Hostname: page-component-cd9895bd7-gbm5v Total loading time: 0 Render date: 2024-12-27T02:08:08.612Z Has data issue: false hasContentIssue false
  • English
  • Français

Identification of Balanus amphitrite larvae from field zooplankton using species-specific primers

Published online by Cambridge University Press:  06 November 2014

Chetan C. Gaonkar ,
Lidita Khandeparker ,
Dattesh V. Desai  and
Arga Chandrashekar Anil
Chetan C. Gaonkar
Affiliation:
National Institute of Oceanography, Council of Scientific and Industrial Research, Dona Paula, Goa 403 004, India
Lidita Khandeparker*
Affiliation:
National Institute of Oceanography, Council of Scientific and Industrial Research, Dona Paula, Goa 403 004, India
Dattesh V. Desai
Affiliation:
National Institute of Oceanography, Council of Scientific and Industrial Research, Dona Paula, Goa 403 004, India
Arga Chandrashekar Anil
Affiliation:
National Institute of Oceanography, Council of Scientific and Industrial Research, Dona Paula, Goa 403 004, India
*
Correspondence should be addressed to:L. Khandeparker, National Institute of Oceanography, Council of Scientific and Industrial Research, Dona Paula, Goa 403 004, India email: [email protected]
Get access Rights & Permissions [Opens in a new window]

Abstract

Identification of marine invertebrate larvae using morphological characters is laborious and complicated by phenotypic plasticity. Balanus amphitrite is a dominant barnacle, important in the context of intertidal ecology and biofouling of manmade structures. Morphological identification of barnacle larval forms in a mixed population is difficult because of their intricacy and similarity in size, shape and developmental stages. We report the development and application of a nucleic acid-based Polymerase Chain Reaction (PCR) method for the specific identification of the barnacle, B. amphitrite, from the heterogeneous zooplankton sample. This method is reliable and accurate thereby overcoming taxonomic ambiguity. Sequence alignment of the 18S rRNA gene region of selected species of barnacles allowed the design of B. amphitrite-specific PCR primers. Assay specificity was evaluated by screening DNA obtained from selected species of barnacles. The oligonucleotide primers used in the study flanked a 1600 bp region within the 18S rRNA gene. The primer is specific and can detect as few as 10 individuals of B. amphitrite larvae spiked in a background of ~186 mg of zooplankton. This technique facilitates accurate identification and the primer can be used as a marker for enumeration of B. amphitrite larvae in the plankton.

Keywords

BarnacleBalanus amphitriteLarvaeZooplankton18S rRNA
Type
Research Article
Information
Journal of the Marine Biological Association of the United Kingdom , Volume 95 , Issue 3 , May 2015 , pp. 497 - 502
Copyright
Copyright © Marine Biological Association of the United Kingdom 2014 

Access options

Get access to the full version of this content by using one of the access options below. (Log in options will check for institutional or personal access. Content may require purchase if you do not have access.)

References

REFERENCES

Amann, R.I., Krumholz, L. and Stahl, D.A. (1990) Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. Journal of Bacteriology 172, 762–770.Google Scholar
Anil, A.C., Chiba, K., Okamoto, K. and Kurokura, H. (1995) Influence of temperature and salinity on larval development of Balanus amphitrite: implications in fouling ecology. Marine Ecology Progress Series 118, 159–166.CrossRefGoogle Scholar
Baldwin, B.S., Black, M., Sanjur, O., Gustafson, R., Lutz, R.A. and Vrijenhoek, R.C. (1996) A diagnostic molecular marker for zebra mussels (Dreissena polymorpha) and potentially co-occurring bivalves: mitochondrial COI. Marine Molecular Biology and Biotechnology 5, 9–14.Google Scholar
Begum, R.A., Yamaguchi, T. and Watabe, S. (2004) Molecular phylogeny of thoracican barnacles based on the mitochondrial 12S and 16S rRNA genes. Sessile Organisms 21, 47–54.Google Scholar
Bell, J.L. and Grassle, J.P. (1998) A DNA probe for identification of larvae of the commercial surf clam (Spisula solidissima). Marine Molecular Biology and Biotechnology 7, 127–137.Google Scholar
Bertness, M.D., Gaines, S.P., Bermudez, D. and Sandford, E. (1991) Extreme spatial variation in the growth and reproductive output of the acorn barnacle Semibalanus balanoides. Marine Ecology Progress Series 75, 91–100.Google Scholar
Bilodeau, A.L., Lankford, W.S., Kim, T.J., Felder, D.L. and Neigel, J.E. (1999) An ultra sensitive method for detection of single crab larvae (Sesarma reticulatum) by PCR amplification of a highly repetitive DNA sequence. Marine Ecology 8, 683–684.Google Scholar
Bleidorn, C., Vogt, L. and Bartolomaeus, T. (2003) New insights into polychaete phylogeny (Annelida) inferred from 18S rDNA sequences. Molecular Phylogenetics and Evolution 29, 279–288.Google Scholar
Branscomb, E.S. and Vedder, K. (1982) A description of the naupliar stages of the barnacles Balanus glandula Darwin, Balanus cariosus Pallas, and Balanus crenatus Bruguière (Cirripedia, Thoracica). Crustaceana 42, 83–95.Google Scholar
Bucklin, A. (2000) Methods for population genetic analysis of zooplankton. In Harris, R.P., Wiebe, P.H., Lenz, J., Skjoldal, H.R. and Huntley, M. (eds) ICES zooplankton methodology manual. London: Academic Press, pp. 533–570.Google Scholar
Chanley, P. and Andrews, J.D. (1971) Aids for identification of bivalve larvae of Virginia. Malacologia 11, 45–119.Google Scholar
Chen, H.-N., Hoeg, J.T. and Chan, B.K.K. (2013) Morphometric and molecular identification of individual barnacle cyprids from wild plankton: an approach to detecting fouling and invasive barnacle species. Biofouling 29, 133–145.Google Scholar
Clare, A.S., Freet, R.K. and McClary, M. Jr (1994) On the antennular secretions of the cyprid of Balanus amphitrite, and its role as a settlement pheromone. Journal of the Marine Biological Association of the United Kingdom 74, 243–250.CrossRefGoogle Scholar
Coffroth, M.A. and Mulawka, J.M. III (1995) Identification of marine invertebrate larvae by means of PCR-RAPD species specific markers. Limnology and Oceanography 40, 181–189.Google Scholar
Connell, J.H. (1985) The consequences of variation in initial settlement vs. post-settlement mortality in rocky intertidal communities. Journal of Experimental Marine Biology and Ecology 93, 11–45.Google Scholar
Crisp, D.J. (1984) Overview of research on marine invertebrate larvae. In Costlow, J.D. and Tipper, R.C. (eds) Marine biodeterioration: an interdisciplinary study. Annapolis: Naval Institute Press, pp. 103–126.Google Scholar
Deagle, B.E., Bax, N.J., Hewitt, C.L. and Patil, J.G. (2003) Development and evaluation of a PCR-based test for detection of Asterias (Echinodermata: Asteroidea) larvae in Australian plankton samples from ballast water. Marine and Freshwater Research 54, 709–719.Google Scholar
Endo, N., Sato, K., Matsumura, K., Yoshimura, E., Odaka, Y. and Nogata, Y. (2010) Species-specific detection and quantification of common barnacle larvae from the Japanese coast using quantitative real time PCR. Biofouling 26, 901–911.Google Scholar
Fernando, S.A. (2006) Monograph on Indian barnacles. Kochi: OSTC Marine Benthos-02, Ocean Science and Technology Cell, CUSAT, 219 pp.Google Scholar
Flowerdew, M.W. (1985) Indices of genetic identity and distance in three taxa within the Balanus amphitrite Darwin complex (Cirripedia, Thoracica). Crustaceana 49, 7–15.Google Scholar
Frischer, M.E., Danforth, J.M., Tyner, L.C., Leverone, J.R., Marelli, D.C., Arnold, W.S. and Blake, N.J. (2000) Development of an Argopecten-specific 18S rRNA targeted genetic probe. Marine Biotechnology 2, 11–20.Google Scholar
Goffredi, S.K., Jones, W.J., Scholin, C.A., Marin, R. and Vrijenhoek, R.C. (2006) Molecular detection of marine invertebrate larvae. Marine Biotechnology 8, 149–160.Google Scholar
Hare, M.P., Palumbi, S.R. and Butman, C.A. (2000) Single-step species identification of bivalve larvae using multiplex polymerase chain reaction. Marine Biology 137, 953–961.CrossRefGoogle Scholar
Hebert, P.D.N. (2002) Life in polar waters. Integrative and Comparative Biology 42, 1242.Google Scholar
Hebert, P.D.N., Cywinska, A., Ball, S.L. and deWaard, J.R. (2003a) Biological identifications through DNA barcodes. Proceedings of the Royal Society of London B 270, 313–321.Google Scholar
Hebert, P.D.N., Ratnasingham, S. and deWaard, J.R. (2003b) Barcoding animal life: cytochrome c oxidase subunit 1 divergences among closely related species. Proceedings of the Royal Society of London B 270, 596–599.Google Scholar
Henry, D.P. and McLaughlin, P.A. (1975) The barnacles of the Balanus amphitrite complex (Cirripedia, Thoracica). Zoologische Verhandelingen Leiden 141, 1–254.Google Scholar
Hillis, D.M., Mable, B.K. and Moritz, C. (1996) Applications of molecular systematics: the state of the field and a look to the future. In Hillis, D.M., Moritz, C. and Mable, B.K. (eds) Molecular systematics. Sunderland, MA: Sinauer Associates, pp. 515–543.Google Scholar
Holm, E.R. (1990) Attachment behavior in the barnacle Balanus amphitrite amphitrite (Darwin): genetic and environmental effects. Journal of Experimental Marine Biology and Ecology 135, 85–98.Google Scholar
Jones, W.J., Preston, C., Marin, R., Scholin, C.A. and Vrijenhoek, R.C. (2008) A robotic molecular method for in situ detection of marine invertebrate larvae. Molecular Ecology Resources 8, 540–550.Google Scholar
Karande, A.A. (1967) On cirripede crustaceans (barnacles), an important fouling group in Bombay waters. Proceedings of the Symposium on Crustaceans. Marine Biological Association of India 2, 1942–1953.Google Scholar
Khandeparker, L. and Anil, A.C. (2011) Role of conspecific cues and sugars in the settlement of cyprids of the barnacle, Balanus amphitrite. Journal of Zoology 284, 206–214.Google Scholar
Khandeparker, L., Anil, A.C. and Raghukumar, S. (2003) Barnacle larval destination: piloting possibilities by bacteria and lectin interaction. Journal of Experimental Marine Biology and Ecology 289, 1–13.CrossRefGoogle Scholar
Khandeparker, L., Anil, A.C. and Raghukumar, S. (2006) Relevance of biofilm bacteria in modulating the larval metamorphosis of Balanus amphitrite Darwin. FEMS Microbiology Ecology 58, 425–438.Google Scholar
Klinbunga, S., Ampayup, P., Tassanakajon, A., Jarayabhand, P. and Yoosukh, W. (2000) Development of species-specific markers of the tropical oyster (Crassostrea belcheri) in Thailand. Marine Biotechnology 2, 476–484.Google Scholar
Larsen, J.B., Frischer, M.E., Rasmussen, L.J. and Hansen, B.W. (2005) Single-step nested multiplex PCR to differentiate between various bivalve larvae. Marine Biology 146, 1119–1129.Google Scholar
Levin, L.A. (1990) A review of methods for labeling and tracking marine invertebrate larvae. Ophelia 32, 115–144.Google Scholar
Maki, J.S., Rittschof, D., Costlow, J.D. and Mitchell, R. (1988) Inhibition of attachment of larval barnacles, Balanus amphitrite, by bacterial films. Marine Biology 97, 199–206.Google Scholar
MaKinster, J.G., Felder, D.L., Chlan, C., Boudreaux, M. and Neigel, J.E. (1999) PCR amplification of a middle repetitive element detects larval stone crabs (Crustacea: Decapoda: Menippidae) in estuarine plankton samples. Marine Ecology Progress Series 181, 161–168.Google Scholar
Morgan, T.S. and Rogers, A.D. (2001) Specificity and sensitivity of microsatellite markers for the identification of larvae. Marine Biology 139, 967–973.Google Scholar
Nichols, S.J. and Black, M.G. (1994) Identification of larvae: the zebra mussel (Dreissena polymorpha), quagga mussel (Dreissena rosteriformis bugensis), and Asian clam (Corbicula fluminea). Canadian Journal of Zoology 72, 406–417.Google Scholar
Patil, J.G., Gunasekera, R.M., Deagle, B.E. and Bax, N.J. (2005a) Specific detection of Pacific oyster (Crassostrea gigas) larvae in plankton samples using nested polymerase chain reaction. Marine Biotechnology 7, 11–20.Google Scholar
Patil, J.G., Gunasekera, R.M., Deagle, B.E., Bax, N.J. and Blackburn, S.I. (2005b) Development and evaluation of a PCR based assay for detection of the toxic dinoflagellate, Gymnodinium catenatum (Graham) in ballast water and environmental samples. Biological Invasions 7, 983–994.Google Scholar
Peplies, J., Glockner, F.O., Amann, R. and Ludwig, W. (2004) Comparative sequence analysis and oligonucleotide probe design based on 23S rRNA genes of Alphaproteobacteria from North Sea bacterioplankton. Systematic and Applied Microbiology 27, 573–580.Google Scholar
Pitombo, F.B. (2004) Phylogenetic analysis of the Balanidae (Cirripedia, Balanomorpha). Zoologica Scripta 33, 261–276.Google Scholar
Pradillon, F., Schmidt, A., Peplies, J. and Dubilier, N. (2007) Species identification of marine invertebrate early stages by whole larvae in situ hybridisation of 18S ribosomal RNA. Marine Ecology Progress Series 333, 103–116.Google Scholar
Raimondi, P.T. (1991) Settlement behavior of Chthamalus anisopoma larvae largely determines the adult distribution. Oecologia 85, 349–360.Google Scholar
Rittschof, D., Branscomb, E.S. and Costlow, J.D. (1984) Settlement and behavior in relation to flow and surface in larval barnacles, Balanus amphitrite Darwin. Journal of Experimental Marine Biology and Ecology 82, 131–146.Google Scholar
Rogers, A. (2001) Molecular ecology and identification of marine invertebrate larvae. In Atkinson, D. and Thorndyke, M. (eds) Environment and animal development: genes, life histories and plasticity. Oxford: BIOS Scientific Publishers, pp. 29–69.Google Scholar
Shanks, A.L. (1986) Tidal periodicity in the daily settlement of intertidal barnacle larvae and an hypothesized mechanism for the cross-shelf transport of cyprids. Biological Bulletin 170, 429–440.Google Scholar
Simon-Blecher, N., Huchon, D. and Achituv, Y. (2007) Phylogeny of coral inhabiting barnacles (Cirripedia; Thoracica; Pyrgomatidae) based on 12S, 16S and 18S rDNA analysis. Molecular Phylogenetics and Evolution 44, 1333–1341.Google Scholar
Stach, T. and Tubeville, J.M. (2002) Phylogeny of Tunicata inferred from molecular and morphological characters. Molecular Phylogenetics and Evolution 25, 408–428.Google Scholar
Strathmann, R.R., Branscomb, E.S. and Vedder, K. (1981) Fatal errors in set as a cost of dispersal and the influence of intertidal flora on set of barnacles. Oecologia 48, 13–18.Google Scholar
Sutherland, J.P. (1990) Recruitment regulates demographic variation in a tropical intertidal barnacle. Ecology 71, 955–972.Google Scholar
Thiyagarajan, V., Venugopalan, V.P., Nair, K.V.K. and Subramoniam, T. (1997a) Larval description of Balanus reticulatus Utinomi (Cirripedia, Balanidae), reared in the laboratory. Journal of Experimental Marine Biology and Ecology 209, 215–231.Google Scholar
Thiyagarajan, V., Venugopalan, V.P., Subramoniam, T. and Nair, K.V.K. (1997b) Description of the naupliar stages of Megabalanus tintinnabulum (Cirrepedia: Balanidae). Journal of Crustacean Biology 17, 332–342.Google Scholar
Thompson, J.D., Gibson, T.J., Plewnaik, F., Jeanmougin, F. and Higgins, D.G. (1997) The Clustal X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Research 25, 4876–4882.Google Scholar
Vadopalas, B., Bouma, J.V., Jackels, C.R. and Friedman, C.S. (2006) Application of real-time PCR for simultaneous identification and quantification of larval abalone. Journal of Experimental Marine Biology and Ecology 334, 219–228.CrossRefGoogle Scholar
Wagh, A.B. and Bal, D.V. (1970) Observations on systematic of sessile barnacles from the west coast of India-1. Journal of the Bombay Natural History Society 71, 109–123.Google Scholar
Webb, K.E., Barnes, D.K.A., Clark, M.S. and Bowden, D.A. (2006) DNA barcoding: a molecular tool to identify Antarctic marine larvae. Deep-Sea Research II 53, 1053–1060.CrossRefGoogle Scholar
Winnepenninckx, B., Backeljau, T. and de Wachter, R. (1995) Phylogeny of protostome worms derived from 18S rRNA sequences. Molecular Biology and Evolution 12, 641–649.Google Scholar
Wood, A., Krogran, N.J., Dover, J., Schneider, J., Heidt, J., Boateng, M.A., Dean, K., Golshani, A., Zhang, Y., Greenblatt, J.F., Johnston, M. and Shilatifard, A. (2003) Bre1, an E3 ubiquitin ligase required for recruitment and substrate selection of Rad6 at a promoter. Molecular Cell 11, 267–274.Google Scholar