SCIENTIFIC CORRESPONDENCE indicators to locate possible gas hydrate zones in the deep continental margins. The association of gas hydrates with submarine mud volcanoes has been recorded in the Adriatic Sea17, Caspian Sea18, Norwegian Sea8, off Barbados19 and Blake Ridge, off USA9. Milkov1 has summarized the association of submarine mud volcanoes and gas hydrates in all the oceans of the world. The above documentations suggest that the submarine mud volcanoes (diapirs) are also reliable indicators of the subsurface occurrence of gas hydrates. In India, the hydrocarbon potential is related to the basin and sedimentary evolution of Arabian Sea, Bay of Bengal and their north-eastern extensions. Since 1950s, the detailed petroleum exploratory work has led to the discovery of several new petroliferous sedimentary basins around India, both onshore and offshore, with good accumulations of oil and natural gas of commercial value. However, the exploration for marine gas hydrates around India is not yet intensified. The eastern continental margin of India adjacent to the major sedimentary basins, viz. Cauvery, Krishna and Godavari is presumed to have great potential for gas hydrate deposits20. Compilation of available data on bathymetry, seabottom temperature and geothermal gradient of continental margin of India has indeed helped to bring out maps depicting the probable thickness of gas hydrate stability zone (GHSZ) and sea-bottom temperature variation21. These maps greatly facilitated preliminary assessment about the depth of occurrence of gas hydrates and clues to search for BSR and its depth around the Indian continental margin. However, a systematic seismic survey in

the offshore areas of India is necessary to confirm the depth of BSR. It is interesting to note that the zones of mud diapirs reported off both Point Calimere and Saurashtra – Mumbai fall in the predicted shallow zones of GHSZ. These (Figure 1) are geologically potential zones for gas hydrates. Collection of additional geoscientific data is now recommended through (i) 3.5 kHz echo sounding in these areas, along closely spaced tracks; (ii) side scan sonar survey to pick up the locations of more diapirs; (iii) multibeam hydro-sweep survey to understand the distribution pattern of the mud diapirs; (iv) sampling of the diapirs using gravity or piston corers, and (v) vertical splitting of the cores, logging and methane gas analysis on-board. 1. Milkov, A. V., Mar. Geol., 2000, 167, 29–42. 2. Sengupta, R., Basu, P. C., Bandyopadhyay, R. R., Bandyopadhyay, A., Rakshit, S. and Sharma, B., Geol. Surv. India Spl. Publ., 1992, 29, 201–207. 3. Reddy, D. R. S., Rao, T. S., Rao, R. B. S. and Biswas, N. R., Geol. Surv. India, Mar. Wing Newsl., 1993, IX, 5–6. 4. Anon., Unpublished Report, Marine Wing, Geol. Surv. India, 1997. 5. Ujiie, Y., Mar. Geol., 2000, 163, 149– 167. 6. Sastri, V. V., Raju, A. T. A., Sinha, R. N., Venkatachala, B. S. and Banerjee, R. K., J. Geol. Soc. India, 1977, 18, 355– 377. 7. Biswas, S. K., Mar. Geol., 1987, 82, 285–291. 8. Bouriak, S., Vanneste, M. and Saoutkine, A., Mar. Geol., 2000, 163, 125–148. 9. Taylor, M. H., Dillon, W. P. and Pecher, I. A., Mar. Geol., 2000, 164, 79–89. 10. Sloan, E. D., Clathrate Hydrates of Natural Gases, Marcel Dekker, NY, 1990.

11. Kvenvolden, K. A., Chem. Geol., 1988, 71, 41–51. 12. Kvenvolden, K. A., Ginsburg, G. D. and Soloviev, V. A., Geo-Mar. Lett., 1993, 13, 32 –40. 13. Hyndman, R. and Davis, E., J. Geophys. Res., 1992, 97, 7025–7041. 14. Brown, K. M., Earth Planet. Sci. Lett., 1996, 139, 471–483. 15. Ginsburg, G. D. and Soloviev, V. A., Mar. Geol., 1997, 137, 49–57. 16. Vogt, P. R., Gardner, J. and Crane, K., Geo-Mar. Lett., 1999, 19, 2–21. 17. Hovland, M. and Curzi, P. V., Mar. Petrol. Geol., 1989, 6, 161–167. 18. Ginsburg, G. D. and Soloviev, V. A., Bull. Geol. Soc. Den., 1994, 41, 95–100. 19. Martin, J. B., Kastner, M., Hendry, P., Le Pichon, X. and Lallemant, S., J. Geophys. Res., 1996, 101, 20325–20345. 20. Sankaran, A. V., Curr. Sci., 1998, 74, 281–282. 21. Hanumantha Rao, Y., Reddy, S. I., Ramesh Khanna, Rao, T. G., Thakur, N. A. and Subramanyam, Y., Curr. Sci., 1998, 74, 466–468.

ACKNOWLEDGEMENTS. I thank the Deputy Director-General, Marine Wing, GSI, Kolkata for permission to publish this paper. Thanks are also due to Dr P. Prabhakara Rao, Director (CT), Marine Wing, GSI, Visakhapatnam for his critical review of the paper. Received 2 April 2001; revised accepted 9 May 2001

G. GAITAN VAZ Operations: East Coast–II, Marine Wing, Geological Survey of India, 41, Kirlampui Layout, Visakhapatnam 530 017, India

Molecular characterization of Piper nigrum L. cultivars using RAPD markers Black pepper, Piper nigrum L., often referred to as the ‘king of spices’ is the most important spice in the world. The pepper of commerce is the dried, mature berries of P. nigrum. Indian black pepper is known for its quality and fetches premium price in the market. Majority of the present-day Indian cultivars, numbering about 100, are land races representing 246

direct introduction from the wild1. Advanced cultivars have been derived mostly by clonal selection from land races, though a few are of hybrid origin. As India is the primary centre of diversity of P. nigrum, the indigenous genetic resources are reservoirs of useful genes for plant improvement programmes. Unambiguous characterization of cultivars

and selected germplasm is an urgent requirement in tune with the globalization of agriculture. Molecular markers offer means of identifying cultivars with much greater reliability than the morphological traits which are governed by complex genetic interactions2. Of the several PCR-based techniques, random amplified polymorphic DNA

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SCIENTIFIC CORRESPONDENCE (RAPD) technique3,4 offers a simple and economical means for rapid identification of a large number of accessions. The information obtained through germplasm characterization using RAPD is extensively used for the identification of germplasm, screening of duplicates, assessing genetic diversity and monitoring the genetic stability of conserved germplasm. In the genus Piper, the technique has been successfully utilized in identifying somaclonal variants of Piper longum L5. The present study is a report on the DNA fingerprinting of P. nigrum cultivars. Thirteen land races and nine advanced cultivars of P. nigrum were used in the present study. DNA was extracted according to the CTAB method with minor modifications. Leaf tissue of 5 g was ground to fine powder in liquid nitrogen, followed by incubation in 15 ml of pre-heated extraction buffer (4% w/v cTAB; 1.4 M NaCl; 100 mM Tris-HCl, pH 8; 20 mM EDTA; 2% PVP, and 0.1% v/v β-mercaptoethanol) for 2 h at 55°C. The homogenate was extracted once with chloroform : iso-amyl alcohol (24 : 1) and centrifuged at 15000 rpm at 25°C. Nucleic acids were precipitated in 0.6 volumes of iso-propanol and collected by centrifugation (15000 rpm, 15 min, 25°C). The precipitate was washed with 70% ethanol, air-dried and dissolved in TE buffer (10 mM Tris HCl, pH 8; 1 mM EDTA). DNA was treated with bovine pancreatic RNase and extracted with phenol : chloroform (1 : 1) and chloroform : iso-amyl alcohol (24 : 1) in succession. The purified DNA was quantified in a fluorometer and dissolved to appropriate dilution in TE buffer. The components of the amplification reaction were optimized and a typical 25 µl PCR mixture comprised 20 ng genomic DNA; 2.5 µl 10 × assay buffer; 2 mM MgCl2; 200 µM each of dATP, dGTP, dCTP and dTTP; 0.75 u Taq DNA polymerase and 1 µM primer. PCR reactions were carried out on a Perkin Elmer 9600 DNA thermal cycler. After a predenaturation step of 4 min at 94°C, amplification reactions were cycled 40 times at 94°C for 1 min, 35°C for 1 min and 72°C for 2 min followed by 5 min at 72°C. Amplification products were separated by electrophoresis on 1.2% agarose gels in 1 × TBE buffer. Same size bands across accessions were treated as identical markers. Polymorphism was confirmed

Table 1.

Cultivar-specific bands generated by random primers in Piper nigrum L.

Cultivar

Primer

Sequence

Band size (bp)

Panniyur-1 Panniyur-2 Panniyur-2 Panniyur-4 Panniyur-4 Panniyur-4 Panniyur-5 Panniyur-5 Sreekara Sreekara Subhakara Panchami Pournami Pournami Pournami Cheriakaniakadan Cheriakaniakadan Karimunda Karimunda Malligeswara

OPA08 OPA08 OPB20 OPA10 OPO16 OPU17 OPA10 OPG11 OPA15 OPA15 OPM04 OPA10 OPA05 OPE12 OPF09 OPF09 OPW11 OPA05 OPO16 OPF09

GTGACGTAGG GTGACGTAGG GGACCCTTAC GTGATCGCAG TCGGCGGTTC ACCTGGGGAG GTGATCGCAG TGCCCGTCGT TTCCGAACCC TTCCGAACCC GGCGGTTGTC GTGATCGCAG AGGGGTCTTG TTATCGCCCC CCAAGCTTCC CCAAGCTTCC CTGATGCGTC AGGGGTCTTG TCGGCGGTTC CCAAGCTTCC

850 500 500 500 550 700 850 1400 2400 2500 550 3000 1500 400 1800 850 300 2000 1031 2400

a

b

Figure 1. RAPD profile of P. nigrum accessions obtained with primer OPA08. a, Lanes represent 1, Wayanadan; 2, Uthiramkotta; 3, Panniyur-1; 4, Cheriakaniakadan; 5, Balankotta; 6, Panniyur-2; 7, Panniyur-4; 8, Perumkody; 9, Panniyur-5; 10, P. longum; 11, Karimunda; 12, Sreekara; b, Lanes represent 13, Subhakara; 14, Aimpiriyan; 15, Panchami; 16, P. colubrinum; 17, Malligeswara; 18, Cholamundi; 19, Dodiga; 20, Sumandy; 21, Kalluvally; 22, Karimunda; 23, Pournami; 24. Panniyur-3.

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SCIENTIFIC CORRESPONDENCE by repeating the experiment at least once. A total of 34 decamer primers (series OPA, OPB etc.) from Operon Tech, CA, USA were screened using three representative genomic DNA samples of black pepper. Of these, 24 primers that yielded consistent and clear band patterns were selected for the final analysis of the 22 accessions used in the present study. The 24 selected 10-mer primers generated 372 amplification products; the total number of markers per primer ranged from 4 (OPV05) to 21 (OPF09). The range of polymorphic markers per primer was 3 (OPV05) to 21 (OPF09), with a mean of 15.3 polymorphic bands per primer. Cultivar-specific bands were obtained for all the released varieties except Panniyur-3 (Table 1). Panniyur-1 and Panniyur-2 generated specific bands with OPA 08 (Figure 1 a and b). OPA10 was efficient in differentiating three varieties, viz. Panniyur-4, Panniyur-5 and Panchami.

Among the land races, cultivar-specific bands were observed for Cheriakaniakadan, Malligeswara and Karimunda. Panniyur-1 is the most popular pepper variety grown in India, while land races Cheriakaniakadan and Karimunda are extensively used for pepper improvement work. The utility of the specific RAPD markers can be increased by sequencing their termini and designing longer primers (SCARS) for specific amplification.

1. Ibrahim, K. K., Pillay, V. S. and Sasikumaran, S., Indian Spices, 1984, 21 and 22, 3–9. 2. Karp, A., Issar, P. G. and Ingram, D. S., Molecular Techniques for Screening Biodiversity, Chapman & Hall, London, 1998. 3. Welsh, H. and Mccleand, M., Nucleic Acids Res., 1990, 19, 303–306. 4. Williams, J. G. K., Kubelik, A. R., Livak, K. J., Rafalski, J. A. and Tingey, S. A., Nucleic Acids Res., 1990, 18, 6531–6535. 5. Parani, M., Anand, A. and Parida, A., Curr. Sci., 1997, 73, 81–83.

ACKNOWLEDGEMENTS. We thank Dr P. L. Gautam, National Director, NATP and Dr G. D. Sharma, Director, NBPGR for providing facilities to undertake this study under NATP Team of Excellence project.

Received 11 September 2000; revised accepted 16 April 2001

T. PRADEEPKUMAR†,* J. L. KARIHALOO# SUNIL ARCHAK# †

Regional Agricultural Research Station, Ambalavayal, Wayanad 673 593, India # National Research Centre on DNA Fingerprinting, National Bureau of Plant Genetic Resources, New Delhi 110 012, India *For correspondence. e-mail: Kumartpradeep@ usa.net

Determination of oxalate in foodstuffs with arylamine glass-bound oxalate oxidase and peroxidase In plants, oxalic acid is derived from oxidative breakdown of carbohydrate and protein. Edible plant tissues contain considerable amount of soluble oxalate, some of which forms an insoluble salt with calcium and thus makes the calcium unavailable in the diet. Patients with calcium oxalate urolithiasis are advised by clinicians to avoid an oxalate-rich diet, as high consumption of oxalate leads to a high level of oxalate in plasma, which might deposit in the kidney as insoluble calcium oxalate to form urinary stones1. Hence the measurement of oxalate in foodstuffs is very important. Different methods have been used to measure oxalate in foodstuffs such as permanganate titrimetry2, colorimetry3, capillary electrophoresis4, spectrophotometry5, GLC3, HPLC6, isotachophoresis7 and ion chromatography8. Of these methods, the first two are non-specific and less accurate, while the rest require expensive apparatus and trained personnel to operate. Although an enzymatic colorimetric method employing oxalate oxidase and peroxidase is simple, sensitive, and spe248

cific and does not require costly equipment9, it becomes expensive for a large number of samples, due to the requirement of bulk quantity of enzyme. Immobilization of enzyme on insoluble support provides its reuse and thus reduces the cost of procedure. Recently, we have reported the immobilization of barley oxalate oxidase onto alkylamine glass beads through glutaraldehyde coupling, for determination of oxalate in a few foodstuffs10. However, the immobilization of an enzyme on alkylamine glass through glutaraldehyde coupling has the disadvantage of extensive self-polymerization nature of glutaraldehyde and protein crosslinking, and hence the support needs to be washed well prior to enzyme addition11. Further, the glutaraldehyde coupling involves Schiff’s base formation, which has a drawback of reversibility of the reaction at low pH12. These problems were overcome by immobilizing oxalate oxidase and peroxidase onto arylamine glass beads through diazotization13,14. The present report describes the use of arylamine glass-bound enzymes for deter-

mination of oxalate in various foodstuffs. Zirconia-coated arylamine glass beads (pore diameter 55 nm) (Corning Glass Works, New York); horseradish peroxidase (RZ = 1.1), oxalic acid, 4-aminophenazone (Sigma Chemicals Co, USA), DEAE-Sephacel and Sephadex G-200 (Pharmacia LKB, Sweden) were used. All other chemicals were of analytical reagent grade. The various foodstuffs analysed in the present study were purchased from a local market. Ten-day-old seedling plants of grain sorghum hybrid (CSH-5) were raised in the laboratory and their leaves were collected as described by Pundir and Nath15. The extraction and purification of oxalate oxidase from sorghum leaves were carried out at 4°C (ref. 16). The leaves were homogenized in cold distilled water in 1 : 3 ratio (w/v) in a chilled mortar with pestle and the homogenate was centrifuged at 15,000 × g for 30 min. The yellowishbrown supernatant was collected and treated as crude enzyme. The crude enzyme was purified by 0–80% (NH4)2SO4 precipitation, ion exchange chromato-

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