 adults of each insect species were exposed individually to 0 IJs (sterile distilled water) and 50 IJs of each EPN. Viability of IJs was checked during the count before adding them to the plates. Plates were incubated at 26°C for 8 days, and adult survival was recorded every 24 hr.

To corroborate whether mortality was due to EPNs and check for EPNs reproduction inside the insects, adults were dissected in Ringer solution (9 g/liter NaCl, 0.4 g/liter KCl, 0.4 g/liter CaCl2, 0.2 g/liter NaHCO3) (Merck, United States). Each bioassay was repeated three times over time.

Statistical analysis

Survival analysis of insects exposed to the EPNs was performed using Graphpad Prism 6.01 software (GraphPad Software, United States) using the Kaplan–Meier method (Kaplan and Meier, 1958). To identify any significant differences between treatments, we used analysis of variance (ANOVA) testing and multiple comparisons with the Tukey test using the area under the curve.

Isolation and identification of bacterial symbiont

Isolation

Bacterial symbiont was isolated from hemolymph of G. mellonella larvae infected with the EPNs recovered, following the procedure described by Kazimierczak et al. (2017). After isolation, bacterial symbionts were stored at −80°C in LB Broth (Scharlab, Spain) with 20% (v/v) glycerol (Merck, United States) and deposited in the Microorganisms Collection of Pontificia Universidad Javeriana – CMPUJ.

Confirmation of bacterial symbiont’s isolate

The correspondence of bacterial isolates to what is expected for EPNs bacterial symbionts of the Xenorhabdus or Photorhabdus genus (Akhurst and Boemare, 2015; Boemare and Akhurst, 2015) was initially confirmed through microscopic and macroscopic morphological characteristics by Gram staining and, on NBTA (13 g/liter nutrient broth, 0.025 g/liter bromothymol blue, 0.04 g/liter chloride 2, 3,5-triphenyltetrazolium, and 15 g/liter agar), MacConkey (Scharlab, Spain) and blood agar (Becton, Dickinson and Company, United States). Samples were incubated at 26°C for 48 hr. To corroborate symptoms of G. mellonella larvae infected with the bacterial isolates, a colony was removed from NBTA medium after 48 hr growth at 26°C, and resuspended in saline solution (0.85% (w/v) NaCl [Merck, United States]). The suspension was serially diluted to 10−4. Then, 10 µl of the last dilution was injected into five larvae of G. mellonella with a syringe (Hamilton Company, United States). The surface of the larvae was disinfected with 1% NaClO (BNS S.A, Colombia) for 1 min, followed by three washes with sterile distilled water. As a negative control, 10 µl of sterile saline solution was injected into five larvae of G. mellonella.

Following injection, larvae were incubated at 26°C for 48 hr, after then, their coloration, mortality, and consistency were all checked.

Biochemical characterization

To identify metabolic characteristics of the bacterial isolate, a colony of the bacteria was exposed to a 3% hydrogen peroxide solution (Sigma-Aldrich, United States) using sterile wooden sticks. Subsequently, a colony was taken from nutrient agar (Scharlab, Spain) after 48 hr growth at 26°C and then resuspended in 0.85% API NaCl medium (BioMérieux, France). The API 20E fast identification system (BioMérieux, France) was then used according to the manufacturer’s instructions and incubated at 26°C for 48 hr. After incubation, results were read and interpreted following the manufacturer’s instructions.

Molecular identification

Bacterial genomic DNA was extracted following the CTAB-based DNA extraction protocol described by Feil et al. (2004). From the extracted DNA, a Multilocus Sequence Typing (MLST) analysis was performed with the partial sequences of five genes used as taxonomic markers. The following five sequences and primers were used: 16S rRNA using 16SP1 5′-GAAGAGTTTGATCATGGCTC-3′ and 16SP2 5′-AAGGAGGTGATCCAGCCGCA-3′ (Tailliez et al., 2006); dnaN (DNA polymerase III beta chain) using dnaN1 5′-GAAATTYATCATTGAACGWG-3′ and dnaN2 5′-CGCATWGGCATMACRAC-3′; recA (DNA recombinase) using recA1 5′-GCTATTGATGAAAATAAACA-3′ and recA2 5′-RATTTTRTCWCCRTTRTAGCT-3′; gltX (glutamyl-tRNA synthetase) using gltX1 5′-GCACCAAGTCCTACTGGCTA-3′ and gltX2 5′- GGCATRCCSACTTTACCCATA-3′; and rplB (50S subunit of ribosomal protein L2) using rplB1 5′-GGCAATTGTTAAATGTAAACC-3′ and rpblBXeno2 5′-GCGGCGTACGATGTATTGAT-3′ (Tailliez et al., 2010).

For molecular markers amplification, 25 µl of PCR mixture was prepared using 1X NH4 Buffer, 3 mM MgCl2, 0.2 mM dNTPs (Bioline, England), 0.5 µg/µl Bovine Serum Albumin (New England Biolabs, United States), forward primer and reverse primer for each 0.5 µM of taxonomic marker, Taq DNApolimerase 2U (Bioline, England) and 20 to 100 ng of bacterial DNA (Hominick et al., 1997; Tailliez et al., 2006).

The PCR protocol consisted of an initial denaturation at 94°C for 5 min, 30 cycles at 94°C for 30 sec, 30 sec at the annealing temperature of each pair of primers (16S: 64.9°C, dnaN: 45°C, recA: 45°C, gltx: 64.9°C and rplB: 58.3°C), extension at 72°C for 1 min, and a final extension at 72°C for 7 min. The specific amplification of all PCR products was verified by electrophoresis in 1% (w/v) agarose gel in 1X TBE buffer stained with 0.5 X Hydragreen (ACTGene, United States).

PCR products were purified for sequencing with Wizard SV Gel and PCR clean-up system (Promega, United States). Three independent amplicons were sequenced in both directions for each molecular marker.

Sequence processing

The six sequences obtained for each marker were visualized, aligned, and edited manually using the MUSCLE tool (Edgar, 2004) in the MEGA X software (Kumar et al., 2018) in order to obtain the respective consensus sequence. The consensus sequence for 16S marker was initially identified by means of EzBioCloud (Yoon et al., 2017), while BLAST (Altschul et al., 1990) was used for initial identification of other markers. The consensus sequences of all markers were deposited in GenBank (NCBI) under the accession numbers registered in Table 1.

Phylogenetic analysis

The consensus sequences of each marker were aligned with sequences of species deposited in GenBank (NCBI) using MUSCLE (Edgar, 2004) in MEGA X software (Kumar et al., 2018). The alignment obtained was edited so that all the sequences used in the analysis were of the same length and genetic region, and phylogenetic trees were constructed independently for each marker. Maximum likelihood estimation (Tamura–Nei model [Tamura and Nei, 1993] with gamma distribution, invariant sites (G + I), and 100 bootstraps) was used for the 16S fragment; maximum likelihood estimation (Kimura 2 model [Kimura, 1980] with gamma distribution, invariant sites (G + I), and 100 bootstraps) was used for the dnaN and rplB fragments; and maximum likelihood estimation (Tamura 3 model [Tamura, 1992] with gamma distribution, invariant sites (G + I), and 100 bootstraps) was used for the gltX and recA fragments.

In addition, the dnaN (815 nts), gltX (654 nts), recA (592 nts), and rplB (678 nts) markers were manually concatenated for construction of the phylogenetic tree using maximum likelihood estimation (General Time Reversible model (Nei and Kumar, 2000) with gamma distribution, invariant sites [G + I], and 100 bootstraps). The accession numbers of all the sequences used are found in Table 1.

Results

Isolation and identification of entomopathogenic nematodes

Only 14 out of 240 soil samples (5.8% recovery frequency) were found to contain EPNs. They were all from two of the 10 sites sampled (20% abundance) (site 1: CC and site 2: NF) (Table 2). However, it was only possible to multiply and maintain the isolate that was found in the soils of site 2. Only dead IJs were found when larvae with symptons of NEPs from site 1 were dissected.

Table 2.

Isolation of EPNs according to types of vegetation cover at sampling sites.

10.21307_jofnem-2020-089-t002.jpg

In total, 1,200 larvae of each host insect were exposed to all soil samples, but only 2.5% of G. mellonella and 0.33% of T. molitor larvae were affected by EPNs. Brown and flaccid larvae affected by nematodes exhibited the typical signs of Steinernematidae infection. On the other hand, the first IJs were recovered eight days after White traps settlement.

Table 2 shows EPNs isolation data in relation to types of vegetation cover and host insects. Nematodes were recovered only at sites whose soil temperatures were close to 16.5°C. Also, in both sites where EPNs were found, more G. mellonella larvae than T. molitor larvae were found dead with symptoms indicating EPNs infection. The greatest recovery frequency of EPNs was obtained from the natural forest plant cover site.

After initial BLAST identification of the 18S, ITS, and 28S markers from the isolate recovered, 100% identity and coverage were obtained with S. carpocapsae. This is consistent with the phylogenetic analysis of the concatenated 18S and ITS-28S markers (Fig. 1A, B) and confirms the specific identity of the isolate named S. carpocapsae BPS.

Figure 1:

Phylogenetic tree of S. carpocapsae BPS. A. Phylogenetic relationships from the 18S marker. B. Phylogenetic relationships from concatenated ITS and 28S markers. The relationships were inferred by maximum likelihood estimation for the 18S marker while UPGMA was used for the concatenated markers. Caenorhabditis. elegans was used as an outgroup. The numbers on the nodes indicate the bootstrap values (1,000 replications).

10.21307_jofnem-2020-089-f001.jpg

Susceptibility of Epitrix cucumeris and Pandeleteius cinereus to EPNs

Between 0 and 6.7% of E. cucumeris adults survived exposure to S. carpocapsae (BPS and FA2015, respectively), while this result was between 0 and 10% for P. cinereus adults, on the other hand, 100% of both pests survived to the sterile distilled water control. This indicates susceptibility of both pests to the EPNs evaluated (Fig. 2). Analysis using the area under the curve found significant differences only in the survival of E. cucumeris when exposed to the two isolates (df = 2; F = 89.32; P = <0.0001). Exposure to S. carpocapsae BPS resulted in less survival for this pest than did the reference isolate S. carpocapsae FA2015, which represents a higher mortality in less time.

Figure 2:

Susceptibility of two Hass avocado (P. americana) pest insects to S. carpocapsae BPS and S. carpocapsae FA2015. A. E. cucumeris. B. P. cinereus.

10.21307_jofnem-2020-089-f002.jpg

Adults and IJs of both S. carpocapsae isolates were present in 100% of the dead insects of both pests. In total, 37.1% of the nematodes found in dead E. cucumeris infected with S. carpocapsae BPS were adults, 41.1% were adults in individuals of this pest infected with S. carpocapsae FA2015, and the rest were IJs in both cases. In contrast, 49.8% of the nematodes found in dead P. cinereus insects infected with S. carpocapsae BPS were adults, 41.1% were adults in insects of this pest infected with S. carpocapsae FA2015, and the rest were IJs in both cases.

Isolation and confirmation of bacterial symbiont

The bacterial symbionts of S. carpocapsae BPS were Gram-negative bacilli of various sizes with irregular borders and with inclusion proteins that varied in quantity and size (Fig. 3A). Macroscopically, they grew in bright, granular, convex colonies with slightly irregular borders. They were blue on NBTA agar (Fig. 3B), pink with translucent margins on MacConkey agar (Fig. 3C), and yellowish-green with total hydrolysis of red blood cells (α-hemolysis) on blood agar (Fig. 3D). In addition, G. mellonella larvae infected with bacterial isolate reached 100% mortality at 48 hr as evidenced by their brown color and sagging.

Figure 3:

Microscopic and macroscopic morphological characteristics of the bacterial symbiont, Xenorhabdus nematophila BPS. A. Gram stain, 100X. B. Colonies on NBTA agar. C. Colony on MacConkey agar. D. Isolate of bacterial symbiont on blood agar, view from the back of the box. The bar measures 0.5 cm.

10.21307_jofnem-2020-089-f003.jpg

Biochemical characterization

As observed in the API biochemical gallery and under incubation conditions, the bacterial symbiont was negative for β-galactosidase, arginine-dihydrolase, lysine decarboxylase, ornithine decarboxylase, citrate assimilation, H2S production, urease, tryptophan deaminase, production of indole, acetoin production (Voges – Proskauer test), gelatinase, production of acid from glucose, mannitol, inositol, sorbitol, rhamnose, sucrose, melibiose, amygdalin, and arabinose.

Molecular identification

The initial identification, using EzBioCloud for the 16S marker and BLAST searches of the nr database of the NCBI for other markers, showed greatest similarity to the respective Xenorhabdus nematophila markers: 99.47% identity and 63.9% coverage were obtained for the 16S marker; 99% identity and 100% coverage were obtained for dnaN; 100% identity and coverage were obtained for recA; 99% identity and 100% coverage were obtained for gltX; and 100% identity and coverage were obtained for rplB. The phylogenetic analysis showed that the bacterial isolate was grouped at the same level with X. nematophila for the 16S marker as well as for the concatenated markers, thus confirming its taxonomic identity (Fig. 4A, B).

Figure 4:

Phylogenetic tree of the bacterial symbiont Xenorhabdus nematophila BPS. A. Phylogenetic relationships from the 16S marker. B. Phylogenetic relationships from concatenation of dnaN, recA, gltX, and rplB markers. The relationships were inferred by maximum likelihood estimation. Morganella morganii subsp. morganii was used as an outgroup for 16S, while Proteus mirabilis and Photorhabdus luminescens subsp. laumondii were used for concatenated markers. The numbers on the nodes indicate the bootstrap values (≥ 50%) (100 replicates).

10.21307_jofnem-2020-089-f004.jpg

Discussion

The recovery frequency gives an estimated of EPNs distribution between sampling points. This study found a low or uneven distribution of EPNs in the rural region evaluated in the municipality of Toribío in the department of Cauca, Colombia, which was evidenced in a low recovery frequency. Despite differences in habitats sampled and methodologies used for soil sampling and EPNs isolation, the recovery frequency obtained in this study was similar to that reported from soils of other Latin American countries such as México (6.6%, 4 positive samples out of 60) (Delgado-Gamboa et al., 2015) and Chile (7%, 97 positive samples out of 1,382) (Edgington et al., 2010), but different from others like Brazil (23.2%, 73 positive samples out of 315) (Foelkel et al., 2017).

In contrast, in this study only one isolate of S. carpocapsae was recovered and multiplied from soils of both natural habitats (natural forest) and crops (coffee cultivation) between 1,757 and 2,963 m.a.s.l, while in México Delgado-Gamboa et al. (2015) recovered four EPN isolates (three S. carpocapsae and one S. websteri) from soils of agricultural areas planted to agave (Agave angustifolia Haw) between 1,400 and 1,600 m.a.s.l, in Chile Edgington et al. (2010) recovered 101 EPN isolates, 94 Steinernema sp. (S. australe, S. unicornum, and S. feltiae) and 7 Heterorhabditis sp.( H. cf. safricana), from soils of natural habitats and agricultural areas between 0 and 2,499 m.a.s.l, while in Brazil Foelkel et al. (2017) recovered five EPN isolates which were closely related to Oscheius sp. from soil samples of an apple orchard (cultivar Eva) at 865 m.a.s.l., which points the variable and cosmopolitan distribution of EPNs (Rizvi et al., 2012). However, in this study both G. mellonella and T. molitor were used in combination as baits for EPNs recovery, in México and Chile studies only last-instar larvae of G. mellonella were used, while in Brazil T. molitor and Anastrepha fraterculus (Diptera: Tephritidae) were used independently as baits for EPNs recovery.

Regarding studies in Colombia, in terms of recovery frequency our result is similar to that reported by López-Núñez et al. (2007) who, in departments of Caldas, Quindío, Risaralda and Cundinamarca, found 3% of samples positive for EPNs (28 out of 945) corresponding to 26 isolates of Steinernema sp. (S. websteri and four undescribed Steinernema sp. taxa) and two of Heterorhabditis sp. (both undescribed) from soils of coffee crops and natural habitats between 1,203 and 1,478 m.a.s.l using only last-instar G. mellonella larvae for EPNs recovery. However, analyzing only positive sites for EPNs, López-Núñez et al. (2007) obtained 2 and 6% of recovery frequency in forests, which is lower than that obtained in this study for natural forest (50%), and indicates a higher distribution of EPNs between sampling points of this habitat in this study. In contrast, they found recovery frequencies for EPNs from 4 to 18% for areas cultivated in coffee (C. arabica), which was similar to the results obtained in this study for coffee cultivation (8.3%).

The differences between the recovery frequencies found in this study compared to the results of the other studies referenced from Latin America and Colombia could be due to that factors such as soil type, distribution of suitable hosts, physiological and behavioral adaptations are key factors affecting the distribution of EPN species (Adams et al., 2006; Stuart et al., 2006). However, some of these resources have heterogeneous distribution, therefore, nematode populations are highly aggregated (Ettema and Lal, 2006).

Despite EPNs were isolated at two kind of habitats; one cultivated site (CC) and at one natural habitat (NF), the greater recovery frequency of the second one indicates that, despite the fact that natural habitats contain greater diversity of insects controlled by one or another natural enemy, it is likely that at NF there is an ecological imbalance which favors the incidence of EPNs in this habitat where they were widely distributed in the soil (Campos-Herrera et al., 2007; Jaffuel et al., 2018). By the other hand, while monocultures may have greater availability of hosts susceptible to attack by EPNs, the continuous use of pesticides can limit availability and negatively affect presence of biocontrol agents, which could be the reason for a low recovery frequency of EPNs at CC or even the no recovery of EPNs in other crop sites evaluated in this study.

In terms of abundance, which gives an estimated about the distribution of EPNs between sampling sites, in this study the EPNs abundance, was lower than registered by López-Llano and Soto-Giraldo (2016) whom obtained 88.2% of abundance (15 positive sites for EPNs out of 17) from sugarcane crops (Saccharum officinarum) in the department of Caldas, in which were recovered 15 isolates of Steinernema sp. and 6 of Heterorhabditis sp, while Melo et al. (2009) obtained 74% of abundance (17 positive sites for EPNs out of 23) from crops between 990 and 1,660 m.a.s.l. in departments of Quindío, Risaralda, Caldas and Cauca. However, in contrast for what was done in this study, both other ones mentioned above used only last-instar larvae of G. mellonella for EPNs recovery.

Although we allegedly detect EPNs at two sites, it was only possible to multiply and maintain the isolate from NF. This may have been due to ignorance of biological, ecological, and temperature conditions required for these nematodes to infect susceptible hosts and indicate a need for additional studies in the area using other detection/extraction methods and/or host insects.

Even though EPNs were allegedly detected at both sites through using the two host insects evaluated, it was evident that G. mellonella was generally more efficient at detecting EPNs than was T. molitor. This indicates that G. mellonella is more susceptible to the EPNs potentially present at both sites. Nevertheless, there were more G. mellonella larvae affected by EPNs at the NF site than at the CC site, while the same number of affected T. molitor larvae was obtained at both sites. This can be attributed to the abundance and nature of the EPNs potentially present at the CC site.

EPNs of the genera Heterorhabditis and Steinernema have been found on all continents: in South America they have been recovered in eight countries (Stock, 2015; Stuart et al., 2015). In Colombia, Steinernema species reported include S. websteri (López-Núñez et al., 2007), a new species named S. colombiense (López-Núñez et al., 2008), and S. kraussei (Melo et al., 2009). The isolate obtained in this study was identified as S. carpocapsae, a nematode that in South America had previously only been reported in Argentina, Brazil, and Perú (Cabi.org, 2018; San-Blas et al., 2019).

Steinernema carpocapsae is found throughout the world (Hominick, 2002), but this species is not frequently isolated (Yan et al., 2016). This is probably due to the species’ ambush behavior. When G. mellonella is used as a target insect to isolate S. carpocapsae, its field frequency is underestimated (Mráček et al., 2005). Despite this, S. carpocapsae has been recovered in grassland areas (602-677 m.a.s.l, average soil temperature of 11°C) (Campos-Herrera et al., 2007) and in forests (1,700 m.a.s.l, soil temperature of 19°C) (Yan et al., 2016). These factors are similar to those in the areas sampled in this study.

Although it has been reported that S. carpocapsae can infect about 95% of the larvae of T. molitor (San-Blas et al., 2012), this study found that a greater number of G. mellonella larvae were affected. This could be attributable to the poor mobility of T. molitor in soil combined with the ambush behavior of S. carpocapsae. This EPN attacks high-mobile insects more efficiently (Banu et al., 2017). Our findings are consistent with those obtained in a study by Yan et al. (2016) in which S. carpocapsae isolate was exposed to the same two insect larvae, but nematodes were only recovered from G. mellonella.

Steinernema carpocapsae has been shown to establish a specific and exclusive symbiosis with X. nematophila (Cowles and Goodrich-Blair, 2008) as its only bacterial symbiont, so its identification as the bacterial isolate of this study was expected. The isolate showed morphological characteristics typical of X. nematophila in phase 1 (Akhurst and Boemare, 2015), while most of its biochemical characteristics obtained. Our results agree with what has been reported for this species elsewhere, except for our negative results for assimilation of citrate, gelatinase activity and acid production from glucose. These differences may be attributable to the fact that metabolic specialization can be found even within the same species (Ponomarova and Patil, 2015) caused by factors such as temperature that affect enzymatic activity (Peterson et al., 2007). Therefore, the conditions in which the biochemical tests were carried out can give rise to different results to what is reported for bacteria of the same species.

Steinernema carpocapsae is the EPN species most commonly used to control insect pests associated with the soil surface and leaf area (Lacey and Georgis, 2012). Although its potential as a control agent has been demonstrated in other insects of the Curculionidae family such as the red weevil, Rhynchophorus ferrugineus (Manachini et al., 2013) and the coffee borer beetle, Hypothenemus hampei (Manton et al., 2012), there had been no previous reports of its efficacy for controlling P. cinereus. Therefore, this is the first report of its susceptibility to S. carpocapsae. On the other hand, the efficacy of S. carpocapsae for controlling two members of the Chrysomelidae family, western corn rootworms, Diabrotica virgifera (Journey and Ostlie, 2000), and potato flea beetles, Epitrix spp. (Parker et al., 2012), has already been demonstrated. This study has shown that P. cinereus and E. cucumeris adults which damage Hass avocado crops are susceptible to both isolates of S. carpocapsae, and that S. carpocapsae BPS has greater capacity to cause less survival of E. cucumeris adults.

The results obtained for susceptibility of both pests to S. carpocapsae are important because these pests affect Hass avocado crops (P. americana) in Colombia. They eat the leaves of the plants causing cuts that decrease leaf area thereby reducing the rate of photosynthesis which in turn reduces the yield of the plant. Similarly, P. cinereus causes damage to the ovary, petals, and newly formed fruit causing them to fall from the trees (Londoño et al., 2014). Consequently, our findings of in vitro susceptibility of both pests to S. carpocaosae now need to be corroborated by field studies that can determine the potential of these EPNs for controlling these two pests of economic interest to Hass avocado cultivation (P. americana) in Colombia.

Conclusions

These results are the first record of S. carpocapase and its bacterial symbiont in Cauca, Colombia. They establish its potential for controlling two pests that affect Hass avocado crops (adults of P. cinereus and E. cucumeris). Additional studies are needed to complement and confirm these results, especially field trials to determine the possibility of using these nematodes in integrated pest management programs for these pests.

Acknowledgements

This work was funded by Pontificia Universidad Javeriana Vicerrectoría de Investigación grant No. 7028.

References


  1. Adams, B. J. , Fodor, A. , Koppenhöfer, H. S. , Stackenbrandt, E. , Stock, S. P. and Klein, M. G. 2006. Biodiversity and systematic of nematode–bacterium entomopathogens. Biological Control 38:4–21.
  2. Akhurst, R. J. and Boemare, N. E. 2015. “ Xenorhabdus ”, Pp. 831–838. In Whitman, W. B. , Rainey, F. , Kämpfer, P. , Trujillo, M. , Chun, J. , DeVos, P. , Hedlund, B. and Dedysh, S. (Eds), Bergey’s manual of systematics of archaea and bacteria. New Jersey: John Wiley & Sons, doi: 10.1002/9781118960608.gbm01173.
  3. Altschul, S. F. , Gish, W. , Miller, W. , Myers, E. W. and Lipman, D. J. 1990. Basic local alignment search tool. Journal of Molecular Biology 215:403–410, doi: 10.1016/S0022-2836(05)80360-2.
  4. Banu, J. G. , Cannayane, I. and Meena, S. K. 2017. “Entomopathogenic nematodes: general biology and behaviour”, In Abd-Elgawad, M. M. M. , Askary, T. H. and Coupland, J. (Eds), Biocontrol agents: entomopathogenic and slug parasitic nematodes CABI Publishing, Wallington, pp. 63–87.
  5. Bedding, R. A. and Akhurst, R. J. 1975. A simple technique for the detection of insect parasitic rhabditid nemadodes in soil. Nematologica 21:109–116..
  6. Boemare, N. E. and Akhurst, R. J. 2015. “ Photorhabdus ”, Pp. 831–838 In Whitman, W. B. , Rainey, F. , Kämpfer, P. , Trujillo, M. , Chun, J. , DeVos, P. , Hedlund, B. and Dedysh, S. (Eds), Bergey’s manual of systematics of archaea and bacteria. New Jersey: John Wiley & Sons, doi: 10.1002/9781118960608.gbm01159.
  7. Cabi.org . 2018. Invasive Species Compendium, Steinernema carpocapsae . available at: https://www.cabi.org/isc/datasheet/51706 (accessed March 26, 2019).
  8. Campos-Herrera, R. and Gutiérrez, C. 2009. A laboratory study on the activity of Steinernema feltiae (Rhabditida: Steinernematidae) Rioja strain against horticultural insect pests. Journal of Pest Science 82:305–309, doi: 10.1007/s10340-009-0247-z.
  9. Campos-Herrera, R. , Escuer, M. , Labrador, S. , Robertson, L. , Barrios, L. and Gutiérrez, C. 2007. Distribution of the entomopathogenic nematodes from La Rioja (Northern Spain). Journal of Invertebrate Pathology 95:125–139, doi: 10.1016/j.jip.2007.02.003.
  10. Çimen, H. , Půža, V. , Nermuť, J. , Hatting, J. , Ramakuwela, T. , Faktorová, L. and Hazir, S. 2016. Steinernema beitlechemi n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from South Africa. Nematology 18:439–453, doi: 10.1163/15685411-00002968.
  11. Cowles, C. E. and Goodrich-Blair, H. 2008. The Xenorhabdus nematophila nilABC genes confer the ability of Xenorhabdus spp. to colonize Steinernema carpocapsae nematodes. Journal of Bacteriology 190:4121–4128, doi: 10.1128/JB.00123-08.
  12. De Brida, A. L. , Rosa, J. M. , Oliveira, C. M. , Castro, B. M. , Serrão, J. E. , Zanuncio, J. C. , Leite, L. G. and Wilcken, S. R. 2017. Entomopathogenic nematodes in agricultural areas in Brazil. Scientific Reports 7:1–7, doi: 10.1038/srep45254.
  13. Delgado-Gamboa, J. R. , Ruíz-Vega, J. , Ibarra-Rendón, J. E. , Aquino-Bolaños, T. and Giron-Pablo, S. 2015. Isolation and Identification of Native Entomopathogenic Nematodes (Nematoda: Rhabditidae) and Potential for Controlling Scyphophorus acupunctatus in a Laboratory. Southwestern Entomologist 40:731–739, doi: 10.3958/059.040.0401.
  14. Edgar, R. C. 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research 32:1792–1797, doi: 10.1093/nar/gkh340.
  15. Edgington, S. , Buddie, A. G. , Moore, D. , France, A. , Merino, L. , Tymo, L. M. and Hunt, D. J. 2010. Diversity and distribution of entomopathogenic nematodes in Chile. Nematology 12:915–928, doi: 10.1163/138855410X498897.
  16. Ettema, C. H. 2006. “Nematodes”, In Lal, R. (Ed.), Encyclopedia of soil science 2nd ed. New York, NY: Taylor & Francis Group, LLC, Vol. 2, pp. 1114–1118.
  17. Feil, W. S. , Feil, H. and Copeland, A. 2004. Bacterial genomic DNA isolation using CTAB. Joint Genome Institute, available at: http://1ofdmq2n8tc36m6i46scovo2e.wpengine.netdna-cdn.com/wp-content/uploads/2014/02/JGI-Bacterial-DNA-isolation-CTAB-Protocol-2012.pdf (accessed July 13, 2018).
  18. Foelkel, E. , Voss, M. , Monteiro, L. B. and Nishimura, G. 2017. Isolation of entomopathogenic nematodes in an apple orchard in Southern Brazil and its virulence to Anastrepha fraterculus (Diptera: Tephritidae) larvae, under laboratory conditions. Brazilian Journal of Biology 77:22–28, doi: 10.1590/1519-6984.08315.
  19. Griffin, C. T. , Boemare, N. T. and Lewis, E. E. 2005. “Biology and behaviour”, In Grewal, P. S. , Ehlers, R. U. and Shapiro-Ilan, D. I. (Eds), Nematodes as biocontrol agents. Wallingford: CABI Publishing, pp. 48–49.
  20. Hazir, S. , Kaya, H. K. , Stock, S. P. and Keskin, N. 2003. Entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) for biological control of soil pests. Turkish Journal of Biology 27:181–202.
  21. Hominick, W. M. 2002. “Biogeography”, in Gaugler, R. (Ed.), Entomopathogenic nematology CABI Publishing, New Brunswick, 118–136.
  22. Hominick, W. M. , Briscoe, B. R. , del Pino, F. G. , Heng, J. , Hunt, D. J. , Kozodoy, E. , Mracek, Z. , Nguyen, K. B. , Reid, A. P. , Spiridonov, S. , Stock, P. , Sturhan, D. , Waturu, C. and Yoshida, M. 1997. Biosystematics of entomopathogenic nematodes: current status, protocols and definitions. Journal of Helminthology 71:271–298, doi: 10.1017/S0022149X00016096.
  23. Jaffuel, G. , Blanco-Pérez, R. , Hug, A. -S. , Chiriboga, X. , Meuli, R. G. , Mascher, F. , Turlings, T. C. J. and Campos-Herrera, R. 2018. The evaluation of entomopathogenic nematode soil food web assemblages across Switzerland reveals major differences among agricultural, grassland and forest ecosystems. Agriculture, Ecosystems & Environment 262:48–57, doi: 10.1016/j.agee.2018.04.008.
  24. Journey, A. M. and Ostlie, K. R. 2000. Biological Control of the Western Corn Rootworm (Coleoptera: Chrysomelidae) Using the Entomopathogenic Nematode, Steinernema carpocapsae . Environmental Entomology 29:822–831, doi: 10.1603/0046-225X-29.4.822.
  25. Kaplan, E. L. and Meier, P. 1958. Nonparametric Estimation from Incomplete Observations. Journal of the American Statistical Association 53:457–481, doi: 10.2307/2281868.
  26. Kazimierczak, W. , Skrzypek, H. , Sajnaga, E. , Skowronek, M. , Waśko, A. and Kraft, A. 2017. Strains of Photorhabdus spp. associated with polish Heterorhabditis isolates: their molecular and phenotypic characterization and symbiont exchange. Archives of Microbiology 199:979–989, doi: 10.1007/s00203-017-1368-z.
  27. Kimura, M. 1980. A simple method for estimating evolutionary rate of base substitutions through comparative studies of nucleotide sequences. Journal of Molecular Evolution 16:111–120, doi: 10.1007/BF01731581.
  28. Kumar, S. , Stecher, G. , Li, M. , Knyaz, C. and Tamura, K. 2018. MEGA X: molecular evolutionary genetics analysis across computing platforms. Molecular Biology and Evolution 35:1547–1549, doi: 10.1093/molbev/msy096.
  29. Labaude, S. and Griffin, C. T. 2018. Transmission success of entomopathogenic nematodes used in pest control. Insects 9:1–20, doi: 10.3390/insects9020072.
  30. Lacey, L. A. and Georgis, R. 2012. Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. Journal of Nematology 44:218–225.
  31. Liu, J. and Berry, R. E. 1995. Natural distribution of entomopathogenic nematodes (Rhabditida: Heterorhabditidae and Steinernematidae) in Oregon soils. Environmental Entomology 24:159–163, doi: 10.1093/ee/24.1.159.
  32. Londoño, M. E. , Kondo, D. T. , Carabalí, A. , Varón, E. and Caicedo, A. M. 2014. “Insectos y Ácaros”, In Bernal, J. , Díaz, C. , Osorio, C. , Tamayo, A. , Osorio, W. , Córdoba, O. , Londoño, M. E. , Kondo, D. T. , Carabalí, A. , Varón, E. , Caicedo, A. M. , Tamayo, P. J. , Sandoval, A. , Forero, F. , García, J. and Londoño, M. (Eds), Actualización Tecnológica y Buenas Práctica Agrícolas (BPA) en el Cultivo de Aguacate Corpoica, Medellín, pp. 227–284.
  33. López-Llano, R. A. and Soto-Giraldo, A. 2016. Aislamiento de nematodos entomopatógenos nativos en cultivos de caña panelera y pruebas de patogenicidad sobre Diatraea saccharalis (Lepidoptera: Crambidae). Boletín Científico. Centro de Museos. Museo de Historia Natural 20:114–123.
  34. López-Núñez, J. C. , Cano, L. , Góngora-Botero, C. E. and Stock, S. P. 2007. Diversity and evolutionary relationships of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) from the Central Andean region of Colombia. Nematology 9:333–341, doi: 10.1163/156854107781351972.
  35. López-Núñez, J. C. , Plichta, K. , Góngora-Botero, C. E. and Stock, S. P. 2008. A new entomopathogenic nematode, Steinernema colombiense n. sp. (Nematoda: Steinernematidae), from Colombia. Nematology 10:561–574, doi: 10.1163/156854108784513879.
  36. Manachini, B. , Schillaci, D. and Arizza, V. 2013. Biological Responses of Rhynchophorus ferrugineus (Coleoptera: Curculionidae) to Steinernema carpocapsae (Nematoda: Steinernematidae). Journal of Economic Entomology 106:1582–1589, doi: 10.1603/EC13031.
  37. Manton, J. L. , Hollingsworth, R. G. and Cabos, R. Y. M. 2012. Potential of Steinernema carpocapsae (Rhabditida: Steinernematidae) Against Hypothenemus hampei (Coleoptera: Curculionidae) in Hawaii. Florida Entomologist 95:1194–1197, doi: 10.1653/024.095.0452.
  38. Melo, E. L. , Ortega, C. A. , Susurluk, A. , Gaigl, A. and Bellotti, A. C. 2009. Poblaciones nativas de nematodos entomopatógenos (Rhabditida) en cuatro departamentos de Colombia. Revista Colombiana de Entomología 35:28–33.
  39. Morise, H. , Miyazaki, E. , Yoshimitsu, S. and Eki, T. 2012. Profiling nematode communities in unmanaged flowerbed and agricultural field soils in Japan by DNA barcode sequencing. PLoS ONE 7:1–10, doi: 10.1371/journal.pone.0051785.
  40. Mráček, Z. , Bečvář, S. , Kindlmann, P. and Jersáková, J. 2005. Habitat preference for entomopathogenic nematodes, their insect hosts and new faunistic records for the Czech Republic. Biological Control 34:27–37, doi: 10.1016/j.biocontrol.2005.03.023.
  41. Nei, M. and Kumar, S. 2000. Molecular Evolution and Phylogenetics. Oxford: Oxford University Press.
  42. Nguyen, K. B. , Malan, A. P. and Gozel, U. 2006. Steinernema khoisanae n. sp (Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa. Nematology 8:157–175, doi: 10.1163/156854106777998728.
  43. Orozco, R. A. , Lee, M. M. and Stock, S. P. 2014. Soil sampling and isolation of entomopathogenic nematodes (Steinernematidae, Heterorhabditidae). Journal of Visualized Experiments 89:1–8, doi: 10.3791/52083.
  44. Parker, J. , Miles, C. , Murray, T. and Snyder, W. 2012. Organic management of flea beetles. Washington State University PNW 640:1–9.
  45. Peterson, M. E. , Daniel, R. M. , Danson, M. J. and Eisenthal, R. 2007. The dependence of enzyme activity on temperature: determination and validation of parameters. Biochemical Journal 402:331–337, doi: 10.1042/BJ20061143.
  46. Poinar, G. O. 1972. Nematodes as facultative parasites of insects. Annual Review of Entomology 17:103–122.
  47. Ponomarova, O. and Patil, K. R. 2015. Metabolic interactions in microbial communities: untangling the Gordian knot. Current Opinion in Microbiology 27:37–44, doi: 10.1016/j.mib.2015.06.014.
  48. Rizvi, P. Q. , Ahmad, S. K. , Choudhury, A. and Ali, A. 2012. “Biopesticides in ecologically-based integrated pest management”, In Abrol, D. P. and Shankar, U. (Ed.), Integrated pest management principles and practice. London: CAB International, pp. 133–161.
  49. San-Blas, E. , Pembroke, B. and Gowen, S. R. 2012. Scavenging and infection of different hosts by Steinernema carpocapsae . Nematropica 42:123–130.
  50. San-Blas, E. , Campos-Herrera, R. , Dolinski, C. , Monteiro, C. , Andaló, V. , Leite, L. G. , Rodríguez, M. G. , Morales-Montero, P. , Sáenz-Aponte, A. , Cedano, C. , López-Nuñez, J. C. , Del Valle, E. , Doucet, M. , Lax, P. , Navarro, P. D. , Báez, F. , Llumiquinga, P. , Ruiz-Vega, J. , Guerra-Moreno, A. and Stock, S. P. 2019. Entomopathogenic nematology in Latin America: a brief history, current research and future prospects. Journal of Invertebrate Pathology 165:22–45, doi: 10.1016/j.jip.2019.03.010.
  51. Shapiro-Ilan, D. I. , Han, R. and Qiu, X. 2014. “Production of entomopathogenic nematodes”, In Morales-Ramos, J. A. , Rojas, M. G. and Shapiro-Ilan, D. I. (Eds), Mass production of beneficial organisms: invertebrates and entomopathogens. Waltham: Academic Press, pp. 321–355.
  52. Sneath, P. H. A. and Sokal, R. R. 1973. Numerical taxonomy: the principles and practice of numerical classification. San Francisco, CA: W. H. Freeman.
  53. Stock, S. P. 2015. “Diversity, biology and evolutionary relationships”, In Campos-Herrera, R. (Ed.), Nematode pathogenesis of insects and other pests: ecology and applied technologies for sustainable plant and crop protection. Neuchâtel, Cham: Springer, pp. 3–27.
  54. Stock, S. P. , Campbell, J. F. and Nadler, S. A. 2001. Phylogeny of Steinernema Travassos, 1927 (Cephalobina: Steinernematidae) inferred from ribosomal DNA sequences and morphological characters. Journal of Parasitology 87:877–889, doi: 10.1645/0022-3395(2001)087[0877: POSTCS]2.0.CO;2.
  55. Stuart, R. J. , Barbercheck, M. E. and Grewal, P. S. 2015. “Entomopathogenic Nematodes in the Soil Environment: Distributions, Interactions and the Influence of Biotic and Abiotic Factors”, In Campos-Herrera, R. (Ed.), Nematode Pathogenesis of insects and other pests: ecology and applied technologies for sustainable plant and crop protection Springer, Neuchâtel, Cham, 97–137.
  56. Stuart, R. J. , Barbercheck, M. E. , Grewal, P. S. , Taylor, R. A. J. and Hoy, C. W. 2006. Population biology of entomopathogenic nematodes: Concepts, issues, and models. Biological Control 38:80–102.
  57. Tailliez, P. , Pagès, S. , Ginibre, N. and Boemare, N. 2006. New insight into diversity in the genus Xenorhabdus, including the description of ten novel species. International Journal of Systematic and Evolutionary Microbiology 56:2805–2818, doi: 10.1099/ijs.0.64287-0.
  58. Tailliez, P. , Laroui, C. , Ginibre, N. , Paule, A. , Pagès, S. and Boemare, N. 2010. Phylogeny of Photorhabdus and Xenorhabdus based on universally conserved protein-coding sequences and implications for the taxonomy of these two genera. Proposal of new taxa: X. vietnamensis sp. nov., P. luminescens subsp. caribbeanensis subsp. nov., P. luminescens subsp. hainanensis subsp. nov., P. temperata subsp. khanii subsp. nov., P. temperata subsp. tasmaniensis subsp. nov., and the reclassification of P. luminescens subsp. thracensis as P. temperata subsp. thracensis comb. nov. International Journal of Systematic and Evolutionary Microbiology 60:1921–1937, doi: 10.1099/ijs.0.014308-0.
  59. Tamura, K. 1992. Estimation of the number of nucleotide substitutions when there are strong transition-transversion and G + C-content biases. Molecular Biology and Evolution 9:678–687, doi: 10.1093/oxfordjournals.molbev.a040752.
  60. Tamura, K. and Nei, M. 1993. Estimation of the number of nucleotide substitutions in the control region of mitochondrial DNA in humans and chimpanzees. Molecular Biology and Evolution 10:512–526, doi: 10.1093/oxfordjournals.molbev.a040023.
  61. Van Zyl, C. and Malan, A. P. 2014. The role of entomopathogenic nematodes as biological control agents of insect pests, with emphasis on the history of their mass culturing and in vivo production. African Entomology 22:235–249, doi: 10.4001/003.022.0222.
  62. Varón de Agudelo, F. and Castillo, G. 2001. Seminario – Taller sobre identificación de nematodos de importancia agrícola ICA and Asociación Colombiana de Fitopatología (Ascolfi), Palmira.
  63. Vrain, T. C. , Wakarchuk, D. A. , Lévesque, A. C. and Hamilton, R. I. 1992. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundamental and Applied Nematology 15:563–573.
  64. White, G. F. 1927. A method for obtaining infective nematode larvae from cultures. Science 66:302–303, doi: 10.1126/science.66.1709.302-a.
  65. Yan, X. , Waweru, B. , Qiu, X. , Hategekimana, A. , Kajuga, J. , Li, H. , Edgington, S. , Umulisa, C. , Han, R. and Toepfer, S. 2016. New entomopathogenic nematodes from semi-natural and small-holder farming habitats of Rwanda. Biocontrol Science and Technology 26:820–834, doi: 10.1080/09583157.2016.1159658.
  66. Yoon, S. H. , Ha, S. M. , Kwon, S. , Lim, J. , Kim, Y. , Seo, H. and Chun, J. 2017. Introducing EzBioCloud: a taxonomically united database of 16S rRNA and whole genome assemblies. International Journal of Systematic and Evolutionary Microbiology 67:1613–1617, doi: 10.1099/ijsem.0.001755.
XML PDF Share

FIGURES & TABLES

Figure 1:

Phylogenetic tree of S. carpocapsae BPS. A. Phylogenetic relationships from the 18S marker. B. Phylogenetic relationships from concatenated ITS and 28S markers. The relationships were inferred by maximum likelihood estimation for the 18S marker while UPGMA was used for the concatenated markers. Caenorhabditis. elegans was used as an outgroup. The numbers on the nodes indicate the bootstrap values (1,000 replications).

Full Size   |   Slide (.pptx)

Figure 2:

Susceptibility of two Hass avocado (P. americana) pest insects to S. carpocapsae BPS and S. carpocapsae FA2015. A. E. cucumeris. B. P. cinereus.

Full Size   |   Slide (.pptx)

Figure 3:

Microscopic and macroscopic morphological characteristics of the bacterial symbiont, Xenorhabdus nematophila BPS. A. Gram stain, 100X. B. Colonies on NBTA agar. C. Colony on MacConkey agar. D. Isolate of bacterial symbiont on blood agar, view from the back of the box. The bar measures 0.5 cm.

Full Size   |   Slide (.pptx)

Figure 4:

Phylogenetic tree of the bacterial symbiont Xenorhabdus nematophila BPS. A. Phylogenetic relationships from the 16S marker. B. Phylogenetic relationships from concatenation of dnaN, recA, gltX, and rplB markers. The relationships were inferred by maximum likelihood estimation. Morganella morganii subsp. morganii was used as an outgroup for 16S, while Proteus mirabilis and Photorhabdus luminescens subsp. laumondii were used for concatenated markers. The numbers on the nodes indicate the bootstrap values (≥ 50%) (100 replicates).

Full Size   |   Slide (.pptx)

REFERENCES

  1. Adams, B. J. , Fodor, A. , Koppenhöfer, H. S. , Stackenbrandt, E. , Stock, S. P. and Klein, M. G. 2006. Biodiversity and systematic of nematode–bacterium entomopathogens. Biological Control 38:4–21.
  2. Akhurst, R. J. and Boemare, N. E. 2015. “ Xenorhabdus ”, Pp. 831–838. In Whitman, W. B. , Rainey, F. , Kämpfer, P. , Trujillo, M. , Chun, J. , DeVos, P. , Hedlund, B. and Dedysh, S. (Eds), Bergey’s manual of systematics of archaea and bacteria. New Jersey: John Wiley & Sons, doi: 10.1002/9781118960608.gbm01173.
  3. Altschul, S. F. , Gish, W. , Miller, W. , Myers, E. W. and Lipman, D. J. 1990. Basic local alignment search tool. Journal of Molecular Biology 215:403–410, doi: 10.1016/S0022-2836(05)80360-2.
  4. Banu, J. G. , Cannayane, I. and Meena, S. K. 2017. “Entomopathogenic nematodes: general biology and behaviour”, In Abd-Elgawad, M. M. M. , Askary, T. H. and Coupland, J. (Eds), Biocontrol agents: entomopathogenic and slug parasitic nematodes CABI Publishing, Wallington, pp. 63–87.
  5. Bedding, R. A. and Akhurst, R. J. 1975. A simple technique for the detection of insect parasitic rhabditid nemadodes in soil. Nematologica 21:109–116..
  6. Boemare, N. E. and Akhurst, R. J. 2015. “ Photorhabdus ”, Pp. 831–838 In Whitman, W. B. , Rainey, F. , Kämpfer, P. , Trujillo, M. , Chun, J. , DeVos, P. , Hedlund, B. and Dedysh, S. (Eds), Bergey’s manual of systematics of archaea and bacteria. New Jersey: John Wiley & Sons, doi: 10.1002/9781118960608.gbm01159.
  7. Cabi.org . 2018. Invasive Species Compendium, Steinernema carpocapsae . available at: https://www.cabi.org/isc/datasheet/51706 (accessed March 26, 2019).
  8. Campos-Herrera, R. and Gutiérrez, C. 2009. A laboratory study on the activity of Steinernema feltiae (Rhabditida: Steinernematidae) Rioja strain against horticultural insect pests. Journal of Pest Science 82:305–309, doi: 10.1007/s10340-009-0247-z.
  9. Campos-Herrera, R. , Escuer, M. , Labrador, S. , Robertson, L. , Barrios, L. and Gutiérrez, C. 2007. Distribution of the entomopathogenic nematodes from La Rioja (Northern Spain). Journal of Invertebrate Pathology 95:125–139, doi: 10.1016/j.jip.2007.02.003.
  10. Çimen, H. , Půža, V. , Nermuť, J. , Hatting, J. , Ramakuwela, T. , Faktorová, L. and Hazir, S. 2016. Steinernema beitlechemi n. sp., a new entomopathogenic nematode (Nematoda: Steinernematidae) from South Africa. Nematology 18:439–453, doi: 10.1163/15685411-00002968.
  11. Cowles, C. E. and Goodrich-Blair, H. 2008. The Xenorhabdus nematophila nilABC genes confer the ability of Xenorhabdus spp. to colonize Steinernema carpocapsae nematodes. Journal of Bacteriology 190:4121–4128, doi: 10.1128/JB.00123-08.
  12. De Brida, A. L. , Rosa, J. M. , Oliveira, C. M. , Castro, B. M. , Serrão, J. E. , Zanuncio, J. C. , Leite, L. G. and Wilcken, S. R. 2017. Entomopathogenic nematodes in agricultural areas in Brazil. Scientific Reports 7:1–7, doi: 10.1038/srep45254.
  13. Delgado-Gamboa, J. R. , Ruíz-Vega, J. , Ibarra-Rendón, J. E. , Aquino-Bolaños, T. and Giron-Pablo, S. 2015. Isolation and Identification of Native Entomopathogenic Nematodes (Nematoda: Rhabditidae) and Potential for Controlling Scyphophorus acupunctatus in a Laboratory. Southwestern Entomologist 40:731–739, doi: 10.3958/059.040.0401.
  14. Edgar, R. C. 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research 32:1792–1797, doi: 10.1093/nar/gkh340.
  15. Edgington, S. , Buddie, A. G. , Moore, D. , France, A. , Merino, L. , Tymo, L. M. and Hunt, D. J. 2010. Diversity and distribution of entomopathogenic nematodes in Chile. Nematology 12:915–928, doi: 10.1163/138855410X498897.
  16. Ettema, C. H. 2006. “Nematodes”, In Lal, R. (Ed.), Encyclopedia of soil science 2nd ed. New York, NY: Taylor & Francis Group, LLC, Vol. 2, pp. 1114–1118.
  17. Feil, W. S. , Feil, H. and Copeland, A. 2004. Bacterial genomic DNA isolation using CTAB. Joint Genome Institute, available at: http://1ofdmq2n8tc36m6i46scovo2e.wpengine.netdna-cdn.com/wp-content/uploads/2014/02/JGI-Bacterial-DNA-isolation-CTAB-Protocol-2012.pdf (accessed July 13, 2018).
  18. Foelkel, E. , Voss, M. , Monteiro, L. B. and Nishimura, G. 2017. Isolation of entomopathogenic nematodes in an apple orchard in Southern Brazil and its virulence to Anastrepha fraterculus (Diptera: Tephritidae) larvae, under laboratory conditions. Brazilian Journal of Biology 77:22–28, doi: 10.1590/1519-6984.08315.
  19. Griffin, C. T. , Boemare, N. T. and Lewis, E. E. 2005. “Biology and behaviour”, In Grewal, P. S. , Ehlers, R. U. and Shapiro-Ilan, D. I. (Eds), Nematodes as biocontrol agents. Wallingford: CABI Publishing, pp. 48–49.
  20. Hazir, S. , Kaya, H. K. , Stock, S. P. and Keskin, N. 2003. Entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) for biological control of soil pests. Turkish Journal of Biology 27:181–202.
  21. Hominick, W. M. 2002. “Biogeography”, in Gaugler, R. (Ed.), Entomopathogenic nematology CABI Publishing, New Brunswick, 118–136.
  22. Hominick, W. M. , Briscoe, B. R. , del Pino, F. G. , Heng, J. , Hunt, D. J. , Kozodoy, E. , Mracek, Z. , Nguyen, K. B. , Reid, A. P. , Spiridonov, S. , Stock, P. , Sturhan, D. , Waturu, C. and Yoshida, M. 1997. Biosystematics of entomopathogenic nematodes: current status, protocols and definitions. Journal of Helminthology 71:271–298, doi: 10.1017/S0022149X00016096.
  23. Jaffuel, G. , Blanco-Pérez, R. , Hug, A. -S. , Chiriboga, X. , Meuli, R. G. , Mascher, F. , Turlings, T. C. J. and Campos-Herrera, R. 2018. The evaluation of entomopathogenic nematode soil food web assemblages across Switzerland reveals major differences among agricultural, grassland and forest ecosystems. Agriculture, Ecosystems & Environment 262:48–57, doi: 10.1016/j.agee.2018.04.008.
  24. Journey, A. M. and Ostlie, K. R. 2000. Biological Control of the Western Corn Rootworm (Coleoptera: Chrysomelidae) Using the Entomopathogenic Nematode, Steinernema carpocapsae . Environmental Entomology 29:822–831, doi: 10.1603/0046-225X-29.4.822.
  25. Kaplan, E. L. and Meier, P. 1958. Nonparametric Estimation from Incomplete Observations. Journal of the American Statistical Association 53:457–481, doi: 10.2307/2281868.
  26. Kazimierczak, W. , Skrzypek, H. , Sajnaga, E. , Skowronek, M. , Waśko, A. and Kraft, A. 2017. Strains of Photorhabdus spp. associated with polish Heterorhabditis isolates: their molecular and phenotypic characterization and symbiont exchange. Archives of Microbiology 199:979–989, doi: 10.1007/s00203-017-1368-z.
  27. Kimura, M. 1980. A simple method for estimating evolutionary rate of base substitutions through comparative studies of nucleotide sequences. Journal of Molecular Evolution 16:111–120, doi: 10.1007/BF01731581.
  28. Kumar, S. , Stecher, G. , Li, M. , Knyaz, C. and Tamura, K. 2018. MEGA X: molecular evolutionary genetics analysis across computing platforms. Molecular Biology and Evolution 35:1547–1549, doi: 10.1093/molbev/msy096.
  29. Labaude, S. and Griffin, C. T. 2018. Transmission success of entomopathogenic nematodes used in pest control. Insects 9:1–20, doi: 10.3390/insects9020072.
  30. Lacey, L. A. and Georgis, R. 2012. Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. Journal of Nematology 44:218–225.
  31. Liu, J. and Berry, R. E. 1995. Natural distribution of entomopathogenic nematodes (Rhabditida: Heterorhabditidae and Steinernematidae) in Oregon soils. Environmental Entomology 24:159–163, doi: 10.1093/ee/24.1.159.
  32. Londoño, M. E. , Kondo, D. T. , Carabalí, A. , Varón, E. and Caicedo, A. M. 2014. “Insectos y Ácaros”, In Bernal, J. , Díaz, C. , Osorio, C. , Tamayo, A. , Osorio, W. , Córdoba, O. , Londoño, M. E. , Kondo, D. T. , Carabalí, A. , Varón, E. , Caicedo, A. M. , Tamayo, P. J. , Sandoval, A. , Forero, F. , García, J. and Londoño, M. (Eds), Actualización Tecnológica y Buenas Práctica Agrícolas (BPA) en el Cultivo de Aguacate Corpoica, Medellín, pp. 227–284.
  33. López-Llano, R. A. and Soto-Giraldo, A. 2016. Aislamiento de nematodos entomopatógenos nativos en cultivos de caña panelera y pruebas de patogenicidad sobre Diatraea saccharalis (Lepidoptera: Crambidae). Boletín Científico. Centro de Museos. Museo de Historia Natural 20:114–123.
  34. López-Núñez, J. C. , Cano, L. , Góngora-Botero, C. E. and Stock, S. P. 2007. Diversity and evolutionary relationships of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) from the Central Andean region of Colombia. Nematology 9:333–341, doi: 10.1163/156854107781351972.
  35. López-Núñez, J. C. , Plichta, K. , Góngora-Botero, C. E. and Stock, S. P. 2008. A new entomopathogenic nematode, Steinernema colombiense n. sp. (Nematoda: Steinernematidae), from Colombia. Nematology 10:561–574, doi: 10.1163/156854108784513879.
  36. Manachini, B. , Schillaci, D. and Arizza, V. 2013. Biological Responses of Rhynchophorus ferrugineus (Coleoptera: Curculionidae) to Steinernema carpocapsae (Nematoda: Steinernematidae). Journal of Economic Entomology 106:1582–1589, doi: 10.1603/EC13031.
  37. Manton, J. L. , Hollingsworth, R. G. and Cabos, R. Y. M. 2012. Potential of Steinernema carpocapsae (Rhabditida: Steinernematidae) Against Hypothenemus hampei (Coleoptera: Curculionidae) in Hawaii. Florida Entomologist 95:1194–1197, doi: 10.1653/024.095.0452.
  38. Melo, E. L. , Ortega, C. A. , Susurluk, A. , Gaigl, A. and Bellotti, A. C. 2009. Poblaciones nativas de nematodos entomopatógenos (Rhabditida) en cuatro departamentos de Colombia. Revista Colombiana de Entomología 35:28–33.
  39. Morise, H. , Miyazaki, E. , Yoshimitsu, S. and Eki, T. 2012. Profiling nematode communities in unmanaged flowerbed and agricultural field soils in Japan by DNA barcode sequencing. PLoS ONE 7:1–10, doi: 10.1371/journal.pone.0051785.
  40. Mráček, Z. , Bečvář, S. , Kindlmann, P. and Jersáková, J. 2005. Habitat preference for entomopathogenic nematodes, their insect hosts and new faunistic records for the Czech Republic. Biological Control 34:27–37, doi: 10.1016/j.biocontrol.2005.03.023.
  41. Nei, M. and Kumar, S. 2000. Molecular Evolution and Phylogenetics. Oxford: Oxford University Press.
  42. Nguyen, K. B. , Malan, A. P. and Gozel, U. 2006. Steinernema khoisanae n. sp (Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa. Nematology 8:157–175, doi: 10.1163/156854106777998728.
  43. Orozco, R. A. , Lee, M. M. and Stock, S. P. 2014. Soil sampling and isolation of entomopathogenic nematodes (Steinernematidae, Heterorhabditidae). Journal of Visualized Experiments 89:1–8, doi: 10.3791/52083.
  44. Parker, J. , Miles, C. , Murray, T. and Snyder, W. 2012. Organic management of flea beetles. Washington State University PNW 640:1–9.
  45. Peterson, M. E. , Daniel, R. M. , Danson, M. J. and Eisenthal, R. 2007. The dependence of enzyme activity on temperature: determination and validation of parameters. Biochemical Journal 402:331–337, doi: 10.1042/BJ20061143.
  46. Poinar, G. O. 1972. Nematodes as facultative parasites of insects. Annual Review of Entomology 17:103–122.
  47. Ponomarova, O. and Patil, K. R. 2015. Metabolic interactions in microbial communities: untangling the Gordian knot. Current Opinion in Microbiology 27:37–44, doi: 10.1016/j.mib.2015.06.014.
  48. Rizvi, P. Q. , Ahmad, S. K. , Choudhury, A. and Ali, A. 2012. “Biopesticides in ecologically-based integrated pest management”, In Abrol, D. P. and Shankar, U. (Ed.), Integrated pest management principles and practice. London: CAB International, pp. 133–161.
  49. San-Blas, E. , Pembroke, B. and Gowen, S. R. 2012. Scavenging and infection of different hosts by Steinernema carpocapsae . Nematropica 42:123–130.
  50. San-Blas, E. , Campos-Herrera, R. , Dolinski, C. , Monteiro, C. , Andaló, V. , Leite, L. G. , Rodríguez, M. G. , Morales-Montero, P. , Sáenz-Aponte, A. , Cedano, C. , López-Nuñez, J. C. , Del Valle, E. , Doucet, M. , Lax, P. , Navarro, P. D. , Báez, F. , Llumiquinga, P. , Ruiz-Vega, J. , Guerra-Moreno, A. and Stock, S. P. 2019. Entomopathogenic nematology in Latin America: a brief history, current research and future prospects. Journal of Invertebrate Pathology 165:22–45, doi: 10.1016/j.jip.2019.03.010.
  51. Shapiro-Ilan, D. I. , Han, R. and Qiu, X. 2014. “Production of entomopathogenic nematodes”, In Morales-Ramos, J. A. , Rojas, M. G. and Shapiro-Ilan, D. I. (Eds), Mass production of beneficial organisms: invertebrates and entomopathogens. Waltham: Academic Press, pp. 321–355.
  52. Sneath, P. H. A. and Sokal, R. R. 1973. Numerical taxonomy: the principles and practice of numerical classification. San Francisco, CA: W. H. Freeman.
  53. Stock, S. P. 2015. “Diversity, biology and evolutionary relationships”, In Campos-Herrera, R. (Ed.), Nematode pathogenesis of insects and other pests: ecology and applied technologies for sustainable plant and crop protection. Neuchâtel, Cham: Springer, pp. 3–27.
  54. Stock, S. P. , Campbell, J. F. and Nadler, S. A. 2001. Phylogeny of Steinernema Travassos, 1927 (Cephalobina: Steinernematidae) inferred from ribosomal DNA sequences and morphological characters. Journal of Parasitology 87:877–889, doi: 10.1645/0022-3395(2001)087[0877: POSTCS]2.0.CO;2.
  55. Stuart, R. J. , Barbercheck, M. E. and Grewal, P. S. 2015. “Entomopathogenic Nematodes in the Soil Environment: Distributions, Interactions and the Influence of Biotic and Abiotic Factors”, In Campos-Herrera, R. (Ed.), Nematode Pathogenesis of insects and other pests: ecology and applied technologies for sustainable plant and crop protection Springer, Neuchâtel, Cham, 97–137.
  56. Stuart, R. J. , Barbercheck, M. E. , Grewal, P. S. , Taylor, R. A. J. and Hoy, C. W. 2006. Population biology of entomopathogenic nematodes: Concepts, issues, and models. Biological Control 38:80–102.
  57. Tailliez, P. , Pagès, S. , Ginibre, N. and Boemare, N. 2006. New insight into diversity in the genus Xenorhabdus, including the description of ten novel species. International Journal of Systematic and Evolutionary Microbiology 56:2805–2818, doi: 10.1099/ijs.0.64287-0.
  58. Tailliez, P. , Laroui, C. , Ginibre, N. , Paule, A. , Pagès, S. and Boemare, N. 2010. Phylogeny of Photorhabdus and Xenorhabdus based on universally conserved protein-coding sequences and implications for the taxonomy of these two genera. Proposal of new taxa: X. vietnamensis sp. nov., P. luminescens subsp. caribbeanensis subsp. nov., P. luminescens subsp. hainanensis subsp. nov., P. temperata subsp. khanii subsp. nov., P. temperata subsp. tasmaniensis subsp. nov., and the reclassification of P. luminescens subsp. thracensis as P. temperata subsp. thracensis comb. nov. International Journal of Systematic and Evolutionary Microbiology 60:1921–1937, doi: 10.1099/ijs.0.014308-0.
  59. Tamura, K. 1992. Estimation of the number of nucleotide substitutions when there are strong transition-transversion and G + C-content biases. Molecular Biology and Evolution 9:678–687, doi: 10.1093/oxfordjournals.molbev.a040752.
  60. Tamura, K. and Nei, M. 1993. Estimation of the number of nucleotide substitutions in the control region of mitochondrial DNA in humans and chimpanzees. Molecular Biology and Evolution 10:512–526, doi: 10.1093/oxfordjournals.molbev.a040023.
  61. Van Zyl, C. and Malan, A. P. 2014. The role of entomopathogenic nematodes as biological control agents of insect pests, with emphasis on the history of their mass culturing and in vivo production. African Entomology 22:235–249, doi: 10.4001/003.022.0222.
  62. Varón de Agudelo, F. and Castillo, G. 2001. Seminario – Taller sobre identificación de nematodos de importancia agrícola ICA and Asociación Colombiana de Fitopatología (Ascolfi), Palmira.
  63. Vrain, T. C. , Wakarchuk, D. A. , Lévesque, A. C. and Hamilton, R. I. 1992. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundamental and Applied Nematology 15:563–573.
  64. White, G. F. 1927. A method for obtaining infective nematode larvae from cultures. Science 66:302–303, doi: 10.1126/science.66.1709.302-a.
  65. Yan, X. , Waweru, B. , Qiu, X. , Hategekimana, A. , Kajuga, J. , Li, H. , Edgington, S. , Umulisa, C. , Han, R. and Toepfer, S. 2016. New entomopathogenic nematodes from semi-natural and small-holder farming habitats of Rwanda. Biocontrol Science and Technology 26:820–834, doi: 10.1080/09583157.2016.1159658.
  66. Yoon, S. H. , Ha, S. M. , Kwon, S. , Lim, J. , Kim, Y. , Seo, H. and Chun, J. 2017. Introducing EzBioCloud: a taxonomically united database of 16S rRNA and whole genome assemblies. International Journal of Systematic and Evolutionary Microbiology 67:1613–1617, doi: 10.1099/ijsem.0.001755.

EXTRA FILES

COMMENTS