Translator Disclaimer
1 August 2013 Inter- and Intraspecific Identification of the Screwworm, Cochliomyia hominivorax, Using Random Amplified Polymorphic DNA-Polymerase Chain Reaction
Steven R. Skoda, James L. Figarola, Saowaluck Pornkulwat, John E. Foster
Author Affiliations +
Abstract

The screwworm, Cochliomyia hominivorax (Coquerel) (Diptera: Calliphoridae), is one of the most devastating arthropod pests of livestock in the Western Hemisphere. Early instars are very difficult to distinguish morphologically from several closely related blow fly species. Random amplified polymorphic DNA-polymerase chain reaction (RAPD-PCR) markers were developed for identifying C. hominivorax from other wound inhabiting species. Forty decameric primers were screened; nine showed clear reproducible RAPD profiles suitable for distinguishing all life stages of C. hominivorax from 7 other species, including C. macellaria (Fabricius). The results from RAPD-PCR with field-collected samples of unknown first instars agreed with morphological identification that the samples were not C. hominivorax. Three different primers showed DNA polymorphisms (intraspecific) for samples originating from Mexico, Costa Rica, Panama, Jamaica, and Brazil. Therefore, RAPD-PCR may be useful for determining the geographic origin of C. hominivorax samples. Comparing products from these primers, used with known and unknown screwworm samples from an outbreak in Mexico, clearly showed that the outbreak did not originate from the mass rearing facility. Accurate identification of suspected C. hominivorax samples is possible using RAPD-PCR. Further development to identify the geographic origin of samples would benefit the ongoing surveillance programs against C. hominivorax and the decision process during suspected outbreaks of this important pest.

Introduction

The screwworm, Cochliomyia hominivorax (Coquerel) (Diptera: Calliphoridae), is an obligate parasite of warm-blooded animals. Larvae feed on living tissues, causing a condition called myiasis (James 1947). C. hominivorax was one of the most devastating pests of livestock in North America before successful eradication using the sterile insect technique (Vargas-Terán et al. 2005). Screwworms remain a threat to humans and domestic animals in currently endemic regions of South America and the Caribbean. Introduction and subsequent eradication of C. hominivorax to new regions or to previously eradicated regions are well documented (Spradberry 1994). There is an ongoing threat of introduction to the U.S., other regions where C. hominivorax has been eradicated, and areas where it is not endemic.

Effective monitoring, surveillance, and quarantine measures against C. hominivorax depend on accurate and timely identification. However the morphologically similar secondary screwworm, Cochliomyia macellaria (Fabricius), co-exists with C. hominivorax, resulting in numerous misidentifications (particularly for first and second instars) using traditional morphological methods. Other flies in the families Calliphoridae and Sarcophagidae are attracted to necrotic wounds of animals (Hall and Wall 1995), further confounding the difficulties of correct identification, especially for the early instars. Additionally, the possibility of sabotage or fraud exists for any samples suspected to be C. hominivorax but in question (e.g. 1992–1993 Mexican outbreak), making it difficult for program managers to respond appropriately. Consequently, emergency releases of sterile flies have been made, costing millions of dollars, even on samples that proved negative on later analysis (Anonymous 1990). Thus, identification methods that are accurate, timely, and potentially capable of indicating the origins of C. hominivorax samples would be an important addition to the current eradication and barrier maintenance program against C. hominivorax.

Techniques in molecular biology have provided powerful tools for discriminating species and populations using molecular markers (Loxdale and Lushai 1998; Semagn et al. 2006; Pereira et al. 2008; Samarakoon et al. 2012), including applications with screwworms (Azeredo-Espin and Lessinger 2006; Alamalakala et al. 2009). One technique useful for inter- and intraspecific identification is the random amplified polymorphic DNApolymerase chain reaction (RAPD-PCR) (Semagn et al. 2006). RAPD-PCR utilizes short, synthetic oligonucleotides of random sequences as a single primer that is able to anneal and prime at multiple locations throughout the genome of an organism; a spectrum of amplification products are pro- duced that are characteristics of the template DNA (Welsh and McClelland 1990; Williams et al. 1990). The presence and absence of a specific PCR product is diagnostic for the oligonucleotide-binding sites on genomic DNA (Williams et al. 1995) and, therefore, can serve as useful molecular markers for taxonomic and population genetic studies (Semagn et al. 2006; Feng et al. 2009). Compared with other DNA-based methods, the advantages of RAPD-PCR include the quick speed and ease with which results can be obtained, relatively low cost, small DNA sample requirements, and the ability to identify hundreds of new markers in a short time (Hadrys et al. 1992; Semagn et al. 2006). Additionally, no preliminary knowledge of the subject genome is necessary, therefore eliminating the requirements for isolation of cloned DNA probes, preparation of filters for hybridization, and nucleotide sequencing (Jain et al. 2010). Thus, RAPD-PCR can be performed in a moderately equipped laboratory for most applications.

RAPD-PCR has been used to identify cryptic, sibling, and related species of mosquitoes (Ballinger-Crabtree et al. 1992; Wilkerson et al. 1993; Sucharit and Komalamisra 1997), honey bees (Suazo et al. 1998), black flies (Duncan et al. 2004), and screwworms (Skoda et al. 2002). Molecular markers generated by RAPD-PCR were useful in determining the geographic origins of a weevil (Williams et al. 1994), stored product moth (Dowdy and McGaughey 1996), gypsy moth (Schreiber et al. 1997), and fruit flies (Reyes and Onchado 1998), and in estimating gene flow and genetic variability in C. hominivorax (InfanteMalachias et al. 1999; Azeredo-Espin and Lessinger 2006) as well as several species of medically-important mosquitoes (de Sousa et al. 2001; Gonzáles et al. 2007; Hiragi et al. 2009). Additionally, inter- and intraspecific differentiation by RAPD-PCR has been applied to the Mediterranean fruit fly (Sonvico et al. 1996), triatomine bugs (Garcia et al. 1998), and horn flies (Castiglioni et al. 2005), with the capability of identifying biotypes or ecotypes/ecoraces of various insect species (Guirao et al. 1997; Pornkulwat et al. 1998; Saha and Kundu 2006).

Here, our objective was to further develop RAPD-PCR for accurate and timely identification of C. hominivorax from other wound inhabiting flies, and to gain insight into the potential of RAPD-PCR to discriminate geographic origin of screwworm samples.

Methods and Materials

Insect specimens

Field collected samples from Brazil (adults and third instars) and the Mexico outbreak (all third instars) were maintained at -70° C until needed. Field collected larvae from a suspected outbreak in Nicaragua (first and early second instars) were initially stored in 95% ethanol; on arrival to our facility, these samples were placed at -20° C. All other C. hominivorax were obtained from laboratory colonies, 9 strains in all, maintained at the USDA-ARS Midwest Livestock Insects Research Unit-Biosecure Screwworm Rearing Laboratory in Lincoln, Nebraska, USA (Table 1). Colonies of C. macellaria, Chrysomya rufifacies (Macquart), Calliphora vicina (Robineau-Desvoidy), Phormia regina Meigen, Lucilia sericata Meigen, Sarcophaga sp. Meigen (Sarcophagidae), and Musca domestica L. (Muscidae), were established from flies collected at blood and liver baited traps outside the laboratory (Table 1). All samples of life stages from colony flies were frozen and stored at -70° C prior to use. Additional samples of C. macellaria adults, collected from 3 locations in Jamaica in 1998, were maintained in 95% ethanol (Table 1). Samples from Mexico, a confirmed outbreak of suspicious origin, included: 1) larvae of known origin, collected and labeled as from the mass production colony, for comparison to unknowns, and 2) unknown samples (both field collected and from the mass production colony); origins of unknowns were not revealed to the authors (a blind test).

Table 1.

Code, origin, and year of collection of specimens of Cochliomyia hominivorax and related flies used for RAPD-PCR.

t01_01.gif

DNA extraction

All chemicals for DNA extraction were from Sigma-Aldrich ( www.sigmaaldrich.com) unless otherwise noted. DNA was extracted from single individuals using only head capsules in the case of the older larvae and pupae, and heads or legs of newly-emerged adults, to minimize contamination and preserve taxonomic value of specimens. For eggs and first instars (< 24 hr old), ∼10 mg and 5 individuals, respectively, were used per DNA sample. The samples were ground in 1.5 mL microcentrifuge tubes, and total genomic DNA from individual samples was extracted by a modified cetyl ammonium bromide method (Rogers and Bendich 1985) using 400 µL of cetyl ammonium bromide buffer (100 mM Tris-HCl, 1.4 M NaCl, 0.02 M EDTA, 2% cetyl ammonium bromide, and 0.2% &Bgrlmercapto ethanol added just before use) and incubating the samples with 10 µL of Proteinase K (20 mg/mL) and 5 µL of RNAse (50 mg/mL) at 55° C and 37° C for 1 and 2 hr respectively. Additionally, phenol-chloroform extraction was performed once, after which DNA was precipitated with ice-cold 95% ethanol. After precipitation in 400 µL of chilled isopropanol (-20° C for > 2 hr) followed by centrifugation for 30 min in a cooled microcentrifuge (4° C; 14,000 rpm), supernatant was discarded, and DNA pellets were resuspended with 50 µL of Tris-EDTA buffer. The DNA concentration was estimated by running an aliquot in 1% agarose gel (in 1x TBE buffer) with a known concentration of Lambda Hind III marker (Gibco-BRL, Invitrogen,  www.invitrogen.com). Aliquots of the resuspended pellet were diluted 5–20 fold (depending on the concentration) to yield a final DNA concentration of ∼10 ng/µL for use in RAPD-PCR.

Polymerase chain reaction

PCRs were performed using the Perkin-Elmer GeneAmp® PCR 9600 thermocycler and reagents ( www.perkinelmer.com). The RAPD protocol described here was adapted from Pornkulwat (1998) using an optimum template concentration of ∼10 ng. Reaction mixtures of 25 µL total volume per tube contained 12.2 µL of sterile distilled water; 1 µL of 1% Nonidet P-40; 2.5 µL of 10x Stoffel buffer; 4 µL of 25 mM MgCl2; 0.75 µL, each of 10 mM dCTP, dATP, dGTP, and dTTP; 1 µL of 10 pmole 10-mer primer (Operon Technologies, Qiagen,  www.qiagen.com); 1 µL of diluted DNA from individual samples; and 0.3 µL of AmpliTaq® DNA Polymerase Stoffel Fragment (Invitrogen). A master mix of all the PCR components, except the template DNA, was made, aliquoted to the appropriate tubes, and then the DNA sample was added. Negative controls included all reaction components, except template DNA, which was replaced with an equal volume of sterile distilled water. Forty decameric primers were used for initial screening (OPA-7, OPA-12, OPA-16, OPB-1, OPB-10, OPB-13, OPB-17, OPD-16, OPD-20, OPE-1 to OPE-20, OPG-5, OPG-6, OPG9, OPG-10, OPG-1, OPI-10, OPI-11, OPI-15, OPI-16, OPJ-8, and OPJ-11).

The following temperature conditions were used: 95° C for 1 min, 94° C for 1 min, 36° C for 30 sec, and 72° C for 1 min (10 cycles); 94° C for 10 sec, 35° C for 30 sec, and 72° C for 30 sec (30 cycles); and a 72° C extension step for 5 min. Eight µL, of RAPD products for individual samples were loaded and electrophoresed in 1.5% agarose gel (in 1x TBE buffer) with DNA molecular size standards (Gibco-BRL) and a negative control at 60 V for about 2 hours. After electrophoresis, gels were stained with ethidium bromide for 15 min and photographed over a UV transilluminator.

Data Analyses

For screening of species-discriminating markers, DNA was extracted from 10 individual adult flies per strain and species and 3 individuals per immature stage. Eight adult flies per population of C. hominivorax were used individually for screening of intraspecific markers. Primers that generated too many bands were eliminated, and only those that produced clear, distinct, and reproducible bands in all life stages were considered. Two replicates, using the same number of individuals as in screening, were done each for interand intraspecific analyses. At least 3 PCR repetitions per DNA sample were performed, and 2 batches of primers were used to ensure reproducibility of results. Photographs of agarose gels were scanned by HP Scan Jet IIc (Hewett-Packard,  www.hp.com), and the sizes of RAPD generated bands were estimated using the DNA ProRLFP® program (DNA ProScan, Nashville, TN). In all RAPD numerical analyses, band size tolerance (the percentage a band can deviate on either side of the size value and still be considered a match) was set at 1–3%.

Genetic distances were calculated, using all scorable bands (presence/absence) from all selected RAPD primers, with the procedure of Nei and Li (1979) using PAUP® (Swofford 2002). Bootstrapping (1000 replicates) was used to determine support for dendrograms that were derived by the unweighted pairgroup method using arithmetic averages using PAUP® (Swofford 2002).

Results

Species discrimination

PCR amplification of DNA from C. hominivorax and related flies by random primers resulted in the production of discrete banding profiles for all primers tested. Primers were selected based on patterns that generated consistent and reproducible diagnostic results for C. hominivorax. To identify a species-specific primer, some or a combination of amplified DNA fragments must be unique to C. hominivorax, and should be present in all individuals and life stages of the species. Of the 40 decameric primers initially screened, 9 gave RAPD patterns that distinguished the 10 populations of screwworms from the other fly species (Table 2) by a single diagnostic band (Figure 1a) or a combination of bands (Figure 1b). Few co-migrating bands were shared across the different fly species, demonstrating that screwworm samples can be readily identified on the basis of their RAPD banding patterns with any one of the nine primers. Additionally, cluster analyses using all scorable RAPD bands for each of the 9 primers correctly grouped and separated all 10 C. hominivorax populations from the other fly species; the screwworm ‘fork’ exhibited 98% bootstrap support (Figure 2). However, the 5 geographic populations (Costa Rica, Mexico, Panama, Jamaica, and Brazil) of C. hominivorax could not be differentiated using these 9 primers.

Table 2.

Nine species-specific primers, their sequences, and their respective RAPD markers diagnostic of Cochliomyia hominivorax.

t02_01.gif

Figure 1.

Representative RAPD profiles for individuals of Cochliomyia hominivorax and other fly species with primer (A) OPA-12 and (B) OPE-14. C. hominivorax are in lanes: (1) CR92 (Costa Rica), (2) CECH (Mexico), (3) CN (Mexico), (4) LH (Mexico), (5) LI (Mexico), (6) RL (Mexico), (7) PA34 (Mexico), (8) P95 (Panama), (9) J98 (Jamaica), (10) BRZ (Brazil); other species are (11) CMN (C. macellaria), (12) CMJ (C. macellaria), (13) CF (Chrysomyia rufifacies), (14) CP (Calliphora vicina), (15) PR (Phormia regina), (16) PS (Lucilia sericata), (17) SR (Sarcophaga sp.), (18) MD (Musca domestica), (19) control. Lane M contains 100 bp ladder markers. High quality figures are available online.

f01_01.jpg

The diagnostic RAPD bands or banding patterns produced by these speciesdiscriminating primers were consistently generated in all life stages of the fly. Despite some variations from individuals within a species and stage, the identified RAPD diagnostic markers were present in all life stages of C. hominivorax, and the banding patterns clearly differed with those of immature C. macellaria (Figure 3).

Figure 2.

Dendogram of the relationships between 10 populations of C. hominivorax CR92 (Costa Rica), CECH (Mexico), CN (Mexico), LH (Mexico), LI (Mexico), RL (Mexico), PA34 (Mexico), P95 (Panama), J98 (Jamaica), BRZ (Brazil) and other closely related myiasigenic flies CMN (C. macellaria), CMJ (C. macellaria), CF (Chrysomyia rufifacies), CP (Calliphora vicina), PR (Phormia regina), PS (Lucilia sericata), SR (Sarcophaga sp.), MD (Musca domestica) using all gels and all scorable RAPD bands amplified by primers OPA-12, OPA-16, OPB-1, OPE-3 and OPE-16. Similarity values calculated by Nei-Li's coefficient; UPGMA clustering; only bootstap values > 50% reported. High quality figures are available online.

f02_01.jpg

Five larvae of unknown first instars collected from cattle wounds in Nicaragua were analyzed together with the same number of larvae of C. hominivorax and the other fly species. The unknown insect was not C. hominivorax (Figure 4a). Morphological identification placed the sample as Phaenicia (=Lucilia). However, RAPD patterns for L. sericata samples from Nebraska did not match (Figure 4a). Cluster analyses using the unweighted pairgroup method indicated that the unknown was closer to C. hominivorax than L. sericata or the other species (Figure 4b).

Figure 3.

Representative gel comparing RAPD profiles between individuals of different stages of Cochliomyia hominivorax (lanes 1–7) with C. macellaria (lanes 10–15) using primer OPB-1. Lanes: (1–2) (CR92; Costa Rica) eggs, (3–4) first instar, (5) third instar, (6) pupa, (7) adult, (8–9) C. macellaria (CMN; Nebraska) eggs, (10–11) first instar, (12) third instar, (13–14) pupa, (15) adult. Lane M contains 100 bp ladder markers. High quality figures are available online.

f03_01.jpg

Table 3.

Potential intraspecific RAPD primers for identification of Cochliomyia hominivorax populations.

t03_01.gif

Variation among C. hominivorax populations

RAPD profiles of individual adults of wild type populations of C. hominivorax from Costa Rica, Mexico, Jamaica, Panama, and Brazil were compared with 3 primers initially identified for potential strain-specific markers. At least 15 potential intraspecific RAPD markers were generated from these 3 primers (Table 3). Discrimination of C. hominivorax populations based on the presence/absence of some RAPD-PCR products was possible. For example, using primer OPJ-8, the presence of ∼600, 540, 420, and 365 bp products distinguished the C. hominivorax populations from Brazil, while a combination of 420 and 365 bp was diagnostic of Jamaican populations (Figure 5a; Table 3). OPG-10 produced several bands, of which the absence of ∼770 bp and 480 bp fragments was diagnostic of Brazilian and Mexican C. hominivorax populations, respectively (Figure 5b; Table 3). In contrast, OPE-12 produced at least 7 bands that could potentially differentiate all the C. hominivorax populations that were analyzed (Table 3).

Figure 4.

(A) Representative gel comparing RAPD profiles of individual unknown first instars from Nicaragua with first instars of Cochliomyia hominivorax in lanes (1) CR92 (Costa Rica) and (2) J98 (Jamaica) and other flies using primer OPA-12 (3–4) CMN (C. macellaria), (5–6) UNK, (7) CF , (Chrysomyia rufifacies) (8) CP (Calliphora vicina), (9) PR (Phormia regina), (10) PS (Lucilia sericata), (11) SR (Sarcophaga sp.). Lanes M1 and M2 contain 100 bp and 50 bp ladder markers, respectively. (B) Dendogram of the relationships between unknowns (UNK) from Nicaragua with screwworms and other flies from all gels and all scorable RAPD bands amplified from primer OPA-12, OPB-1, OPE-3 and OPE-14. Abbreviations same as for A above. Similarity values calculated by Nei-Li's coefficient; unweighted pair-group method using arithmetic averages clustering; only bootstap values > 50% reported. High quality figures are available online.

f04_01.jpg

Figure 5.

Representative gels of RAPD profiles with primer OPJ-8 (A), and OPG-10 (B) using individuals of Cochliomyia hominivorax originating from 5 different locations. Lanes: (1–4) P95 (Panama), (5–8) J98 (Jamaica), (9–12) PA-34 (Mexico), (13–16) BRZ (Brazil), (17–20) CR92 (Costa Rica). Lane M contains 100 bp ladder markers. High quality figures are available online.

f05_01.jpg

Numerical analyses of RAPD-PCR products from the 3 primers (OPG-10, OPE-12, and OPJ-8) resulted in clear-cut population grouping of each fly sample according to its geographical origin (Figure 6). Analysis of RAPD patterns showed strong support for each population ‘branch.’ Overall cluster analysis using all scorable RAPD bands generated from the 3 primers suggests that flies from Brazil form an independent group separate from the other 4 populations. C. hominivorax from Costa Rica were more closely associated with the Mexican populations while Panama and Jamaica C. hominivorax clustered together (Figure 6).

The same 3 primers used to detect intraspecific variation, along with OPA-12 and OPI-10, were used with samples from the Mexico outbreak. The results, including photographs of gels and dendrograms generated with unweighted pair-group analysis, were sent to the sample curator (who designated the field vs. colony unknowns). The pattern of RAPD amplification products for the field-collected outbreak samples did not match that from the mass production facility, as evidenced from gel images and dendrograms (data not shown); therefore, sabotage from the facility for mass production of screwworms, as originally feared/suspected, was ruled out.

Figure 6.

Dendogram of the relationships between Cochliomyia hominivorax from 5 geographic locations using all gels and all scorable RAPD bands amplified by 3 primers (OPE-12, OPG-10, OPJ-8); P95 = Panama, J98 = Jamaica, MX = Mexico, CR = Costa Rica, BR = Brazil. Similarity values calculated by Nei-Li's coefficient; unweighted pair-group method using arithmetic averages clustering; only bootstap values > 50% reported. High quality figures are available online.

f06_01.jpg

Discussion

Several studies have shown that the number of loci amplified with RAPD primers depends on factors such as reagents and reaction conditions, sample conditions, and DNA quality and extraction methods (Williams et al. 1990; 1995; Black 1993; Ellsworth et al. 1993; Kernodle et al. 1993; MacPherson et al. 1993; Meunier and Grimont 1993; Micheli et al. 1994; Gallego and Martinez 1997; Khandka et al. 1997). Variations were minimized and amplifications of artifacts in the RAPD profiles were prevented by using heads and legs of individual frozen flies, by using similar amounts of template DNA for PCR runs, by running a negative control in each reaction, and by using 2 batches of each potential primer and AmpliTaq® DNA Polymerase Stoffel Fragment during the replication process. Stoffel Fragment is a 61 kDa modified form of recombinant AmpliTaq® DNA Polymerase from which the N-terminal 289 amino acids have been deleted to increase stringency at lower ionic strength and reduce misextension. The use of Stoffel Fragment is very important for reproducibility of results, a negative attribute often attributed to RAPDs (Pereira et al. 2008). Additionally, amplification conditions that previously resulted in reproducible RAPD profiles were used (Pornkulwat 1998; Skoda et al. 2002). Primer selection was based on discrete and reproducible fragments, i.e., those that produced a smaller number of intense, diagnostic bands, as seen on agarose gels stained with ethidium bromide. Reproducible genetic markers were identified from the decameric primers used for species-specific and intraspecific identifications. Sample identification can de done in < 10 hr, starting from DNA extraction to actual visualization of amplified products on agarose gels.

The diagnostic bands for each of the 9 primers suitable for C. hominivorax identification were selected based on their reproducibility and consistency of amplification products in all C. hominivorax life stages. In addition, all diagnostic markers were present in all 10 C. hominivorax populations analyzed in this study. The diagnostic marker can be a single RAPD band (e.g., OPA-12, OPE-13), or a combination of bands (OPB-1, OPE-3, OPE-14, OPE-16, OPG-9). In practice, we recommend that several primers should be used for accurate diagnosis of samples, especially for poorly-preserved specimens. For example, a loss of some marker bands was observed from samples preserved in 70% alcohol for more than 3 months (S. Pornkulwat, personal observation) compared with 95% ethanol, such that identification through amplification products from a single primer may not be reliable. Similar findings have been reported for L. sericata (Stevens and Wall 1995). We further recommend that samples be preserved in 95% ethanol (if alcohol is the only storage medium and for long-term storage) for this RAPDPCR method to be accurate and reliable.

The accuracy of this technique for identifying screwworms was evaluated when unknown Nicaraguan samples of first instars taken from myiasis wounds were analyzed. Although Nicaragua had been declared screwworm-free since 1997, the possibility of reintroduction in this country due to livestock trade and movement is always present. RAPD-PCR of DNA from the unknown showed that the sample was not C. hominivorax. This conclusion was supported by results from morphological identification. The unknown was identified as Lucilia sp. based on available keys, but RAPD banding patterns of the L. sericata collected from Nebraska were different from the unknowns. Interestingly, cluster analyses indicated that the unknowns were closely related to C. hominivorax. Polymorphisms have been shown to exist among populations of L. sericata worldwide (Stevens and Wall 1997), and probably all species of flies from different geographical areas, such that identification of negative samples by RAPD-PCR is impractical, if not impossible at this time, based on the limited genetic markers for these non-screwworm species. Nevertheless, as noted above, all the RAPD diagnostic markers generated from all 9 species-specific primers were always present in all 10 C. hominivorax populations used here; these populations originated from distinctly different geographic regions. Additionally, results from cluster analyses of RAPD banding patterns for each of the 9 species-discriminating primers, strongly supported by bootstrapping analyses, resulted in the correct separation of C. hominivorax samples from non-screwworm specimens (using 1–3% band size tolerance matches). This further indicates the utility of RAPD-PCR for identification and emphasizes the need to expand the molecular genetic database in order to identify which species is present if samples are identified as not being C. hominivorax. The possibility of using some of these RAPD markers (particularly the single RAPD marker from OPA-12 and OPE-13) as species-specific DNA probes for C. hominivorax and their potential for field use should be investigated. Such RAPD-derived probes, sequence characterized amplified regions (SCAR), have been successfully used to identify related bacterial (Argenton et al. 1996; Oakey et al. 1998), protozoan (trypanosome) (Oury et al. 1997), silkworms (Saha and Kundu 2006), and other species.

Although not the main focus, analyses of C. hominivorax samples originating from 5 geographic locations using RAPD-PCR suggested that this technique may be able to discriminate between these populations. Genetic analyses based on pair-wise comparisons of RAPD bands showed that intraspecific genetic variation existed within and among these populations, in general agreement with results from earlier studies (Infante-Malachias et al. 1999; Skoda et al. 2002; Azeredo-Espin and Lessinger 2006). However, all individuals within a population were observed to cluster, suggesting that this technique may be suitable for molecular fingerprinting and possible identification of the geographical origin of a C. hominivorax sample. The analyses using all scorable RAPD bands generated by the 3 primers indicated that flies from Brazil form separate populations different from those of Jamaica, Mexico, and Central America. This finding was also indicated in the dendrogram generated from data of RAPDs using the primers for identifying screwworms from other species. These results differ somewhat from the conclusions made by Taylor et al. (1996), who divided C. hominivorax populations into 3 assemblages (North and Central America, South America, and Jamaica) but were using RFLP of mtDNA and had fewer samples and fewer bands for analysis.

The data presented here provide useful molecular markers only for possible identification of the geographical origins of the C. hominivorax sample. The results are still preliminary and should not be regarded as indicators of the level of genetic variability or relationships among various C. hominivorax populations. It would be necessary to analyze several more C. hominivorax samples from South America and the Caribbean to develop more comprehensive genetic ‘fingerprints’ for later comparisons with suspected C. hominivorax infestation samples. Obviously, much greater diversity might be found if specimens representing the entire geographical range of C. hominivorax are examined. Compared with PCR-RLFP, AFLP, or microsatellites, the ease with which RAPD-PCR can be used increases the number of samples that can be tested, thereby allowing an increased representation of populations in future RAPD analyses. Additionally, the use of 2 or more primers in RAPD reactions (Welsh and McClelland 1991; Micheli et al. 1993) could potentially increase the number of molecular markers that can be generated, thereby increasing the chances of developing RAPD ‘fingerprints’ for all populations and/or strains of C. hominivorax.

In conclusion, RAPD-PCR is a useful tool for interspecific, and promising for intraspecific, identification of C. hominivorax. This technique provides information helpful in the positive identification and monitoring of C. hominivorax in areas where the species is currently endemic, while its use does not demand a sophisticated laboratory. In case of suspected introduction, rapid and reliable identification allows eradication program officials to take appropriate steps to prevent further spread of the infestation (which can then lead to timely, appropriate corrective measures), but does not preclude the further verification of results from other laboratories or using other techniques. Aside from routine identification of field-collected immature samples and the potential for determination of sample origin, future applications of this technique may include checking suspected contamination in screwworm colonies in rearing facilities and assessing loss of genetic variability caused by selection and inbreeding in a colony. Moreover, developing RAPD markers that can distinguish males from females during early life stages would be valuable in the current plan of developing a male-only strain of C. hominivorax flies in mass-production facilities.

Acknowledgements

We thank Valerie Lacky, Leela Alamalakala, and Jessica Jones for their assistance in the laboratory, as well as Mark Meadows and Dr. Dennis Berkebile for colony maintenance. This work was done in cooperation with the Institute of Agriculture and Natural Resources, University of Nebraska, Lincoln, NE. Mention of a proprietary product does not constitute endorsement or recommendation for its use by the USDA. USDA is an equal opportunity provider and employer.

Glossary

Abbreviations:

RAPD-PCR

,

random amplified polymorphic DNA-polymerase chain reaction

References

1.

L Alamalakala , SR Skoda , JE Foster . 2009. Amplified fragment length polymorphism used for inter- and intraspecific differentiation of screwworms (Diptera: Calliphoridae). Bulletin of Entomological Research 99: 139–149. Google Scholar

2.

Anonymous. 1990. Suspected screwworms in Texas. Texas A & M Agricultural Experiment Station, Foreign Animal Disease Report 18: 4. Google Scholar

3.

F Argenton , D DeMas , C Malocco , Valle L Dalla , G Giorgetti , L Colombo . 1996. Use of random DNA amplification to generate specific molecular probes for hybridization tests and PCR-based diagnosis of Yersinia ruckeri. Diseases of Aquatic Organisms 24: 121–127. Google Scholar

4.

AML Azeredo-Espin , AC Lessinger . 2006. Genetic approaches for studying myiasiscausing flies: molecular markers and mitochondrial genomics. Genetica 126: 111–131. Google Scholar

5.

ME Ballinger-Crabtree , WC Black IV , BB Miller . 1992. Use of genetic polymorphism detected by random pylymerase chain reaction (RAPD-PCR) for differentiation of Aedes aegypti subspecies and populations. The American Journal of Tropical Medicine and Hygiene 47: 893–901. Google Scholar

6.

WC Black IV . 1993. PCR with arbitrary primers: approach with care. Insect Molecular Biology 2: 1–6. Google Scholar

7.

L Castiglioni , HEM de Campos Bicudo . 2005 Molecular characterization and relatedness of Haematobia irritons (horn fly) populations, by RAPD-PCR. Genetica 124: 11–21. Google Scholar

8.

GB de Sousa , A Blanco , CN Gardenal . 2001. Genetic relationships among Aedes aegypti (Diptera: Culicidae) populations from Argentina using random amplified polymorphic DNA polymerase chain reaction markers. Journal of Medical Entomology 38: 371–375. Google Scholar

9.

AK Dowdy , WH McGaughey . 1996. Using random amplified polymorphic DNA to differentiate strains of the Indian meal moth (Lepidoptera: Pyralidae). Environmental Entomology 25: 396–400. Google Scholar

10.

GA Duncan , PH Adler , KP Pruess , TO Powers . 2004. Molecular differentiation of two sibling species of the black fly Simulium vittatum (Diptera: Simuliidae) based on random amplified polymorphic DNA. Genome 47: 373–379. Google Scholar

11.

DL Ellsworth , KD Rittenhouse , RL Honeycutt . 1993. Artifactual variation in randomly amplified polymorphic DNA banding patterns. BioTechniques 14: 214–217. Google Scholar

12.

K Feng , S Wang , DJ Hu , FQ Yang , HX Wang , SP Li . 2009. Random amplified polymorphic DNA (RAPD) analysis and the nucleosides assessment of fungal strains isolated from natural Cordyceps sinensis. Journal of Pharmaceutical and Biomedical Analysis 50: 522–526 Google Scholar

13.

FJ Gallego , I Martinez . 1997. Method to improve reliability of random-amplified polymorphic DNA markers. BioTechniques 23: 663–664. Google Scholar

14.

AL Garcia , HJ Carraco , CJ Schofield , JR Stothhard , IA Frame , SAS Valente , MA Miles . 1998. Random amplification of polymorphic DNA as a tool for taxonomic studies of triatomine bugs (Reduviidae: Hemiptera). Journal of Medical Entomology 35: 38–45. Google Scholar

15.

R González , R Wilkerson , MF Suárez , F García , G Gallego , H Cárdenas , CE Posso , MC Duque . 2007. A population genetics study of Anopheles darlingi (Diptera: Culicidae) from Colombia based on random amplified polymorphic DNA-polymerase chain reaction and amplified fragment length polymorphism markers. Memórias do Instituto Oswaldo Cruz 102: 255–262. Google Scholar

16.

P Guirao , F Beitia , JL Cenis . 1997. Biotype determination of Spanish population of Bemisia tabaci (Hemiptera: Aleyrodidae). Bulletin of Entomological Research 87: 587–593. Google Scholar

17.

H Hadrys , M Balick , B Schierwater . 1992. Applications of random amplified polymorphic DNA (RAPD) in molecular ecology. Molecular Ecology 1: 55–63. Google Scholar

18.

M Hall , R Wall . 1995. Myiasis of humans and domestic animals. Advances in Parasitology 35: 257–334. Google Scholar

19.

C Hiragi , K Simões , E Martins , P Queiroz , L Lima , R Monnerat . 2009. Genetic variability in Aedes aegypti (L.) (Diptera: Culicidae) populations using RAPD markers. Neotropical Entomology 38: 542–547. Google Scholar

20.

ME Infante-Malachias , KSC Yotoko , Espin AML de Azeredo . 1999. Random amplified polymorphic DNA of screwworm fly populations (Diptera: Calliphoridae) from southeastern Brazil and northern Argentina. Genome 42: 772–779. Google Scholar

21.

SK Jain , B Neekhra , D Pandey , K Jain . 2010. RAPD marker system in insect study: a review. Indian Journal of Biotechnology 9: 7–12. Google Scholar

22.

MT James . 1947. The flies that cause myiasis in man. USDA Miscellaneous Publication 631. Google Scholar

23.

SP Kernodle , RE Cannon , JG Scandalios . 1993. Concentration of primer and template qualitatively affects products in randomamplified polymorphic DNA PCR. BioTechniques 14: 362–364. Google Scholar

24.

DK Khandka , M Tuna , M Tal , A Nejidat , A Golangoldhisrh . 1997. Variability in the pattern of random amplified polymorphic DNA. Electrophoresis 18: 2852–2856. Google Scholar

25.

HD Loxdale , G Lushai . 1998. Molecular markers in entomology. Bulletin of Entomological Research 88: 577–600. Google Scholar

26.

JM MacPherson , PE Ecktein , GJ Scoles , AA Gajadhar . 1993. Variability of the random amplified polymorphic DNA assay among thermal cyclers, and effects of primer and DNA concentration. Molecular and Cellular Probes 7: 293–299. Google Scholar

27.

JR Meunier , PAD Grimont . 1993. Factors affecting reproducibility of random amplified polymorphic DNA fingerprinting. Research in Microbiology. 144: 373–379. Google Scholar

28.

M Micheli , RR Bova , P Calissano , E D'Ambrossio . 1993. Randomly amplified polymorphic DNA fingerprinting using combinations of oligonucleotide primers. BioTechniques 15: 388–390. Google Scholar

29.

MR Micheli , R Bova , E Pascale , E D'Ambrosio . 1994. Reproducible DNA fingerprinting with the random amplified polymorphic DNA (RAPD) method. Nucleic Acids Research 22: 1921–1922. Google Scholar

30.

M Nei , WH Li . 1979. Mathematical model for studying genetic variation in terms of restriction endonucleases. Proceedings of the National Academy of Sciences USA 76: 5269–5273. Google Scholar

31.

HJ Oakey , JT Ellis , LF Gibson . 1998. The development of random DNA probes specific for Aeromonas salmonicida. Journal of Applied Microbiology 84: 37–46. Google Scholar

32.

B Oury , N Dutrait , B Bastrenta , M Tibayrenc . 1997. Trypanosoma cruzi: evaluation of a RAPD synamorphic fragment as a speciesspecific DNA probe. Journal of Parasitology 83: 52–57. Google Scholar

33.

F Pereira , J Carneiro , A Amorim . 2008. Identification of species with DNA-based technology: current progress and challenges. Recent Patents on DNA and Gene Sequencing 2: 187–199. Google Scholar

34.

S Pornkulwat . 1998. Molecular genetic markers among and within populations of the European corn borer (Lepidoptera: Pyralidae) and screwworms (Diptera: Calliphoridae). Ph.D. Dissertation. University of Nebraska, Lincoln, NE. Google Scholar

35.

S Pornkulwat , SR Skoda , GD Thomas , JE Foster . 1998. Random amplified polymorphic DNA used to identify genetic variation in ecotypes of the European corn borer (Lepidoptera: Pyralidae). Annals of the Entomological Society of America 91: 719–725. Google Scholar

36.

A Reyes , MD Onchado . 1998. Use of molecular markers for detecting the geographical origin of Ceratitis capitata (Diptera: Tephritidae) populations. Annals of the Entomological Society of America 91: 222–227. Google Scholar

37.

SO Rogers , AJ Bendich . 1985. Extraction of DNA from milligram amounts of fresh, herbarium and mummified plant tissues. Plant Molecular Biology 5: 69–76. Google Scholar

38.

M Saha , SC Kundu . 2006. Molecular identification of tropical tasar silkworm (Antheraea mylitta) ecoraces with RAPD and SCAR markers. Biochemical Genetics 44: 75–88. Google Scholar

39.

U Samarakoon , SR Skoda , FP Baxendale , JE Foster . 2012. A molecular key for the identification of blow flies in southeastern Nebraska. Journal of Forensic Sciences DOI:  10.HH/j.1556-4029.2012.02182.x  Google Scholar

40.

DE Schreiber , KJ Garner , JM Slavicek . 1997. Identification of three randomly amplified polymorphic DNA-polymerase chain reaction markers for distinguishing Asian and North American gypsy moths. Annals of the Entomological Society of America 90: 667–674. Google Scholar

41.

K Semagn , Å Bjørnstad MN Ndjiondjop . 2006. An overview of molecular marker methods for plants. African Journal of Biotechnology 5: 2540–2568. Google Scholar

42.

SR Skoda , S Pornkulwat , JE Foster . 2002. Random amplified polymorphic DNA markers for discriminating Cochliomyia hominivorax from C. macellaria (Diptera: Calliphoridae). Bulletin of Entomological Research 92: 89–96. Google Scholar

43.

A Sonvico , F Manso , LA Quesada-Allue . 1996. Discrimination between the immature stages of Ceratitis capitata and Anastrepha fraterculus (Diptera: Tephritidae) populations by random amplified polymorphic DNA polymerase chain reaction. Journal of Economic Entomology 89: 1208–1212. Google Scholar

44.

JP Spradberry . 1994. Screw-worm fly: a tale of two species. Agricultural Zoology Reviews 6: 1–62. Google Scholar

45.

J Stevens , R Wall . 1995. The use of random amplified polymorphic DNA (RAPD) analysis for studies of genetic variation in populations of the blowfly Lucilia sericata (Diptera: Calliphoridae) in southern England. Bulletin of Entomological Research 85: 549–555. Google Scholar

46.

J Stevens , R Wall . 1997. Genetic variation in populations of the blow flies Lucilia cuprina and Lucilia sericata (Diptera: Calliphoridae). Random amplified polymorphic DNA and mitochondrial DNA sequences. Biochemical Systematics and Ecology 25: 81–97. Google Scholar

47.

A Suazo , R McTiernan , HG Hall . 1998. Differences between African and European honey bees (Apis mellifera L.) in random amplified polymorphic DNA (RAPD). Journal of Heredity 89: 32–36. Google Scholar

48.

S Sucharit , N Komalamisra . 1997. Differentiation of Anopheles minimus species complex by RAPD-PCR technique. Journal of the Medical Association of Thailand 80: 598–602. Google Scholar

49.

DL Swofford . 2002. PAUP*, Phylogenetic Analysis Using Parsimony (*and Other Methods), Version 4. Sinauer Associates. Google Scholar

50.

DB Taylor , AL Szalanski , RD Peterson . 1996. Mitochondrial DNA variation in screwworm. Medical and Veterinary Entomology 10: 161–169. Google Scholar

51.

M Vargas-Terán , HC Hofmann , NE Tweddle . 2005. Impact of screwworm eradication programmes using the sterile insect technique. In: VA Dyck , AS Robinson , Editors. Sterile insect technique, principles and practice in area-wide integrated pest management. pp. 629–650. Springer. Google Scholar

52.

J Welsh , M McClelland . 1990. Fingerprinting genomes using PCR with arbitrary primers. Nucleic Acids Research 18: 7213–7218. Google Scholar

53.

J Welsh , M McClelland . 1991. Genomic fingerprinting using arbitrary primed PCR and a matrix of pairwise combinations of primers. Nucleic Acids Research 19: 5275–5280. Google Scholar

54.

RC Wilkerson , TJ Parsons , DG Albright , TA Klein , MJ Braun . 1993. Random amplified polymorphic DNA (RAPD) markers readily distinguish cryptic mosquito species (Diptera: Culicidae: Anopheles). Insect Molecular Biology 1: 205–211. Google Scholar

55.

CL Williams , SL Goldson , DB Baird , DW Bullock . 1994. Geographical origin of an introduced insect pest, Listronotus bonariensis (Kuschel) determined by RAPD analysis. Heredity 72: 412–419. Google Scholar

56.

JGK Williams , AN Kubelik , KJ Livak , JA Rafalski , SV Tingey . 1990. DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Research 18: 6531–6535. Google Scholar

57.

JGK Williams , MK Hanafey , JA Rafalski , SV Tingey . 1995. Genetic analysis using random amplified polymorphic DNA markers. , In: R Wu , Editor. Recombinant DNA methodology II. pp. 849–883. Academic Press. Google Scholar
This is an open access paper. We use the Creative Commons Attribution 3.0 license that permits unrestricted use, provided that the paper is properly attributed.
Steven R. Skoda, James L. Figarola, Saowaluck Pornkulwat, and John E. Foster "Inter- and Intraspecific Identification of the Screwworm, Cochliomyia hominivorax, Using Random Amplified Polymorphic DNA-Polymerase Chain Reaction," Journal of Insect Science 13(76), 1-15, (1 August 2013). https://doi.org/10.1673/031.013.7601
Received: 6 March 2012; Accepted: 1 October 2012; Published: 1 August 2013
JOURNAL ARTICLE
15 PAGES


SHARE
ARTICLE IMPACT
Back to Top