Benchmarks served difference in efficiency of amplification between nuclear and mitochondrial targets, depending on whether lysis was used or not, is probably due to the relative ease with which quantities of DNA from rupturing mitochondria are released, as compared with the release of more sequestered DNA from the cell nucleus. REFERENCES 1.Barstead, R.J. and R.H. Waterston. 1991. Vinculin is essential for muscle function in the nematode. J. Cell. Biol. 114:715-724. 2.Brown, T.A. 1998. Genetics a Molecular Approach. Chapman and Hall, London. 3.Coffroth, M.A. and J.M. Mulawka III. 1995. Identification of marine invertebrate larvae by means of PCR-RAPD species-specific markers. Limnol. Oceanogr. 40:181-189. 4.Côrte-Real, H.B.S.M., D.R. Dixon and P.W.H. Holland. 1994. Intron-targeted PCR: a new approach to survey neutral DNA polymorphism in bivalve populations. Mar. Biol. 120:407-413. 5.Côrte-Real, H.B.S.M., P.W.H. Holland and D.R. Dixon. 1994. Inheritance of a nuclear DNA polymorphism assayed in single bivalve larvae. Mar. Biol. 120:415-420. 6.Crossland, S., D. Coates, J. Grahame and P.J. Mill. 1993. Use of random amplified polymorphic DNAs (RAPDs) in separating two sibling species of Littorina. Mar. Ecol. Prog. Ser. 96:301-305. 7.Evans, B.S., R.W.G. White and R.D. Ward. 1998. Genetic identification of asteroid larvae from Tasmania, Australia, by PCR-RFLP. Mol. Ecol. 7:1077-1082. 8.Gasser, R.B., N.B. Chilton, H. Hoste and I. Beveridge. 1993. Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Res. 21:2525-2526. 9.Geller, J.B., J.T. Carlton and D.A. Powers. 1994. PCR-based detection of mtDNA haplotypes of native and invading mussels on the northeastern Pacific coast: latitudinal pattern of invasion. Mar. Biol. 119:243-249. 10.Higuchi, R. 1989. Simple and rapid preparation of samples for PCR, p. 31-38. In H.A. Erlich (Ed.), PCR Technology. Stockton Press, NY. 11.Hull, S.L., J. Grahame and P.J. Mill. 1996. Morphological divergence and evidence for reproductive isolation in Littorina saxatilis (Olivi) in northeast England. J. Mol. Stud. 62:89-99. 12.Mikhailova, N. and K. Johannesson. 1998. A comparison of different protocols for RAPD analysis of Littorina. Hydrobiologia 378:33-42. 13.Reid, D.G. 1996. Systematics and Evolution of Littorina. The Ray Society, London. 14.White, L.R., B.A. McPheron and J.R. Stauffer, Jr. 1994. Identification of freshwater mussel glochidia on host fishes using restriction fragment length polymorphisms. Mol. Ecol. 3:183-185. 15.Wilson, A.C., R.L. Cann, S.M. Carr, M. George, U.B. Gyllensten, K.M. Helmby1052 BioTechniques
chowski, R.G. Higuchi, S.R. Palumbi, E.M. Prager, R.D. Sage and M. Stoneking. 1985. Mitochondrial DNA and two perspectives on evolutionary genetics. Biol. J. Linn. Soc. 26:375-400. 16.Wray, C.G., J.J. Lee and R. DeSalle. 1993. Extraction and enzymatic characterization of foraminiferal DNA. Micropaleontology 39:69-73.
This study was funded in part by Contract No. MAS3-CT95-0042 from the European Union. We thank the School of Biology at the University of Leeds for the use of their facilities and Dr. I.A. Hope and his research group for their assistance. Address correspondence to Mr. Robert J. Simpson, School of Biology, The University of Leeds, Leeds, UK. Internet:
[email protected] Received 19 November 1998; accepted 1 March 1999.
Robert J. Simpson, Craig S. Wilding and John Grahame The University of Leeds Leeds, UK
Extracting High-Quality DNA from Shed Reptile Skins: A Simplified Method BioTechniques 26:1052-1054 (June 1999)
Molecular studies involving reptiles often overlook shed skins as a source for high-quality DNA. In most cases, tissues or blood samples are preferred by researchers, but the process of sampling for these tissue types can be harmful or otherwise adversely affect the animals involved. While reptile breeders or zoological institutions are a potential source of specimens, most will likely decline requests for samples if the sampling will harm their prized animals. Generally, breeders and curators would be much more amenable to part with a shed skin—which is something they usually discard anyway.
Figure 1. Agarose gel depicting the quality of the genomic DNA extractions produced by this method (lanes 1–5 and 13). Subsequent cytochrome-b PCRs generated from these extractions (lanes 7–11 and 14) using the LGL765 primer (5′-GAAAAACCAYCGTTGTWATTCAACT-3′) of Bickham et al. (2) and the H15919 primer (5′-GACCCAKCTTTGRYTTACAAGGACAA-3′) from this study are also shown. These primers produce a product that is approximately 1160 bp in length. Extractions from the different species are as follows: L. mexicana, lane 1; L. g. floridana, lane 2; L. p. pyromelana, lane 3; L. alterna, lane 4; L. z. zonata, lane 5. Lane 6 contains a 1-kb DNA Ladder (Life Technologies). Lanes 7–11 contain the same species (in the same order) as lanes 1–5. Lane 12 contains a 100-bp DNA ladder (Life Technologies). Lanes 13 and 14 contain extracted DNA and resulting PCR product, respectively, from the day gecko P. madagascariensis grandis. Vol. 26, No. 6 (1999)
The ability to extract DNA from a shed skin found in the field (assuming the species can be identified) is also of use when trying to collect samples of rare, elusive or endangered species. One investigator showed that sufficient quantities of DNA for polymerase chain reaction (PCR) amplification can be obtained from shed skins that have been exposed to the elements (4). Results of the present study show that sufficient quantities of DNA can also be obtained from sheds that are several months old. In the past few years several methods (1,3–5) have been proposed for extracting DNA from shed reptile skins. Although these methods will work, they are either labor-intensive or involve potentially harmful or hazardous chemicals. In this paper, I describe a relatively quick, inexpensive and nonhazardous method of extracting DNA from shed reptile skins that yields sufficient quantities of purified DNA for use in PCR. Shed snake skins for this study were obtained from two private collectors and included the following five Lampropeltis species: L. mexicana, L. getula floridana, L. p. pyromelana, L. alterna and L. z. zonata. Sheds were collected from cages, placed in individual Ziploc bags and shipped to the
laboratory for analysis. All skins were allowed to dry completely before shipment and then kept at room temperature. Also included in the study was a shed skin from a day gecko (Phelsuma madagascariensis grandis). The DNA extraction process described in Table 1 is a slightly modified version of the Puregene DNA Isolation Kit, which is commercially available from Gentra Systems (Minneapolis, MN, USA). Extractions were conducted in 1.5-mL microcentrifuge tubes, but are easily scaleable to larger volumes (up to 15 mL, if necessary) with only a slight increase in cost per sample. The DNA obtained by this extraction protocol is of high quality (see Figure 1, lanes 1–5 and 13) and can be used directly in many applications such as PCR, restriction enzyme digestion and sequencing. The extractions that I have conducted on shed skins using this method (n >75) have consistently yielded high-quality DNA and tend to produce high yields in PCR (Figure 1, lanes 7–11). The resulting PCR products (Figure 1, lanes 7–11 and 14) have been used in sequencing reactions performed with an ABI PRISM 377XL DNA Sequencer (PE Biosystems, Foster City, CA, USA) and have resulted in
Color Photo - For position only
Figure 2. Example of sequence data generated on an automated sequencer using the DNA extraction protocol described here. Sequences were generated on an ABI 377XL sequencer using the LGL765 cytochrome-b primer of Bickham et al. (2). Sequences shown are from three different regions of the electrophoretogram out to 700 bp. The data depicted are from the L. mexicana sample as described in the text. Vol. 26, No. 6 (1999)
BioTechniques 1053
Benchmarks Table 1. Extraction Protocol for Shed Reptile Skins
1.An approximately 1-inch square piece of shed skin (cut up) is placed into 900 µL of cell lysis buffer (10 mM Tris-base, 100 mM EDTA, 2% sodium dodecyl sulfate [SDS], pH 8.0) along with 9 µL of proteinase K (20 mg/mL). The sample is mixed thoroughly and then placed in a dry heat block (or water bath) at 55°C for several hours to overnight, with occasional mixing for the first few hours. 2.The sample is then removed from the heat block and cooled to room temperature (Note: the skin does not dissolve after proteinase K digestion, but the DNA is released into solution). Once the sample reaches room temperature, 4 µL of RNase A (10 mg/mL) are added and the sample is mixed and placed in a 37°C water bath for 1 h. 3.The sample is then cooled to room temperature again, and 300 µL of 7.5 M ammonium acetate are added. The sample is then vortex mixed for 10 s and placed on ice for 10–15 min. 4.The sample is then removed from the ice and centrifuged at top speed (ca. 13–14k rpm) for 3 min to pellet the skin remnants, SDS and cell debris. (Sometimes the skin remnants do not pellet properly, so the following step is needed.) 5.As much of the supernatant as possible is drawn off and transferred to a new 1.5-mL microcentrifuge tube, which is then centrifuged at top speed for an additional 2–3 min to pellet any remaining particles. 6.The supernatant from the second spin is then transferred to a new 2-mL tube containing 900 µL of isopropanol and inverted gently about 20× to mix and pellet the DNA. Depending on whether a pellet is observed after addition of the supernatant to the isopropanol, the sample can either be placed at -20° C overnight or centrifuged immediately at top speed for (16 000× g) 2 min to pellet the DNA to the bottom of the tube. 7.After centrifugation, the isopropanol is poured off, and the pellet is washed with 500 µL of 70% ethanol. The sample is then centrifuged as before for 2 min, the ethanol is removed and then the sample is placed in a vacuum concentrator (or the tube can be inverted at room temperature for 15–20 min) until all traces of ethanol have evaporated. 8.The DNA pellet is then resuspended in 30–100 µL of TE buffer (10 mM Trisbase, 0.1 MM EDTA, pH 8.0), depending on the size of the pellet.
high-quality (low background) sequences that consistently have read lengths of over 700 bp (Figure 2). In several cases, I have extracted DNA from shed skins kept at room temperature that are almost nine months old (one such extraction is included in this study; see Figure 1, lane 2). It appears that the DNA from this sample is more degraded than the DNA from other recently collected (one month old) samples, but suitable quantities of the fulllength (1160 bp) cytochrome-b PCR product were still obtained from this sample for sequencing purposes (Figure 1, lane 8). This suggests that the DNA is stable for quite some time as long as the shed is kept dry. 1054 BioTechniques
I have also used the above DNA extraction protocol on a variety of different tissue types (e.g., heart, muscle, liver or whole blood) and on various organisms (e.g., snakes, fish and invertebrates) with much of the same success. Additionally, blood samples can be kept in the cell lysis buffer (see Table 1, step 1) for extended periods of time (several weeks) at room temperature without any noticeable degradation of the DNA. This is helpful when collecting specimens in the field without access to liquid nitrogen or other cryopreservation sources. Overall, this method is very versatile, inexpensive, requires little hands-on work and, best of all, does not require overly haz-
ardous chemicals or organic solvents. Additionally, this technique should prove useful to many researchers for a variety of applications. REFERENCES 1.Ausubel, F.M., R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith and K. Sturhl (Eds.). 1995. Current Protocols in Molecular Biology, Vol. 1, p. 1.01-14.8. John Wiley & Sons, NY. 2.Bickham, J.W., C.C. Wood and J.C. Patton. 1995. Biogeographic implications of cytochrome-b sequences and allozymes in sockeye (Oncorhynchus nerka). J. Hered. 80:140144. 3.Bricker, J., L.M. Bushar, H.K. Reinert and L. Gelbert. 1996. Purification of high quality DNA from shed skins. Herpetol. Rev. 27:133134. 4.Clark, A. 1998. Reptile sheds yield high quality DNA. Herpetol. Rev. 29:17-18. 5.Hillis, D.M., C. Moritz and B.K. Mable (Eds.). 1996. Molecular Systematics. 2nd ed. Sinauer Associates, Sunderland, MA.
I would like to thank Matt Smith (High Mountain Reptiles) and Kenney Kristo (University of Florida) for supplying shed skins from their private collections for use in this study. I am also indebted to Keith Crandall and Mike Whiting for the use of lab space, chemicals and equipment during the course of this study. This study was funded by the National Science Foundation (NSF) Grant Nos. IBN9702338 and DEB9615269 to K.C. and M.W., respectively. Address correspondence to Mr. James W. Fetzner, Jr., Department of Zoology, Brigham Young University, 574 Widtsoe Building, Provo, UT 84602-5255, USA. Internet:
[email protected] Received 21 December 1998; accepted 16 March 1999.
James W. Fetzner, Jr. Brigham Young University Provo, UT, USA
Vol. 26, No. 6 (1999)
