Morphological and Molecular Identification of Globodera pallida Associated with Potato in Idaho

Abstract

The identity of a newly discovered population of pale potato cyst nematode Globodera pallida associated with potato in eastern Idaho was established by morphological and molecular methods. Morphometrics of cysts and second-stage juveniles were generally within the expected ranges for G. pallida with some variations noted. The Idaho population and paratype material from Epworth, Lincolnshire, England, both showed variations in tail shape, with bluntly rounded to finely pointed tail termini. Compared to literature values for the paratypes, second-stage juveniles of the Idaho population had a somewhat shorter mean body length, and cysts had a slightly higher mean distance from the anus to the nearest edge of the fenestra. PCR-RFLP of the rDNA ITS region, sequence-specific multiplex PCR and DNA sequence comparisons all confirmed the identity of the Idaho population as G. pallida. The ITS rDNA sequence of the Idaho isolate was identical to those from York, England, and the Netherlands. Species-specific primers that can positively identify the tobacco cyst nematode Globodera tabacum were also developed, providing a new assay for distinguishing this species from G. pallida and the golden potato cyst nematode Globodera rostochiensis.

Like the golden potato cyst nematode Globodera rostochiensis (Wollenweber, 1923) Behrens, 1975, the pale potato cyst nematode G. pallida (Stone, 1973) Behrens, 1975 is a regulated pathogen of potato (Solanum tuberosum) in the United States and many other countries. Within the Americas, G. pallida has been found in South America, Panama and Newfoundland, Canada (Marks and Brodie, 1998; EPPO, 2004), but had not previously been reported in the U.S. In March 2006, cyst nematodes were discovered in tare soil from a potato processing facility in eastern Idaho. The nematodes were found during a routine survey conducted jointly by the Idaho State Department of Agriculture and the USDA Animal and Plant Health Inspection Service through the Cooperative Agricultural Pest Survey program. Six cysts from the new detection were received by the USDA-ARS Nematology Laboratory for species identification. Subsequent extensive sampling traced the nematode to two fields in northern Bingham County, Idaho (Hafez et al., 2007). Morphological and molecular methods that were used to identify and characterize this population as G. pallida are described herein. The new discovery of G. pallida in Idaho constitutes a significant threat to the U.S. potato industry, which is valued at over $2.6 billion (Agricultural Statistics Board, 2006). The availability of germplasm resistant to G. pallida is currently limited. Thus, the ability to distinguish G. pallida from other cyst nematodes that may be present is critical for the application of rational regulatory decisions aimed at preventing further infestation.

A few females of G. pallida have been experimentally demonstrated to reproduce on tobacco (Nicotiana tabacum) (Parrot and Miller, 1977). Similarly, only a few females of morphologically similar G. tabacum (Lownsbery and Lownsbery, 1954) Behrens, 1975 have been shown to reproduce on potato (Stone and Miller, 1974; Baldwin and Mundo-Ocampo, 1991). Globodera tabacum has been found in the states of Virginia, Connecticut and North Carolina, as well as in Mexico and various countries in Europe and northern Africa (Marché et al., 2001; Syracuse et al., 2004). Most recently, G. tabacum was detected in tobacco from Ontario, Canada, although the subspecies was not reported (Bélair and Miller, 2006). The G. tabacum complex contains three subspecies that in nature parasitize solanaceous weeds (G. tabacum virginiae) and tobacco (G. tabacum tabacum and G. tabacum solanacearum) (Baldwin and Mundo-Ocampo, 1991; Syracuse et al., 2004).

The morphometrics from juveniles or females cannot clearly differentiate G. pallida from the three subspecies of G. tabacum (Baldwin and Mundo-Ocampo, 1991; EPPO, 2004). Neither light microscopy (Stone, 1983; Baldwin and Mundo-Ocampo, 1991; Mota and Eisenback, 1993a–c; EPPO, 2004) nor scanning electron microscopy (Othman et al., 1988) of juveniles or females can clearly distinguish G. pallida from the three subspecies of G. tabacum. While a number of immunological, protein and DNA-based techniques can separate G. rostochiensis from G. pallida (reviewed in Fleming and Powers, 1998 and Ibrahim et al., 2001), only a few have included both European and non-European populations (Subbotin et al., 2000; Grenier et al., 2001) or included related species such as G. tabacum, “G. mexicana” (Campos-Vela, 1967), or G. achilliae (Thiéry and Mugniéry, 1996). With the appearance of G. pallida now confirmed in the U.S. (Hafez et al., 2007), there is an urgent need for new molecular diagnostics capable of differentiating G. pallida from species other than G. rostochiensis that occur in the Americas. A modified multiplex PCR assay that can distinguish G. tabacum from G. pallida and G. rostochiensis is described.

Materials and Methods

Populations: Globodera spp. populations described and discussed are listed in Table 1. Heterodera avenae from Idaho was also included in this investigation because of its occasional occurrence in survey samples and utility as a control for the PCR-RFLP analysis.

Table 1.

List of nematode species and populations used in this study

Morphological Characterization: Juveniles for morphological observations were hatched from cysts that had been sieved from fresh soil and kept in water in Syracuse watch glasses in the laboratory. Juveniles were then fixed in 3% formaldehyde and processed to glycerine by the formalin-glycerine method (Hooper, 1970; Golden, 1990). Cysts were similarly removed from soil samples by sieving, fixed for 12 hr in 3% formaldehyde and processed to glycerine. Photomicrographs of cyst vulval cones and J2 were made with an automatic 35-mm camera attached to a compound microscope fitted with an interference contrast system. Light microscopic images of fixed nematodes were taken on a Leica Wild MPS48 Leitz DMRB compound microscope, and measurements were made with an ocular micrometer on the same microscope. Specimens were then morphologically identified with recent taxonomic keys and a compendium for identification of Globodera spp. (Golden, 1986; Baldwin and Mundo-Ocampo, 1991). All measurements are in micrometers unless otherwise stated. Differential Interference Contrast (DIC) images of live specimens mounted on an agar pad were taken on a Zeiss UltraPhot light microscope, with contrast enhanced in Adobe Photoshop CS v. 8.

Molecular characterization: Nematode juveniles or cysts were mechanically disrupted with an eye-knife or sharp forceps tips in 20 μl nematode extraction buffer as described (Thomas et al., 1997) and stored at −80°C. For preparation of nematode extracts, samples were thawed, an additional 1 μl proteinase K (from 2 mg/ml stock solution) was added, and the tubes were incu bated at 60°C for 60 min, followed by 95°C for 15 min to deactivate the proteinase K. Two or five microliters of the extract was used for each PCR reaction (see be low). Where noted for some PCR reactions, bulk G. rostochiensis genomic DNA was included at 20 ng/ reaction.

The rDNA PCR primers used in this study are listed in Table 2. Amplifications were performed on two or more individuals from each nematode population. Negative controls with water instead of template were included with each experiment. PCR reactions contained 0.2 μM each of primers TW81 (Joyce et al., 1994) and AB28 (Howlett et al., 1992), 5 μl nematode extract, 200 μM dNTP, 1 U Eppendorf Hot Master Taq (Brinkmann, Westbury, NY) and supplied enzyme reaction buffer at 1x in a total volume of 50 μl. Cycling conditions included an initial denaturation step of 95°C for 2 min, followed by 35 cycles of 95°C for 30 sec, 55°C for 30 sec and 72°C for 90 sec, and finished with one cycle at 72°C for 5 min. A 5μl aliquot of each PCR reaction was analyzed by electrophoresis on 1% agarose/Trisacetate-EDTA (TAE), and the rest was saved for subsequent restriction enzyme digestion. Gels were stained with ethidium bromide and visualized by UV illumination. Gel images were captured with a Chemi Imager (Alpha Innotech, San Leandro, CA).

Table 2.

Primers used in this study

Restriction enzymes Alu I, Rsa I and Taq I were purchased from New England Biolabs (Ipswich, MA). Bsh 1236I was purchased from Fermentas (Hanover, MD). Restriction digests contained an 8 μl aliquot from the PCR reaction, 1 U restriction enzyme and 1x restriction enzyme buffer in a 20 μl reaction volume and were incubated overnight at 37°C. Products were separated on 2% agarose-TAE gels, stained with ethidium bromide and visualized by UV illumination. PCR-RFLP tests were performed a minimum of two times for each population.

PCR products were excised from agarose gels, purified with the QIAquick Gel Extraction Kit (Qiagen, Valencia, CA) and cloned into the vector pCR2.1 using the Topo-TA Cloning kit (Invitrogen, Carlsbad, CA). Plasmid DNA was prepared with the Qiagen miniprep kit and digested with Eco RI to verify the presence of insert. Sequencing was performed at the University of Maryland Center for Biosystems Research. Nine clones representing three juveniles from two cysts from the initial tare soil sample were sequenced. Direct sequencing was used to analyze PCR products from: Field 1 (four J2); Field 2 (two J2); as well as G. pallida York, England (one J2, one cyst); G. pallida Chile, G. rostochiensis NY, G. rostochiensis York, England, G. tabacum solanacearum Richmond, VA, G. tabacum tabacum CT (three J2 each); and H. avenae ID (one J2). Sequences were submitted to the GenBank database under accession numbers EF153834 − EF153843.

DNA sequences were assembled using Sequencher 4.7 (Genecodes, Ann Arbor, MI). Alignments were made with Clustal W (Thompson et al., 1994) and checked by eye for consistency of conserved positions among sequences. Alignments were edited in GeneDoc (Nicholas et al., 1997). Globodera artemesiae, G. achilleae, G. millefolii and G. hypolysi were specified as outgroups. ModelTest (Posada and Crandall, 2001) was used to generate parameters useful for constructing a distance tree. Using hLRT criteria, the HKY85 model (Hasegawa et al., 1985) was selected, involving a time-reversible process, non-uniform nucleotide distribution and different transition and transversion rates (HKY+G). Parameters for this model included gamma shape 0.2972, Ti/Tv 2.2957, proportion invariant 0 and −lnL 2514.7830. AIC criteria gave a transversion model with gamma distributed rates across sites and a fraction of sites assumed to be invariable (TVM + I + G), gamma shape 0.7477, proportion invariant 0.3968, −lnL 2509.4099, AIC 5036.8198. Both were implemented in Neighbor Joining (NJ) trees. For Maximum Parsimony trees involving 85 parsimony-informative characters of 846 total characters, fast, stepwise-addition and slow heuristic searches with 1,000 bootstrap replications employed ACCTRAN (accelerated transformation) char acter-state optimization and TBR (tree bisection-reconnection) branch-swapping. This was performed during the slow search on 57,200 trees, with 85,373 rearrangements. All procedures were implemented in PAUP*, v. 4.0b4a (Swofford, 1998).

Species-specific PCR was performed as described by Bulman and Marshall (1997). Briefly, 2 μl nematode extract and 250 μM of each primer (ITS5, PITSr3 and PITSp4) were included in a 25 μl multiplex reaction, with 160μM dNTP, 0.6 U HotMaster Taq and reaction buffer, as described above. Reactions were cycled once at 94°C for 2 min, followed by 35 cycles of 94°C (30 sec), 60°C (30 sec) and 72°C (30 sec), and completed by one cycle at 72°C for 5 min. Negative controls with no DNA were included in each experiment. Primers PITSr3 and PITSp4 were adapted to amplify Globodera tabacum and were named PITSt3 and PITSt4, respectively (Table 2). The modified multiplex reactions included ITS5 and were performed as described above, but had PITSt3 substituted for PITSr3 or PITSt4 substituted for PITSp4. Reaction products from the multiplex assays were analyzed by gel electrophoresis as described above. The experiment was repeated at least twice for each isolate.

Results and Discussion

Second-stage juveniles (n = 80): Measurements are listed in Table 3 and are in micrometers. The morphological characters of second-stage juveniles of this Idaho population agree with those of G. pallida, but exhibited a slightly shorter body mean length (452 ± 36 μm) than the lengths reported by Stone (1972) for the Epworth, Lincolnshire, England, population (486 ± 23 μm) and for the Duddingston, Scotland, population (482 ± 18 μm). Due to the observed variations in tail shape in the Idaho population, additional specimens from Idaho were then compared with cultured specimens of G. pallida originating from Epworth that had been deposited in the USDA Nematode Collection at Beltsville in 1972 (Fig. 1, A-W). The same variations in tail shape, not reported in the original description of G. pallida from England, were observed in both the Idaho population and the type material from England (Fig. 1J,R-W). Representative Idaho specimens with blunt or pointed tail shapes were also subjected to molecular analysis as described below.

Table 3.

Morphometrics of diagnostic characters of second-stage juveniles and cysts of Globodera pallida from Idaho.

Fig. 1.

Photomicrographs of second-stage juveniles of Globodera pallida from Idaho and England. Specimens from Idaho: A-C) heads, G-I) tails. Paratypes from England deposited in the USDA Nematode Collection at Beltsville, MD (slides T-1441p, T-1449p): D) head, J) tail. Specimens from Idaho: E) esophageal region, F) lateral field showing four incisures. K-P) Variations in tail shape of Idaho population (bluntly rounded to finely rounded pointed tip) with (arrows) in L, P and M showing refractive bodies and anal area, respectively. R-W) Variations in tail shape (bluntly rounded to finely rounded pointed tip) of paratypes from England deposited in the USDA Nematode Collection at Beltsville, MD (slide T-1441p for R-V, and slide T-1449p for W), with (arrows) in T and V showing refractive bodies. Q) Whole juvenile, Idaho population.

Cysts (n = 80): Measurements are listed in Table 3 and are in micrometers, excluding ratios. The quality of cysts from which measurements were taken was good, and the vulval regions were intact (Fig. 2). The morphological characters of cysts of this population agree with those of G. pallida except for slightly higher mean in the distance from anus to the nearest edge of fenestra 53.5 (30–80 μm). Both cyst and J2 morphometrics and J2 tail morphology of the Idaho specimens fit well within ranges observed for G. pallida, indicating that the Idaho specimens must represent G. pallida (Stone, 1973) Behrens, 1975. Table 4 shows updated diagnostic morphometrics of four Globodera species genetically related to G. pallida, including typographic corrections to a previous table (EPPO, 2004), and new measurements from G. achilliae type material deposited in the USDA Nematode Collection at Beltsville, MD (slide numbers: T-1478p to T-1492p). Most notably for G. pallida and G. achilleae cysts, the maximum values for the vulva to anus distance and the number of cuticular ridges are expanded.

Fig. 2.

Photomicrographs of the anterior and terminal areas of Globodera pallida cysts from Idaho. A) Anterior region. B-K) Anal-vulval regions with D, E, G-I, J and K (arrows) showing perineal tubercles, vulval-slit, anal areas, punctations and cuticular ridges, respectively. L) Cysts isolated from soil.

Table 4.

Morphometrics for Globodera Behrens, 1975 amended from EPPO (2004)

Molecular identification: To complement the morphological diagnosis of the Idaho population as G. pallida, three molecular assays were performed: PCR-RFLP of rDNA (Blok et al. 1998; Subbotin et al., 2000; Radivojevic et al., 2001; Reid and Pickup, 2005); phylogenetic analysis of DNA sequences; and sequence-specific multiplex PCR (Bulman and Marshall, 1997; Pylypenko et al., 2005). The ease and accuracy of these methods for identifying G. pallida have been previously verified by a number of laboratories using numerous worldwide populations of G. pallida and G. rostochiensis. However, few studies (Thiéry and Mugniéry, 1996; Subbotin et al., 2000) included direct comparison of G. pallida, G. rostochiensis and G. tabacum, prompting us to further validate and extend existing rDNA-based diagnostics to include all three species.

PCR-RFLP: Using primers TW81 and AB28, the isolates from Idaho yielded PCR products of approximately 927 bp, including 18S rDNA (partial 3′) ITS1, 5.8S, ITS2 and 28S (partial 5′). These products appeared similar in size to those from G. rostochiensis and G. pallida (not shown). Based on diagnostic RFLP profiles described previously (Thiéry and Mugniéry, 1996; Blok et al., 1998; Subbotin et al., 2000; Reid and Pickup, 2005), restriction digestion of ITS PCR products with Alu I, Rsa I, Bsh 1236I or Taq I was used to characterize the Idaho isolates. In addition to known populations of G. pallida, G. rostochiensis and Heterodera avenae, isolates of G. tabacum solanacearum and G. tabacum tabacum were included for comparison. Alu I digestion (Fig. 3A) showed that the specimens from Idaho (lanes 15–20) were consistent with G. pallida (lanes 11–14; fragment sizes 505 and 383 bp) and distinct from G. rostochiensis (lane 4; fragment sizes 381, 360 and 148 bp) and H. avenae (lanes 2, 3; uncut). However, Alu I digestion could not discriminate G. pallida from G. tabacum. The Rsa I digestion (Fig. 3B) clearly identified the Idaho population (lanes 15–20) as G. pallida (lanes 11–14; fragment sizes 587 and 385 bp), distinguishing it from G. tabacum (lanes 5–10) and G. rostochiensis (lane 4), each with fragments of 587, 222 and 162 bp. Restriction patterns from the Bsh 1236I digests clearly differentiated the G. pallida isolates (Fig. 3C, lanes 2–6; fragment sizes 503, 347 and 126 bp) from G. rostochiensis (lanes 7–10; fragment sizes 842 and 126 bp), G. tabacum solanacearum (lanes 11–14; fragment sizes 430, 332, 124 and < 100 bp) and G. tabacum tabacum (lanes 15–18; fragment sizes 430, 328 and 114 bp). Taq I digestion also positively identified the Idaho population as G. pallida, but did not discriminate between G. rostochiensis and G. tabacum (not shown). Occasional faint partial digest products and poorly visible fragments less than 100 bp were not included in the analysis and did not preclude detection of diagnostic banding patterns for each species.

Fig. 3.

Amplified PCR products from Globodera spp. digested by two enzymes. Panel A, Alu I and Panel B, Rsa I: Lanes 1, 21) 100 bp ladder (New England Biolabs, Ipswich, MA); lanes 2, 3) H. avenae Idaho; lane 4) G. rostochiensis NY (bulk genomic DNA, 20 ng); lanes 5, 6) G. tabacum solanacearum VA; lanes 7, 8) G. tabacum tabacum CT; lanes 9, 10) G. tabacum solanacearum VA; lanes 11–14) G. pallida York; lanes 15–20) G. pallida Idaho, tare soil; lane 16) blunt-tailed J2; lane 17) pointed-tailed J2. All PCR reactions included DNA extracts from individual juveniles except for lane 4 (20 ng bulk genomic DNA) and lanes 13, 14 (single cyst extracts). Panel C, Bsh 1236I: Lanes 1, 19) 100 bp ladder; lanes 2, 3) G. pallida Idaho field samples; lanes 4, 5) G. pallida York; lane 6) G. pallida Chile; lanes 7–10) G. rostochiensis NY; lanes 11–14) G. tabacum solanacearum VA; lanes 15–18) G. tabacumtabacum CT. Numbers to the left indicate sizes of selected marker bands in the ladder.

The faint products present at approximately 650 bp in H. avenae reactions (Fig. 3A, lanes 2, 3; 3B, lanes 2, 3) are non-specific PCR products, as they also appeared in the undigested ITS PCR reactions of several H. avenae specimens tested (not shown). Among the G. pallida specimens tested from Idaho were some that exhibited unusually blunt or pointed tail shapes (as shown in Fig. 1). These specimens (Fig. 3A,B, lane 16, blunt tail; lane 17, pointed tail) appeared the same in the digests as others having more typical juvenile morphology, thus providing molecular evidence that the tail shape variants were G. pallida. While all populations in this study gave rise to the expected banding patterns, it would not be unusual to find populations with polymorphisms in the ITS region that result in missing restriction sites. Therefore, the use of more than one enzyme is generally recommended to achieve a reliable diagnosis of G. pallida.

DNA sequence analysis: To further confirm the identity of the Idaho isolates as G. pallida, DNA sequences were obtained for the ITS rDNA PCR products obtained with primers TW81 and AB28. A total of 21 point mutations in the amplified region was observed among nine clones representing three J2 from Idaho tare soil. The consensus sequence derived from these clones (EF153837) was identical to the directly sequenced products of specimens from Field 1 (EF153836) and Field 2 (EF153835). Since each base change only appeared in a single clone and none were reflected as heterozygotes in the directly sequenced products, these differences were most likely due to random sequencing errors. No other patterns of variation were observed in the Idaho population that would suggest heterogeneity among different rDNA copies. One overall consensus sequence (designated Gp ID) was used for further phylogenetic comparison to the entire ITS region obtained for other populations of G. pallida, G. rostochiensis and G. tabacum, plus several others obtained from GenBank or published works (Table 1). Relationships among the species and populations inferred from the ITS rDNA are shown in Figure 4. The Gp ID sequence from Idaho was identical to those from Gp Y (York) and Gp D375 (Netherlands) and highly similar to the ITS region from other European populations. With the exception of Gp A from Argentina (DQ097514), all G. pallida populations appeared in a single highly supported clade. Using ModelTest parameters estimated on NJ trees, the Gp A population from Argentina grouped with G. rostochiensis populations with 71 to 91% bootstrap support; fast MP bootstrapped searches with TBR gave a similar grouping with 57% support (trees not shown). However, slow heuristic searches (318 tree consensus) resulted in a different topology, grouping Gp A with other G. pallida populations with 57% bootstrap support, as shown in Figure 4. In considering the equivocal tree position of this population based on ITS, it should be noted that, because many populations of Globodera spp. have demonstrated hybridization potential (Miller, 1983; Baldwin and Mundo-Ocampo, 1991), existence of genetically intermediate populations would not be surprising. The Gp A population could therefore be a hybrid or a naturally occurring variant that diverged from the European type G. pallida populations.

Fig. 4.

Maximum Parsimony phylogram of Globodera species and populations made from a ClustalW alignment of 29 taxa with 846 nucleotide characters from the ITS1 and ITS2 regions of rDNA. Trees were generated using a heuristic search bootstrapped 1,000 times, employing TBR branch-swapping on 57,200 trees, with 85,373 rearrangements and ACCTRAN character-state optimization as implemented in PAUP*. Bootstrap support values occur above branches. TL = 264, CI = 0.814, RI = 0.889, RC = 0.724, HI = 0.186, goodness of fit = −74.707. GenBank accession numbers and abbreviations for nematode populations are given in Table 1. Differential Interference Contrast images were taken of live specimens with distance bars equivalent to 10 μm.

Based upon ITS rDNA, all G. tabacum populations also grouped together in a highly supported clade, with the G. tabacum tabacum sequence from Connecticut (Gtt CT) identical to another from the U.S. of unknown specific geographic origin (Gt Tab1) and one from Japan (Gt J). Gtt L (DQ097515) appeared closer to a clade containing both isolates of G. tabacum solanacearum (Gt Sol1 and Gts VA) and the population of G. tabacum virginiae (Gt Vir1). Likewise, the Gr NY and Gr Y populations appeared together in a clade with populations from Japan (Gr J) and Russia (Gr Ja1). As demonstrated by the head and tail photographs of live cyst nematodes shown in Figure 4, significant variation in qualitative morphological features between individuals makes interspecific comparisons difficult. For instance, while stylet morphology among the cyst nematodes shows considerable overlap (Manduric et al., 2004), the nearly symmetrical mid-tail constriction is a notable characteristic of G. pallida individuals within this population.

Additional GenBank entries, including AF016865 to AF016881, allowed further comparison of the Idaho G. pallida population with the ITS1 region from several other Globodera species and populations (data not shown). Compared to this group of sequences, the Idaho G. pallida ITS1 sequence showed closest similarity to isolates from Romania (differing at 2 bp within 549 bp aligned), Spain and Peru (each differing at 4 bp) and Northern Ireland (5 bp). Thus, all of the phylogenetic analyses of Gp ID clearly placed the Idaho population within G. pallida.

Sequence-specific PCR: As further confirmation of the species identity, a multiplex PCR assay using species-specific primers was performed according to the method of Bulman and Marshall (1997). Globodera pallida positive control specimens from York (Fig. 5, lanes 8, 9), all J2 obtained from tare (lanes 2, 3) or field samples (lanes 4–7) and the isolate from Chile (lanes 14–16) were positive for the expected G. pallida-specific 265 bp band. As was done for the restriction digests, nematodes from the Idaho population that exhibited unusually blunt or pointed tail shapes were also subjected to this multiplex assay. These specimens also tested positive for G. pallida, adding further confirmation for their identity (not shown). Globodera rostochiensis samples from New York (lanes 17, 18) and York, England (lanes 20–22), gave the 434 bp band expected for this species. One reaction (lane 19) gave no product; further testing showed this was due to a poor quality extract. Because primers ITS5, PITSr3 and PITSp4 had not previously been tested with G. tabacum, we examined isolates of G. tabacum solanacearum (lanes 10, 11) and G. tabacum tabacum (lanes 12, 13). All G. tabacum isolates, including G. tabacum solanacearum from Nottoway, VA (not shown), and water control (lane 25) reactions were negative for PCR product, confirming the expected results based upon sequence information. The H. avenae reactions (lanes 23, 24) had faint products that were most likely due to mismatch priming with these templates, since they were larger than the expected size and did not consistently appear with repeated testing (not shown).

Fig. 5.

Multiplex polymerase chain reactions of ITS rDNA from Globodera spp. using primers PITSr3, PITSp4 and ITS5. Single juvenile DNA extracts used for these reactions are the same as for Figure 7. Lanes 1) and 26)100 bp ladder (New England Biolabs, Ipswich, MA); lanes 2, 3) Idaho tare soil; lanes 4, 5) Idaho Field 1; lanes 6, 7) Idaho Field 2; lanes 8, 9) G. pallida York; lanes 10, 11) G. tabacum solanacearum, VA; lanes 12,13) G. tabacum tabacum CT; lanes 14–16) G. pallida, Chile; lanes 17–19) G. rostochiensis NY; lanes 20–22) G. rostochiensis York; lanes 23, 24) H. avenae Idaho; lane 25) no DNA control.

We then used DNA sequence alignments to design new primers that produced a positive PCR reaction for G. tabacum. While ITS sequence variation among the species was limited, we were able to modify the sequences of primers PITSr3 and PITSp4. The new primers, named PITSt3 and PITSt4 (Table 2), were each paired separately with ITS5. As predicted from the sequence information, PITSt4 and ITS5 amplified a 265 bp band from both G. tabacum and G. rostochiensis but not from G. pallida or the controls (Fig. 6A). Included among the Idaho nematodes testing negative for G. tabacum with this primer set were blunt-tailed (Fig. 6A, lane 12) and pointed-tailed (lane 13) specimens.

Fig. 6.

Conventional PCR reaction of Globodera populations using species-specific primers modified from PITSr3 and PITSp4. A. Reactions with primers PITSt4 and ITS5. B. Reactions with primers PITSt3 and ITS5. Lanes 1, 19) 100 bp ladder; lane 2) G. rostochiensis NY; lane 3, 4) G. tabacum solanacearum Richmond, VA; lane 5, 6) G. tabacum solanacearum Nottoway County; lane 7, 8) G. tabacum tabacum; lane 9, 10) G. pallida York; lanes 11–16) G. pallida Idaho, tare soil; lane 17) H. avenae Idaho; lane 18) no DNA control.

Primers PITSt3 and ITS5 amplified a 434 bp band from all three G. tabacum isolates but not from G. rostochiensis or any of the G. pallida specimens from Idaho, including tail shape variants (Fig. 6B, lanes 12, 13). This primer pair occasionally produced a faint PCR band at 434 bp from the G. pallida York isolate (Fig. 6B, lane 9). Assuming this was due to contamination of the York specimen, we repeated the assay several times using fresh reagents and template preparations (not shown). This phenomenon appeared only sporadically with the G. pallida isolate from York but not with specimens from any other location. It is not clear why the York population was uniquely affected, since the sequence of its ITS region was identical to G. pallida from Idaho. Because only two nucleotide changes separate the PITSt3 primer from an exact match to the G. pallida sequence, mismatch primer binding on those templates or priming from a rare variant ITS repeat are possible. Raising the annealing temperature slightly from 60°C or otherwise changing the cycling conditions might eliminate this effect, but we did not investigate this any further. Because relatively few nucleotide differences separate all three species and intraspecific variation is known to occur in Globodera ITS regions, phenomena such as this highlight the need for caution when applying and interpreting species-specific PCR assays.

Because the PITSt4 primer performed more reliably overall than PITSt3, we then tested this primer in mul tiplex PCR with ITS5 and PITSp4 (Fig. 7). As expected, PCR of G. rostochiensis showed bands at both 434 bp and 265 bp (Fig. 7, lanes 17–22). Globodera tabacum isolates showed only the 265 bp band (lanes 10–13); G. pallida (lanes 2–9), H. avenae (lanes 23, 24) and control reactions (lane 25) were negative. According to these results, the appearance of both bands is diagnostic for G. rostochiensis, one band at 265 bp indicates G. tabacum and the absence of product is consistent with G. pallida, although not definitive. Thus, this multiplex PCR modification of the Bulman and Marshall (1997) assay provides a means to positively identify G. tabacum in a similarly simple and straightforward manner. When combined with the original assay, it is now possible to discriminate between G. pallida, G. tabacum and G. rostochiensis. This additional test could be invaluable in diagnostic situations that may arise in the Americas, where the potential for these species to spread across borders has already caused heightened regulatory concern and had significant trade implications.

Fig. 7.

Multiplex PCR reaction of Globodera populations using primers PITSt4, PITSr3 and ITS5. DNA extracts used for these reactions were the same as in Figure 5. See Table 1 for population codes.