SciELO - Scientific Electronic Library Online

 
vol.103 issue5Genetic relationships between sympatric populations of Bacillus cereus and Bacillus thuringiensis, as revealed by rep-PCR genomic fingerprintingSerine protease activities in Oxysarcodexia thornax (Walker) (Diptera: Sarcophagidae) first instar larva author indexsubject indexarticles search
Home Pagealphabetic serial listing  

Memórias do Instituto Oswaldo Cruz

Print version ISSN 0074-0276

Mem. Inst. Oswaldo Cruz vol.103 no.5 Rio de Janeiro Aug. 2008

http://dx.doi.org/10.1590/S0074-02762008000500017 

SHORT COMMUNICATIONS

 

Cheap, rapid and efficient DNA extraction method to perform multilocus microsatellite genotyping on all Schistosoma mansoni stages

 

 

S Beltran; R Galinier; JF Allienne; J Boissier*

UMR 5244 CNRS-EPHE, Parasitologie Fonctionnelle et Evolutive, Biologie et Ecologie Tropical et Méditerranéene, Université de Perpignan Via Domitia, 52 Av. Paul Alduy, 66860 Perpignan, France

 

 


ABSTRACT

Schistosomes are endoparasites causing a serious human disease called schistosomiasis. The quantification of parasite genetic diversity is an essential component to understand the schistosomiasis epidemiology and disease transmission patterns. In this paper, we propose a novel assay for a rapid, low costly and efficient DNA extraction method of egg, larval and adult stages of Schistosoma mansoni. One euro makes possible to perform 60,000 DNA extraction reactions at top speed (only 15 min of incubation and 5 handling steps).

Key words: Schistosoma mansoni - DNA extraction - microsatellites


 

 

Schistosomes (Plathyhelminth, Digenea) are endo-parasites causing a serious human disease called schistosomiasis. Schistosomiasis ranks second only to malaria in terms of parasite-induced human morbidity and mortality, with more than 200 million people infected (Crompton 1999, Chitsulo et al. 2000). The quantification of parasite genetic diversity is an essential component to understand schistosomiasis epidemiology and disease transmission patterns. This genetic diversity could be assessed either at the adult stage (Theron et al. 2004) or, more recently, at the larval stage (Shrivastava et al. 2005, Sorensen et al. 2006). The use of adult worms to quantify the genetic diversity in the definitive host is only relevant when worms can be directly recovered from naturally infected rodents (Theron et al. 2004). The quantification of parasite genetic diversity from intra-human (Brouwer et al. 2001, Curtis et al. 2002, Stohler et al. 2004) or intra-mollusk stages (Dabo et al. 1997, Eppert et al. 2002, Sire et al. 2001) requires a long time for a passaging through experimental hosts. However, mollusk or vertebrate experimental host may induce a bias due to this host selective pressure. Indeed, such laboratory passage may be predicted to result in genetic bottlenecking of the parasite population and impose selection pressures not encountered in field conditions. Firstly, exposure of individual snails to single miracidia results in only 5-50% of successful infections, depending on the parasite strain used (Theron et al. 1997), thus between 50% and 95% of the parasite genetic diversity is lost. Secondly, as far as the vertebrate host is concerned, it has been shown that passaging through experimental models decreases the parasite genetic diversity in comparison to field isolates (Loverde et al. 1985). To circumvent ethical, technical and epidemiological disadvantages of the use of experimental hosts, methods for genotyping larvae have been recently proposed (Shrivastava et al. 2005, Sorensen et al. 2006). Due to their small size (450 µm for cercariae and 150 µm for miracidia), the main technical limitations of these studies were the available quantity of DNA to perform PCR amplifications. In 2005, Shrivastava et al. proposed a DNA extraction protocol allowing sufficient DNA for only one PCR reaction by larvae, thus for only one locus analyses. In 2006, Sorensen et al. proposed a more complex protocol, only tested on eggs and that required liquid nitrogen to disrupt the eggshell by heat shock and Instagen Matrix (Bio-Rad) to capture DNA. This last protocol permits multi-locus analyses but it requires a particular material and finally, the resulting analysis have been performed only on eggs. In this paper, we propose a novel assay for a very rapid, very low costly and efficient DNA extraction method of adult and free larval stages from individual Schistosoma mansoni.

To investigate the efficiency of the method, we have performed DNA extraction of individual schistosome from all life cycle stages (except intra-mollusk stages) and used five microsatellite markers of various sizes (i) on 10 individual eggs derived from faeces of infected mice, (ii) on 10 individual miracidia obtained from eggs purified from the livers of infected mice, (iii) on 10 individual cercariae derived from monomiracidially infected mollusks, (iv) and finally, on 10 adults obtained from infected mice.

The S. mansoni strain was isolated from naturally infected mollusks collected in Guadeloupe (French West Indies) in December 2002. The intermediate host used was a Guadeloupean strain of Biomphalaria glabrata and the definitive host was the Swiss OF1 mouse strain. Detailed methods for the mollusk and mouse infections were previously described (Boissier & Moné 2000). S. mansoni eggs were recovered from faeces of two experimentally infected mice; 10 eggs were individually isolated in 5 µl of NaCl 8% and transferred in a PCR reaction tube using a 20 µl micropipette. Miracidia were hatched from eggs purified from the liver of one infected mouse; 10 miracidia were individually isolated in 5 µl of spring water and transferred in a PCR reaction tube using a 20 µl micropipette. Mollusks were individually exposed to individual miracidium which all originated from the same mouse. Five weeks later, mollusks were individually placed in spring water and exposed to artificial light to stimulate cercarial release; 10 cercariae derived from mollusks were individually isolated in 5 µl of purified spring water and transferred in a PCR reaction tube using a 20 µl micropipette. The presence of only one egg, one miracidium or one cercariae in each respective tube was checked under a binocular microscope. One mouse was infected using 120 cercariae. Seven weeks later, the mouse was sacrificed and 10 worms were recovered and individually isolated.

The same DNA extraction procedure was used, for either adult or larval stages. Before DNA extraction, individual eggs, miracidium, cercariae or adult worms were individually vacuum-dried for 15 min in a Speedvac evaporator. Next, 20 µl of NaOH (250 mM) was added to each tube. After a 15 min incubation period at 25°C, the tubes were heated at 99°C for 2 min. Then, 10 µl HCl (250 mM), 5 µl of Tris-HCl (500 mM) and 5 µl Triton X-100 (2%) were added and a second heat shock at 99°C for 2 min was performed. The products were stored at -20°C. The PCR amplifications were performed in duplicate using five microsatellite markers (Table I). The PCR reactions were carried out in a total volume of 20 µl containing 4 µl of 5X buffer (10 mM Tris-HCl, pH 9.0 at 25°C, 50 mM KCl, 0.1% Triton X-100), 0.2 µM of each oligonucleotide primer, 200 µM of each dNTP (Promega), 1 unit of GoTaq polymerase (Promega, Madison, Wisconsin), 1 µl of extracted DNA and DNase-free water q.s.p. 20 µl. The PCR programme consisted in an initial denaturation phase at 95°C for 5 min, followed by 40 cycles at 95°C for 30 s, annealing temperature for 20 s (Table I), 72°C for 30 s, and a final extension at 72°C for 10 min in a thermocycler (Bio-Rad, Hercules, USA). For each marker, the forward PCR primer was 5' fluorescein labelled (Proligo, Cambridge, UK) allowing a precise analysis in an automated DNA sequencer. The microsatellite PCR products were diluted in sample loading solution (Beckman Coulter, Villepinte, France) with a red labeled size standard (CEQTM DNA size standard kit, 400 Beckman Coulter), and electrophoresed using an automatic sequencer (CEQTM 8000, Beckman Coulter) with CEQTM 8000 sequence analysis software. The sizes of the alleles were calculated with the fragment analyzer package.

From the 10 eggs, we obtained, during the first amplification, 52% of success. A second amplification on the same extracted DNA gave 92% of success. From the 10 miracidia, we obtained, during the first amplification, 90% of success. A second amplification on the same extracted DNA gave 100% of success. From the 10 cercariae, during the first PCR amplifications performed, we obtained 98% of success. A second amplification of the same extracted DNA gave 100% of success. From the 10 adults, during the first PCR amplifications performed, we obtained 98% of success. A second amplification of the same extracted DNA gave 100% of success. The amplification failures were independent from the locus tested. Furthermore, it is likely that 100% of DNA had been extracted because after one or two PCR reactions, all expected PCR products gave at least one result in one microsatellite marker. DNA extraction methods are generally complex and time consuming, or quick and usually more expensive due to the use of commercial kits. Table II shows a comparison between our method and the two previous ones (Shrivastava et al. 2005, Sorensen et al. 2006). Our DNA extraction protocol is efficient on all parasite stages and makes it possible to obtain an extracted DNA for PCR amplification at top speed (15 min incubation), with few handling steps (5) and at a very low cost (1 euro is sufficient to perform more than 60,000 DNA extraction reactions). This extraction procedure yields 40 µl of DNA from individual egg, miracidium, cercaria or adult that allows for 40 PCR amplifications, according to our protocol. This method could be perform in 96-well microplates allowing several hundreds DNA extractions in one hour.

 

ACKNOWLEDGEMENTS

To Valérie Bech, for reading the manuscript and helpful comments, and Bernard Dejean and Pierre Tisseyre, for technical assistance. The experiments comply with the current French laws.

 

REFERENCES

Boissier J, Moné H 2000. Experimental observations on the sex ratio of adult Schistosoma mansoni, with comments on the natural male bias. Parasitology 121: 379-383.         [ Links ]

Brouwer KC, Ndhlovu P, Munatsi A, Shiff CJ 2001. Genetic diversity of a population of Schistosoma haematobium derived from schoolchildren in east central Zimbabwe. J Parasitol 87: 762-769.         [ Links ]

Chitsulo L, Engels D, Montressor A, Savioli L 2000. The global status of schistosomiasis and its control. Acta Trop 77: 41-51.         [ Links ]

Crompton DWT 1999. How much human helminthiasis is there in the world? J Parasitol 85: 397-403.         [ Links ]

Curtis J, Sorensen RE, Kristen Page L, Minchella DJ 2001. Microsatellite loci in the human blood fluke Schistosoma mansoni and their utility for other schistosome species. Mol Ecol Notes 1: 143-145.         [ Links ]

Curtis J, Sorensen RE, Minchella DJ 2002. Schistosome genetic diversity: the implications of population structure as detected with microsatellite markers. Parasitology 125 (Suppl.): S51-59.         [ Links ]

Dabo A, Durand P, Morand S, Diakite M, Langand J, Imbert-Establet D, Doumbo O, Jourdane J 1997. Distribution and genetic diversity of Schistosoma haematobium within its bulinid intermediate hosts in Mali. Acta Trop 66: 15-26.         [ Links ]

Durand P, Sire C, Theron A 2000. Isolation of microsatellite markers in the digenetic trematode Schistosoma mansoni from Guadeloupe island. Mol Ecol 9: 997-998.         [ Links ]

Eppert A, Lewis FA, Grzywacz C, Coura-Filho P, Caldas I, Minchella DJ 2002. Distribution of schistosome infections in molluscan hosts at different levels of parasite prevalence. J Parasitol 88: 232-236.         [ Links ]

Loverde PT, De Wald J, Minchella DJ, Bosshardt SC, Damian RT 1985. Evidence for host-induced selection in Schistosoma mansoni. J Parasitol 71: 297-301.         [ Links ]

Rodrigues NB, Silvia MR, Pucci MM, Minchella DJ, Sorensen R, Loverde PT, Romanha AJ, Oliveira G 2007. Microsatellite-enriched genomic libraries as a source of polymorphic loci for Schistosoma mansoni. Mol Ecol Notes 7: 263-265.         [ Links ]

Shrivastava J, Gower CM, Balolong EJ, Wang TP, Qian BZ, Webster JP 2005. Population genetics of multi-host parasites-the case for molecular epidemiological studies of Schistosoma japonicum using larval stages from naturally infected hosts. Parasitology 131: 617-626.         [ Links ]

Sire C, Langand J, Barral V, Theron A 2001. Parasite (Schistosoma mansoni) and host (Biomphalaria glabrata) genetic diversity: population structure in a fragmented landscape. Parasitology 122: 545-554.         [ Links ]

Sorensen RE, Rodrigues NB, Oliveira G, Romanha AJ, Minchella DJ 2006. Genetic filtering and optimal sampling of Schistosoma mansoni populations. Parasitology 133: 443-451.         [ Links ]

Stohler RA, Curtis J, Minchella DJ 2004. A comparison of microsatellite polymorphism and heterozygosity among field and laboratory populations of Schistosoma mansoni. Int J Parasitol 34: 595-601.         [ Links ]

Theron A, Pages JR, Rognon A 1997. Schistosoma mansoni: distribution patterns of miracidia among Biomphalaria glabrata snail as related to host susceptibility and sporocyst regulatory processes. Exp Parasitol 85: 1-9.         [ Links ]

Theron A, Sire C, Rognon A, Prugnolle F, Durand P 2004. Molecular ecology of Schistosoma mansoni transmission inferred from the genetic composition of larval and adult infrapopulations within intermediate and definitive hosts. Parasitology 129: 571-585.         [ Links ]

 

 

Received 18 September 2007
Accepted 28 July 2008
Financial support: Ministère de l'Enseignement Supérieur et de la Recherche, CNRS

 

 

* Corresponding author: boissier@univ-perp.fr