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Fingerprinting sequence variation in ribosomal DNA of parasites by DGGE
Molecular and Cellular Probes (1996) 10, 99–105
Fingerprinting sequence variation in ribosomal DNA of parasites by DGGE Robin Gasser,1∗,2 Peter Nansen2 and Per Guldberg3 1
The University of Melbourne, Department of Veterinary Science, Werribee, Australia, Danish Centre for Experimental Parasitology, DK-1870 Frederiksberg C, Denmark, and 3 Danish Cancer Society, Division for Cancer Biology, DK-2100 Copenhagen, Denmark
2
(Received 14 August 1995, Accepted 27 October 1995) Although there is a tendency for rDNA genes within a species to maintain sequence homogeneity, there can be significant levels of variation among rDNA repeat sequences within populations or individuals of a species as a consequence of mutation mechanisms. To date, there have been no practical techniques available in molecular parasitology that allow the extent of sequence variation among the repeats (ie, number of sequence types) to be displayed visually. In this report, we describe the use of the denaturing gradient gel electrophoresis (DGGE) technique for the rapid screening of parasite rDNA for sequence variation without the need for exhaustive cloning or DNA sequencing. The resolution of this variation by DGGE provides a diagnostic fingerprint for a species. 1996 Academic Press Limited KEYWORDS: DGGE, ribosomal DNA, internal transcribed spacer, sequence variation, parasite, species identification.
INTRODUCTION Nuclear ribosomal (r) DNA of eukaryotic organisms is a large multigene family consisting of tandem repeat sequences, usually found in clusters in specific chromosomes.1 Homogeneity within a species coupled with differences between species for sequences of tandemly repeated genes is termed concerted evolution, which appears to arise from either a sudden or gradual correction mechanism (=molecular drive).2,3 Molecular drive acts upon the constant turnover in rDNA sequence and spreads variants through the multiple copies. Although there is a tendency for the individual rDNA genes within a species to maintain sequence homogeneity, there can be significant levels of variation among rDNA repeat sequences within a species,4,5 which appear to be a result of slippage mutations during DNA replication.6,7
Recent research has demonstrated that internal transcribed spacer rDNA (ITS1 and ITS2) provides reliable genetic markers for a range of parasites for diagnostic purposes and establishing systematic relationships.8–11 Our work has focused particularly on the ITS2 of helminth parasites and has shown that this spacer provides reliable species markers.12–16 Recently, we have observed that some degree of polymorphism may exist in the ITS2 sequence (=different sequence types) of some species of parasitic nematodes.13,15,16 Although DNA sequencing, PCR-linked RFLP and cloning techniques can be used to detect such sequence variation, 14–16 these techniques do not allow the number of different sequence types to be determined accurately. Advances in DNA screening procedures have included the development and improvement of de-
∗ Author to whom correspondence should be addressed at: The University of Melbourne, Werribee, Australia.
0890–8508/96/020099+07 $18.00/0
99
1996 Academic Press Limited
100
R. Gasser et al. Table 1.
DNA isolates used in this study
Species
Isolate∗
Host origin
Geographical origin
Trichostrongylus axei
Ta2 Ta3
Sheep Cattle
Victoria, Australia New South Wales, Australia
T. colubriformis
Tc1 Tc2 Tc3 Tc5 Tc9
Sheep Sheep Sheep/drug susceptible Sheep/drug resistant Goat
Victoria, Australia Port Lincoln, South Australia Nouzilly, France Nouzilly, France Poitou Charentes, France
T. vitrinus
Tv1 Tv4 Tv6
Sheep Sheep Sheep
Victoria, Australia Edinburgh, Scotland Nouzilly, France
T. T. T. T.
Tp1 Tru1 Tre1 Tt1
Sheep Sheep Rabbit Grouse
Adelaide, South Australia Strathalbyn, South Australia Victoria, Australia Inverness, Scotland
probolurus rugatus retortaeformis tenuis
∗All isolates except Ta3 were male worms. Ta3 was an isolate of third stage larvae.
naturing gradient gel electrophoresis (DGGE), temperature gradient gel electrophoresis (TGGE) and single stranded conformational polymorphism (SSCP) techniques.17 Particularly DGGE has found a broad applicability in a range of fields of biology to screen for allelic variation, because of its versatility, high resolution power and reproducibility.18,19 The mutational resolving power of DGGE relies on the physical separation of similar DNA fragments differing in melting properties as a consequence of differences in nucleotide composition.20 In this report, we describe the use of DGGE for the direct display of different ITS2 rDNA types in closely-related species of parasitic helminth. The technique exploits the existence of sequence variation in ITS2 repeats among species (‘interspecific heterogeneity’) coupled with absent or low levels of variation within a species (‘intraspecific homogeneity’) to generate a diagnostic fingerprint for a parasite species.
MATERIALS AND METHODS Parasites For the current study, we used parasites of animal health importance as a model system (closely-related species of Trichostrongylus (Table 1)). Adult worms of each species were isolated from the gastrointestinal tract of mammalian or bird hosts, and individual adult male worms were identified morphologically to the species level21 and frozen at −70°C until use. Each worm isolate was from a single host, and a larval isolate (Ta3) was obtained from a monospecific laboratory culture of T. axei.
Isolation and purification of genomic DNA Genomic DNA was isolated from 5–10 worms of each isolate by a standard small-scale method of sodium dodecyl-sulfate and proteinase K treatment, phenol/chloroform extraction, ethanol precipitation and purification with PrepaGene (Bio-Rad).22
PCR ITS2 (including flanking sequence) was amplified by PCR23 from worm DNA (1–5 ng template) using oligonucleotide primers NC1>: 5′-ACG TCT GGT TCA GGG TTG TT-3′ (forward) and
DGGE of parasite rDNA
101
Fig. 1. DGGE analysis of ITS2 rDNA of three species of Trichostrongylus. Top (‘rDNA’): Amplification of ITS2 from the rDNA transcriptional unit using conserved primers NC1> and
were examined on 1·5% agarose-TBE (89 m TrisHCl, 89 m boric acid, 1 m EDTA, pH 7·4) gels after ethidium bromide staining.25 Approximate molecular weight of fragments was determined by comparison to UX174-Hae III or pGEMTM markers (Promega), and gels were photographed with transillumination using Polaroid 667 film (Kodak).
the ITS2 sequences of T. axei, T. colubriformis and T. vitrinus15 using the computer algorithm MELT87.28 Based on these parameters, ITS2 PCR products (15 ll) were directly subjected to electrophoresis in a 6% polyacrylamide gel containing a 15–50% gradient of denaturant, and run at 150V and 54°C for 6 h. Gels were then incubated in ethidium bromide (0.5 lg ml−1) for 5 min and photographed.
DGGE PCR-linked RFLP DGGE was performed essentially as described previously.26,27 One relevant melting domain with an estimated temperature of 66·1°C was determined for
Bands excised from denaturing gradient gels were suspended in 100 ll H2O overnight (4°C), the gel
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slice removed and 1 ll of the suspension diluted to 1/10000. Then, 1 ll of the diluted suspension was subjected to PCR using NC1> and
RESULTS In a first step, the ITS2 PCR products of different isolates of T. axei, T. colubriformis and T. vitrinus were examined by DGGE (Fig. 1). On agarose gels, there was no detectable size difference among the products of the 3 species (Fig. 1, middle). Examination of these products by DGGE revealed different levels of ITS2 sequence variation among the species, which generated a ‘fingerprint’ for each species/isolate (Fig. 1, bottom). PCR products were examined several times on different days under the same DGGE conditions, and fingerprints of sequence variation were found to be highly reproducible. For each species examined, the most dominant band in DGGE was considered to be the ‘parent’ ITS2 molecule, while fainter bands were considered to be ‘mutant’ molecules (Fig. 1, bottom). T. vitrinus exhibited a larger number (n=12–15) of bands corresponding to variant molecules than did T. axei or T. colubriformis (up to 6 bands). These findings are in agreement with previous results achieved by DNA sequencing, which demonstrated that a higher level of polymorphism existed in the ITS2 sequence of T. vitrinus compared with T. axei and T. colubriformis.15,29 The examination of multiple isolates of each species (Table 1) in DGGE demonstrated that there was some difference in the number of bands (sequence variants) among isolates of T. vitrinus, which was not the case for the other 2 species examined (Fig. 1, bottom). Although it is possible that some of these differences may be a consequence of biased PCR amplification of certain variant sequences in some isolates, they may reflect actual population variation in the ITS2 repeats among isolates of T. vitrinus. Due to the small number of isolates examined in this study, it would be premature to conclude that there is no difference in the number of ITS2 sequence variants in T. axei and T. colubriformis. In order to confirm the identity of some bands (considered to be clonal ITS2 variant molecules), bands a–i (Fig. 1, bottom) were excised from denaturing gels and examined by PCR-linked RFLP. The
molecular weight of the products re-amplified from these bands corresponded to that of the ITS2 PCR products amplified from genomic DNA (Fig. 2, left). Based on DNA sequence data and restriction maps of the ITS2 of T. axei, T. colubriformis and T. vitrinus,15 endonuclease digestion of PCR products amplified from bands a–h with Rsa I or Dra I revealed that they corresponded with ITS2 sequences (Fig. 2, right). The PCR product derived from band-i remained undigested with Dra I, while it was digested with Rsa I. The difference in the Rsa I digestion of this product compared with the products from bands a–h reflected the sequence polymorphism previously identified at position 83 in the ITS2 sequence of T. vitrinus.15 In the same study,15 we were not able to accurately record sequence polymorphism at the Dra I recognition (TTTAAA; positions 162–167, 192–197). Re-reading of original ITS2 sequencing gels of 3 individual T. vitrinus worms revealed that there was in fact polymorphism at positions 162 and 192 in the sequence derived from one of the worms. In addition to the T previously recorded, a faint A was detected at each of the positions. Although the A had a low intensity on the sequencing gel compared with the T (approx. 1:5 intensity), these observations would imply the loss of the two Dra I recognition sites (ATTAAA instead of TTTAAA) in the ITS2 repeat population corresponding to band-i. The current findings indicate that a ‘mutational’ sequence type (band-i) lacking the Dra I sites was isolated by DGGE in spite of its low abundance (compare Fig. 1, bottom, lane 6, band-i). It is very unlikely that, over this short sequence, the base changes in these sites are the result of base misincorporations during PCR, because it has been estimated that the misincorporation rate per nucleotide per cycle for Taq polymerase is approximately 2×10−4.30 Having shown that there was no or relatively little variation in DGGE banding patterns within T. axei, T. colubriformis and T. vitrinus and that bands excised from denaturing gels corresponded with ITS2, the next step was to assess the capacity of DGGE to fingerprint ITS2 sequence variation in a broader range of closely-related species (Fig. 3). Although there was no detectable difference among the 7 species in size of the PCR products examined on agarose gels (not all data shown; compare Fig. 1, middle), DGGE was able to reproducibly resolve significant differences in the level of sequence variation among the species. Also, the relative positions of individual parent bands on DGGE gels appeared to be associated with the percentage of nucleotide sequence differences (approx. 1·5–7%) in the ITS2 among some of the species.15
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Fig. 2. Analysis of bands a–i (see fig. 1) by PCR-linked RFLP. Agarose gel electrophoresis of PCR products amplified from the individual bands (left), and the same products digested with Dra I or Rsa I (right).
Fig. 3. DGGE fingerprinting of ITS2 rDNA sequence variation in seven species of Trichostrongylus. Lanes 1, T. axei (Ta2); lane 2, T. colubriformis (Tc3); lane 3, T. vitrinus (Tv1); lane 4, T. probolurus; lane 5, T. rugatus; lane 6, T. retortaeformis and lane 7, T. tenuis.
DISCUSSION Separation of DNA in DGGE is based on the electrophoretic mobility of a partially melted molecule in polyacrylamide gels, which is decreased compared with that of the helical form of the molecule. The melting of fragments proceeds in discrete melting domains, which are regions of sequence with an identical melting domain. Once the melting domain with the lowest melting temperature is at a particular position in the gel, a transition of helical to partially melted molecules occurs, and the migration of the molecules will almost stop. Sequence variation within such domains causes the melting temperatures of the variant (mutant) molecules to differ. As a consequence, sequence variants will stop migrating at different positions in the denaturing gradient and will be separated. DGGE can be used to analyse genomic DNA, but recently the technique has been adapted to PCR products in order to analyse sequences of interest. A modification to the latter approach has been to incorporate a GC-rich sequence into one
of the primers used in PCR, which alters the melting behaviour of the fragment and permits the separation of almost 100% of variant molecules.24 Based on the results of the present study, we have demonstrated that DGGE is a powerful analytical technique to screen for sequence variation in parasite rDNA. The technique displays visually the different sequence types of ITS2 in one step. Given that different levels of mutational variation were observed among the species examined and similar (or the same) levels within a species, the technique allowed a diagnostic fingerprint of a species to be generated using a defined rDNA region. This feature should be of particular use for the species identification of individual developmental stages of parasites where morphological characters are unreliable, and therefore has important implications for diagnosis of infection(s). Given that we have already shown that ITS2 can be amplified from single worms, eggs and larvae of a range of strongylid nematodes,13,15,16,22 this approach has formidable potential for the identification of these stages without the need for restriction endonuclease analysis or DNA sequencing. The DGGE technique also has valuable preparative qualities, in that individual variant sequences (even if in low abundance) can be excised from gels, subsequently reamplified and directly subjected to PCR-RFLP or DNA sequence analysis. In this way, it should be possible to quantitatively determine levels of sequence variation among rDNA repeats within and among parasite species without the need for time-consuming isolation of variant sequences using DNA cloning procedures. Although in this study, DGGE has been used to analyse the ITS2 of parasites of veterinary significance, it has the potential to be applied to a broad range of parasites and other eukaryotic organisms of medical importance.
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ACKNOWLEDGEMENTS This work was supported by the Australian Research Council (no. A19531013), Danish Research Academy (DANVIS no. D930061) and Danish National Research Foundation. Thanks to Henrik Bøgh and Birgitte Sønderby for their support. We are also grateful to H. Hoste, C. Chartier (France), I. Beveridge (Melbourne), M. O’Callaghan (South Australia), P. Hudson (Inverness) and R. Coop (Edinburgh) who assisted in collecting or provided parasite isolates.
REFERENCES 1. Gerbi, S. A. (1986). The evolution of eukaryotic ribosomal DNA. BioSystems 19, 247–58. 2. Arnheim, N. (1983). Concerted evolution of multigene families. In Evolution of Genes and Proteins (M. Nei & R. K. Koehn, eds.) pp. 38–61. Sunderland, Massachusetts: Sinauer. 3. Dover, G. A. (1989). Linkage disequilibrium and molecular drive in the rDNA gene family. Genetics 122, 249–52. 4. Gonzalez, I. L., Sylvester, J. E. & Schmickel, R. D. (1988). Human 28S ribosomal RNA sequence heterogeneity. Nucleic Acids Research 16, 10213–23. 5. Crease, T. J. & Lynch, M. (1991). Ribosomal DNA variation in Daphnia pulex. Molecular Biology and Evolution 8, 620–40. 6. Hancock, J. M., Tautz, D. & Dover, G. A. (1989). Evolution of the secondary structures and compensatory mutations of ribosomal RNAs of Drosophila melanogaster. Molecular Biology and Evolution 5, 393–414. 7. Hancock, J. M. & Dover, G. A. (1990). Compensatory slippage in the evolution of ribosomal RNA genes. Nucleic Acids Research 18, 5949–54. 8. Luton, K., Walker, D. & Blair, D. (1992). Comparisons of ribosomal internal transcribed spacers from two congeneric species of flukes (Platyhelminthes: Trematoda: Digenea). Molecular and Biochemical Parasitology 56, 323–8. 9. Anderson, G. R. & Barker, S. C. (1993). Species differentiation in the Didymozoidae (Digenea): restriction length differences in internal transcribed spacer and 5.8S ribosomal DNA. International Journal for Parasitology 23, 133–6. 10. Bowles, J. & McManus, D. P. (1993). Rapid discrimination of Echinococcus species and strains using a polymerase chain reaction-based RFLP method. Molecular and Biochemical Parasitology 57, 231–40. 11. Wachira, T. M., Bowles, J. & McManus, D. P. (1993). Molecular examination of the sympatry and distribution of sheep and camel strains of Echinococcus granulosus. American Journal of Tropical Medicine and Hygiene 48, 473–9. 12. Gasser, R. B., Chilton, N. B., Hoste, H. & Stevenson, L. A. (1994). Species identification of trichostrongyle nematodes by PCR-linked RFLP. International Journal for Parasitology 24, 291–3. 13. Campbell, A. J. D., Gasser, R. B. & Chilton, N. B. (1995). Differences in a ribosomal sequence of Strongylus species allows identification of single eggs. International Journal for Parasitology 25, 359–65. 14. Gasser, R. B. & Chilton, N. B. (1995). Characterisation
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of taeniid cestode species by PCR-RFLP of ITS2 ribosomal DNA. Acta Tropica 59, 31–40. Hoste, H., Chilton, N. B., Gasser, R. B. & Beveridge, I. (1995). Differences in the second internal transcribed spacer (ribosomal DNA) between five species of Trichostrongylus (Nematoda: Trichostrongylidae). International Journal for Parasitology 25, 75–80. Stevenson, L. A., Chilton, N. B. & Gasser, R. B. (1995). Differentiation of Haemonchus placei from H. contortus (Nematoda: Trichostrongylidae) by the ribosomal second internal transcribed spacer. International Journal for Parasitology 25, 483–8. Lessa, E. P. & Applebaum, G. (1993). Screening techniques for detecting allelic variation in DNA sequences. Molecular Ecology 2, 119–29. Sheffield, V. C., Beck, J. S., Nichols, B., Cousineau, A., Lidral, A. & Stone, E. M. (1992). Detection of multiallele polymorphisms within gene sequences by GC-clamped denaturing gradient gel electrophoresis. American Journal of Human Genetics 50, 567–75. Guldberg, P. & Gu¨ttler, F. (1994). Broad-range DGGE for single step mutation scanning of entire genes: application to human phenylalanine hydroxylase gene. Nucleic Acids Research 22, 880–1. Myers, R. M., Maniatis, T. & Lerman, L. S. (1987). Detection and localization of single base changes by denaturing gradient gel electrophoresis. Methods in Enzymology 155, 501–27. Nagaty, H. F. (1932). The genus Trichostrongylus Looss, 1905. Annals of Tropical Medicine and Hygiene 26, 457–518. Gasser, R. B., Chilton, N. B., Hoste, H. & Beveridge, I. (1993). Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Research 21, 2525–6. Mullis, K. B., Faloona, F., Scharf, S., Saiki, R., Horn, G. & Erlich, H. (1986). Specific enzymatic amplification of DNA in vitro: the polymerase chain reaction. Cold Spring Harbor Symposium of Quantitative Biology 51, 263–73. Sheffield, V. C., Cox, D. R., Lerman, L. S. & Myers, R. M. (1989). Attachment of a 40-base pair G+C-rich sequence (GC-clamp) to genomic DNA fragments by polymerase chain reaction results in improved detection of single-base changes. Proceedings of the National Academy of Science, USA 86, 232–6. Sambrook, J., Fritsch, E. F. & Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, 2nd ed. Salem: Cold Spring Harbour Laboratory Press. Myers, R. M., Fischer, S. G., Lerman, L. S. & Maniatis, T. (1985). Nearly all base substitutions in DNA fragments joined to a GC-clamp can be detected by denaturing gradient gel electrophoresis. Nucleic Acids Research 13, 3131–45. Guldberg, P., Henriksen, K. F. & Gu¨ttler, F. (1993). Molecular analysis of phenylketonuria in Denmark: 99% of the mutations detected by denaturing gradient gel electrophoresis. Genomics 17, 141–6. Lerman, L. S. & Silverstein, K. (1987). Computational simulation of DNA melting and its application to denaturing gradient gel electrophoresis. Methods in Enzymology 155, 482–501. Hoste, H., Gasser, R. B., Chilton, N. B., Mallet, S. & Beveridge, I. (1993). Lack of intraspecific variation in the second internal transcribed spacer (ITS-2) of Trichostrongylus colubriformis ribosomal
DGGE of parasite rDNA DNA. International Journal for Parasitology23, 1069–71. 30. Saiki, R. K., Gelfand, D. H., Stoffel, S., Scharf, S. J., Higuchi, R., Horn, G. T., Mullis, K. B. & Erlich, H.
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Fingerprinting sequence variation in ribosomal DNA of parasites by DGGE Robin Gasser,1∗,2 Peter Nansen2 and Per Guldberg3 1
The University of Melbourne, Department of Veterinary Science, Werribee, Australia, Danish Centre for Experimental Parasitology, DK-1870 Frederiksberg C, Denmark, and 3 Danish Cancer Society, Division for Cancer Biology, DK-2100 Copenhagen, Denmark
2
(Received 14 August 1995, Accepted 27 October 1995) Although there is a tendency for rDNA genes within a species to maintain sequence homogeneity, there can be significant levels of variation among rDNA repeat sequences within populations or individuals of a species as a consequence of mutation mechanisms. To date, there have been no practical techniques available in molecular parasitology that allow the extent of sequence variation among the repeats (ie, number of sequence types) to be displayed visually. In this report, we describe the use of the denaturing gradient gel electrophoresis (DGGE) technique for the rapid screening of parasite rDNA for sequence variation without the need for exhaustive cloning or DNA sequencing. The resolution of this variation by DGGE provides a diagnostic fingerprint for a species. 1996 Academic Press Limited KEYWORDS: DGGE, ribosomal DNA, internal transcribed spacer, sequence variation, parasite, species identification.
INTRODUCTION Nuclear ribosomal (r) DNA of eukaryotic organisms is a large multigene family consisting of tandem repeat sequences, usually found in clusters in specific chromosomes.1 Homogeneity within a species coupled with differences between species for sequences of tandemly repeated genes is termed concerted evolution, which appears to arise from either a sudden or gradual correction mechanism (=molecular drive).2,3 Molecular drive acts upon the constant turnover in rDNA sequence and spreads variants through the multiple copies. Although there is a tendency for the individual rDNA genes within a species to maintain sequence homogeneity, there can be significant levels of variation among rDNA repeat sequences within a species,4,5 which appear to be a result of slippage mutations during DNA replication.6,7
Recent research has demonstrated that internal transcribed spacer rDNA (ITS1 and ITS2) provides reliable genetic markers for a range of parasites for diagnostic purposes and establishing systematic relationships.8–11 Our work has focused particularly on the ITS2 of helminth parasites and has shown that this spacer provides reliable species markers.12–16 Recently, we have observed that some degree of polymorphism may exist in the ITS2 sequence (=different sequence types) of some species of parasitic nematodes.13,15,16 Although DNA sequencing, PCR-linked RFLP and cloning techniques can be used to detect such sequence variation, 14–16 these techniques do not allow the number of different sequence types to be determined accurately. Advances in DNA screening procedures have included the development and improvement of de-
∗ Author to whom correspondence should be addressed at: The University of Melbourne, Werribee, Australia.
0890–8508/96/020099+07 $18.00/0
99
1996 Academic Press Limited
100
R. Gasser et al. Table 1.
DNA isolates used in this study
Species
Isolate∗
Host origin
Geographical origin
Trichostrongylus axei
Ta2 Ta3
Sheep Cattle
Victoria, Australia New South Wales, Australia
T. colubriformis
Tc1 Tc2 Tc3 Tc5 Tc9
Sheep Sheep Sheep/drug susceptible Sheep/drug resistant Goat
Victoria, Australia Port Lincoln, South Australia Nouzilly, France Nouzilly, France Poitou Charentes, France
T. vitrinus
Tv1 Tv4 Tv6
Sheep Sheep Sheep
Victoria, Australia Edinburgh, Scotland Nouzilly, France
T. T. T. T.
Tp1 Tru1 Tre1 Tt1
Sheep Sheep Rabbit Grouse
Adelaide, South Australia Strathalbyn, South Australia Victoria, Australia Inverness, Scotland
probolurus rugatus retortaeformis tenuis
∗All isolates except Ta3 were male worms. Ta3 was an isolate of third stage larvae.
naturing gradient gel electrophoresis (DGGE), temperature gradient gel electrophoresis (TGGE) and single stranded conformational polymorphism (SSCP) techniques.17 Particularly DGGE has found a broad applicability in a range of fields of biology to screen for allelic variation, because of its versatility, high resolution power and reproducibility.18,19 The mutational resolving power of DGGE relies on the physical separation of similar DNA fragments differing in melting properties as a consequence of differences in nucleotide composition.20 In this report, we describe the use of DGGE for the direct display of different ITS2 rDNA types in closely-related species of parasitic helminth. The technique exploits the existence of sequence variation in ITS2 repeats among species (‘interspecific heterogeneity’) coupled with absent or low levels of variation within a species (‘intraspecific homogeneity’) to generate a diagnostic fingerprint for a parasite species.
MATERIALS AND METHODS Parasites For the current study, we used parasites of animal health importance as a model system (closely-related species of Trichostrongylus (Table 1)). Adult worms of each species were isolated from the gastrointestinal tract of mammalian or bird hosts, and individual adult male worms were identified morphologically to the species level21 and frozen at −70°C until use. Each worm isolate was from a single host, and a larval isolate (Ta3) was obtained from a monospecific laboratory culture of T. axei.
Isolation and purification of genomic DNA Genomic DNA was isolated from 5–10 worms of each isolate by a standard small-scale method of sodium dodecyl-sulfate and proteinase K treatment, phenol/chloroform extraction, ethanol precipitation and purification with PrepaGene (Bio-Rad).22
PCR ITS2 (including flanking sequence) was amplified by PCR23 from worm DNA (1–5 ng template) using oligonucleotide primers NC1>: 5′-ACG TCT GGT TCA GGG TTG TT-3′ (forward) and
DGGE of parasite rDNA
101
Fig. 1. DGGE analysis of ITS2 rDNA of three species of Trichostrongylus. Top (‘rDNA’): Amplification of ITS2 from the rDNA transcriptional unit using conserved primers NC1> and
were examined on 1·5% agarose-TBE (89 m TrisHCl, 89 m boric acid, 1 m EDTA, pH 7·4) gels after ethidium bromide staining.25 Approximate molecular weight of fragments was determined by comparison to UX174-Hae III or pGEMTM markers (Promega), and gels were photographed with transillumination using Polaroid 667 film (Kodak).
the ITS2 sequences of T. axei, T. colubriformis and T. vitrinus15 using the computer algorithm MELT87.28 Based on these parameters, ITS2 PCR products (15 ll) were directly subjected to electrophoresis in a 6% polyacrylamide gel containing a 15–50% gradient of denaturant, and run at 150V and 54°C for 6 h. Gels were then incubated in ethidium bromide (0.5 lg ml−1) for 5 min and photographed.
DGGE PCR-linked RFLP DGGE was performed essentially as described previously.26,27 One relevant melting domain with an estimated temperature of 66·1°C was determined for
Bands excised from denaturing gradient gels were suspended in 100 ll H2O overnight (4°C), the gel
102
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slice removed and 1 ll of the suspension diluted to 1/10000. Then, 1 ll of the diluted suspension was subjected to PCR using NC1> and
RESULTS In a first step, the ITS2 PCR products of different isolates of T. axei, T. colubriformis and T. vitrinus were examined by DGGE (Fig. 1). On agarose gels, there was no detectable size difference among the products of the 3 species (Fig. 1, middle). Examination of these products by DGGE revealed different levels of ITS2 sequence variation among the species, which generated a ‘fingerprint’ for each species/isolate (Fig. 1, bottom). PCR products were examined several times on different days under the same DGGE conditions, and fingerprints of sequence variation were found to be highly reproducible. For each species examined, the most dominant band in DGGE was considered to be the ‘parent’ ITS2 molecule, while fainter bands were considered to be ‘mutant’ molecules (Fig. 1, bottom). T. vitrinus exhibited a larger number (n=12–15) of bands corresponding to variant molecules than did T. axei or T. colubriformis (up to 6 bands). These findings are in agreement with previous results achieved by DNA sequencing, which demonstrated that a higher level of polymorphism existed in the ITS2 sequence of T. vitrinus compared with T. axei and T. colubriformis.15,29 The examination of multiple isolates of each species (Table 1) in DGGE demonstrated that there was some difference in the number of bands (sequence variants) among isolates of T. vitrinus, which was not the case for the other 2 species examined (Fig. 1, bottom). Although it is possible that some of these differences may be a consequence of biased PCR amplification of certain variant sequences in some isolates, they may reflect actual population variation in the ITS2 repeats among isolates of T. vitrinus. Due to the small number of isolates examined in this study, it would be premature to conclude that there is no difference in the number of ITS2 sequence variants in T. axei and T. colubriformis. In order to confirm the identity of some bands (considered to be clonal ITS2 variant molecules), bands a–i (Fig. 1, bottom) were excised from denaturing gels and examined by PCR-linked RFLP. The
molecular weight of the products re-amplified from these bands corresponded to that of the ITS2 PCR products amplified from genomic DNA (Fig. 2, left). Based on DNA sequence data and restriction maps of the ITS2 of T. axei, T. colubriformis and T. vitrinus,15 endonuclease digestion of PCR products amplified from bands a–h with Rsa I or Dra I revealed that they corresponded with ITS2 sequences (Fig. 2, right). The PCR product derived from band-i remained undigested with Dra I, while it was digested with Rsa I. The difference in the Rsa I digestion of this product compared with the products from bands a–h reflected the sequence polymorphism previously identified at position 83 in the ITS2 sequence of T. vitrinus.15 In the same study,15 we were not able to accurately record sequence polymorphism at the Dra I recognition (TTTAAA; positions 162–167, 192–197). Re-reading of original ITS2 sequencing gels of 3 individual T. vitrinus worms revealed that there was in fact polymorphism at positions 162 and 192 in the sequence derived from one of the worms. In addition to the T previously recorded, a faint A was detected at each of the positions. Although the A had a low intensity on the sequencing gel compared with the T (approx. 1:5 intensity), these observations would imply the loss of the two Dra I recognition sites (ATTAAA instead of TTTAAA) in the ITS2 repeat population corresponding to band-i. The current findings indicate that a ‘mutational’ sequence type (band-i) lacking the Dra I sites was isolated by DGGE in spite of its low abundance (compare Fig. 1, bottom, lane 6, band-i). It is very unlikely that, over this short sequence, the base changes in these sites are the result of base misincorporations during PCR, because it has been estimated that the misincorporation rate per nucleotide per cycle for Taq polymerase is approximately 2×10−4.30 Having shown that there was no or relatively little variation in DGGE banding patterns within T. axei, T. colubriformis and T. vitrinus and that bands excised from denaturing gels corresponded with ITS2, the next step was to assess the capacity of DGGE to fingerprint ITS2 sequence variation in a broader range of closely-related species (Fig. 3). Although there was no detectable difference among the 7 species in size of the PCR products examined on agarose gels (not all data shown; compare Fig. 1, middle), DGGE was able to reproducibly resolve significant differences in the level of sequence variation among the species. Also, the relative positions of individual parent bands on DGGE gels appeared to be associated with the percentage of nucleotide sequence differences (approx. 1·5–7%) in the ITS2 among some of the species.15
DGGE of parasite rDNA
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Fig. 2. Analysis of bands a–i (see fig. 1) by PCR-linked RFLP. Agarose gel electrophoresis of PCR products amplified from the individual bands (left), and the same products digested with Dra I or Rsa I (right).
Fig. 3. DGGE fingerprinting of ITS2 rDNA sequence variation in seven species of Trichostrongylus. Lanes 1, T. axei (Ta2); lane 2, T. colubriformis (Tc3); lane 3, T. vitrinus (Tv1); lane 4, T. probolurus; lane 5, T. rugatus; lane 6, T. retortaeformis and lane 7, T. tenuis.
DISCUSSION Separation of DNA in DGGE is based on the electrophoretic mobility of a partially melted molecule in polyacrylamide gels, which is decreased compared with that of the helical form of the molecule. The melting of fragments proceeds in discrete melting domains, which are regions of sequence with an identical melting domain. Once the melting domain with the lowest melting temperature is at a particular position in the gel, a transition of helical to partially melted molecules occurs, and the migration of the molecules will almost stop. Sequence variation within such domains causes the melting temperatures of the variant (mutant) molecules to differ. As a consequence, sequence variants will stop migrating at different positions in the denaturing gradient and will be separated. DGGE can be used to analyse genomic DNA, but recently the technique has been adapted to PCR products in order to analyse sequences of interest. A modification to the latter approach has been to incorporate a GC-rich sequence into one
of the primers used in PCR, which alters the melting behaviour of the fragment and permits the separation of almost 100% of variant molecules.24 Based on the results of the present study, we have demonstrated that DGGE is a powerful analytical technique to screen for sequence variation in parasite rDNA. The technique displays visually the different sequence types of ITS2 in one step. Given that different levels of mutational variation were observed among the species examined and similar (or the same) levels within a species, the technique allowed a diagnostic fingerprint of a species to be generated using a defined rDNA region. This feature should be of particular use for the species identification of individual developmental stages of parasites where morphological characters are unreliable, and therefore has important implications for diagnosis of infection(s). Given that we have already shown that ITS2 can be amplified from single worms, eggs and larvae of a range of strongylid nematodes,13,15,16,22 this approach has formidable potential for the identification of these stages without the need for restriction endonuclease analysis or DNA sequencing. The DGGE technique also has valuable preparative qualities, in that individual variant sequences (even if in low abundance) can be excised from gels, subsequently reamplified and directly subjected to PCR-RFLP or DNA sequence analysis. In this way, it should be possible to quantitatively determine levels of sequence variation among rDNA repeats within and among parasite species without the need for time-consuming isolation of variant sequences using DNA cloning procedures. Although in this study, DGGE has been used to analyse the ITS2 of parasites of veterinary significance, it has the potential to be applied to a broad range of parasites and other eukaryotic organisms of medical importance.
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ACKNOWLEDGEMENTS This work was supported by the Australian Research Council (no. A19531013), Danish Research Academy (DANVIS no. D930061) and Danish National Research Foundation. Thanks to Henrik Bøgh and Birgitte Sønderby for their support. We are also grateful to H. Hoste, C. Chartier (France), I. Beveridge (Melbourne), M. O’Callaghan (South Australia), P. Hudson (Inverness) and R. Coop (Edinburgh) who assisted in collecting or provided parasite isolates.
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