Evaluation of Chitinase Enzyme Production in Some of the Fusarium Isolates Obtained from Globodera rostochiensis

پژوهش های حفاظت گیاهان ایران

Article 2, Volume 32, Issue 4 - Serial Number 42, December 2019, Pages 463-479   

Document Type: Research Article

DOI: 10.22067/jpp.v32i4.66605

Authors

khadijeh abbasi^{* 1}; Doustmorad Zafari²

¹Bu Ali Sina

²bu ali sina

Abstract

Introduction: The plant parasitic nematodes are among the most important agents causing losses in agricultural crops (Nicol et al. 2013). The golden cyst nematode, Globodera rostochiensis (Wollenweber, 1923; Behrens, 1975) is the most destructive nematode of potato in the world (Brodie, 1984). Chitin is the most abundant polymer in structure of nematode eggshell. Chitinases are widely distributed in fungi and play important roles in degradation of chitin. The aim of this study was to assay chitinase activity in 16 isolates of various species of Fusarium obtained from golden potato cyst nematode, Globodera rostochiensis.
Materials and Methods: In the current study, 16 Fusarium isolates were recovered from infected eggs of the golden potato cyst nematode, G. rostochiensis. These isolates were identified based on morphological (Seifert, 1996) and molecular features including internal transcribed spacer (ITS) regions (ITS1, ITS2 and 5.8S gene) of ribosomal DNA. Total genomic DNA was extracted from lyophilized mycelia with a QIAGEN DNeasy Plant Mini Kit (Germany). ITS regions of ribosomal DNA were amplified with the ITS1 (forward primer) and ITS4 (reverse primer) primers. PCR amplifications were performed with total reaction volume of 25 µl using a Takara EmeraldAmp GT PCR Master Mix. PCR products were separated by electrophoresis technique using 1% agarose gel in 1X Tris-Borate-EDTA (TBE) buffer by adding 12 μl SYBR-safe 10,000X concentrate DNA stain to melted agarose before running the gel and finally visualized under ultraviolet illuminator. The ITS1, ITS2 and 5.8S sequences were obtained by sequencing both strands in opposite directions using the PCR amplification primers, ITS1 and ITS4 in genomics resource laboratory at Massachusetts University, USA. Colloidal chitin was prepared using the procedure of Tikhonov et al. (2002). For enzyme assay in liquid media, the 16 isolates were grown in minimal synthetic medium (MSM) containing colloidal chitin (1 g l-1) based on the method described by Zeilinger et al. (1999). The culture medium was filtered through Whatman paper No. 3 filter followed by filtration through 0.2-mm Millipore polydifluoropropilene membranes. The filtrate obtained was analyzed for chitinolytic activity. Chitinase activity was determined by measuring the release of reducing saccharides from colloidal chitin by the N-acetyl-glucosamine-dinitrosalicylate method according to the method described by Monreal and Reese (1969). Protein concentration was determined according to Bradford (1976) with bovine serum albumin (Sigma) as the standard. Chitinase specific activity was obtained by dividing the enzyme activity rate by total protein mass. Enzyme assay was done to determine the most promising isolates for biological control of G. rostochiensis in 24 and 96 h after fungal growth. The chitinase specific activity data was subjected to analysis of variance (ANOVA) using software SAS, version 9.0 (Statistical Analysis System Institute Inc., USA) in a CRD (completely randomized design) with three replicates.
Two selected isolates were assayed using a chitinase assay, fluorimetric kit-CS1030 (Sigma). The assay included three substrates, 4-Methylumbelliferyl N,N′-diacetyl-β-D-chitobioside, 4-Methylumbelliferyl N-acetyl-β-D-glucosaminide and 4-Methylumbelliferyl β-D-N,N′,N′′-triacetylchitotriose for the detection of the chitinolytic isozymes Chitobiosidase, β-N-acetylglucosaminidase and Endochitinase. The experiment was a randomized complete block design (RCB) with three replicates. Chitinase activity was subjected to analysis of variance using software SAS, version 9.0. Enzyme activity was expressed over time using the Logistic-Peak, software Slide Write, version 2.0. Two isolates (the strongest and weakest ones) were selected to determine the optimum conditions for chitinase production. The measured conditions were pH, temperature and reaction time between enzyme and substrate in a N-acetyl-glucosamine-dinitrosalicylate method. According to measurements taken by an enzyme activity assay kit, the fourth day (96 h) showed the highest activity for the two selected isolates. We then tested our two isolates at pHs of 3, 4, 5, 6, 7, 8 and 9, temperatures of 22°C, 25°C, 28°C, 31°C for differing periods of time (1 h, 6 h, and 24 h). The experiment was based on factorial experiment in a completely randomized design with three replicates. Analysis of variance chitinase activity was performed using SAS 9.0 software.
Results and Discussion: Among 16 isolates, F12 (1/02 U/mg) and F15 (0/04 U/mg) had the maximum and minimum amount of specific activity, respectively. According to morphological features and sequencing of ITS regions (ITS1, ITS2 and 5.8S gene) of ribosomal DNA, these 16 isolates were classified in three species i.e. F. solani, F. oxysporum and F. equiseti. The optimum conditions to produce chitinase in isolates of F12 and F15 were pH=6, 96 hours after fungal growth in 25°C and the reaction between the enzyme and substrate during one hour achieved by the N-acetyl-glucosamine-dinitrosalicylate method. As a producer of various chitinase enzymes, the filamentous fungus, Fusarium spp. seems to be recommendable to biologically control Globodera rostochiensis.

Keywords

Biological control; Enzymatic reaction; Isozyme; Substrate

References

1. Bradford M.M. 1976. A rapid and sensitive method for quantification of microgram. Analytical Biochemestry, 72: 248–254.

2. Brodie B.B. 1984. Nematode parasites of potato. In: Nickle W.R. (ed.): Plant and Insect Nematodes. Marcell Dekker, Inc, New York and Basel, Pp:169–181.

3. Chen S.Y., Dickson D.W., and Whitty E.B. 1996. Fungi associated with egg masses of Meloidogyne incognita and M. javanica in a Florida tobacco field. Nematropica, 26(2): 153–157.

4. Dackman C. 1990. Fungal parasites of the potato cyst nematode Globodera rostochiensis: isolation and reinfection. Journal of Nematology, 22(4): 594–597.

5. Dong J.Y., Zhao Z.X., Cai L., Liu S.Q., Zhang H.R., Duan M., and Zhang K.Q. 2004. Nematicidal effect of fresh water fungal cultures against the pine-nematode, Bursaphelenchus xylophilus. Fungal Diversity, 15: 125–135.

6. Ibrahim G.H., Al Rehiayani S.M., and Bellal M.M. 2009. Use of biocontrol fungi, Bacillus thuringiensis and organic soil amendment to control root-knot nematode (Meloidogyne incognita) in tomato and eggplant. Mansoura University Journal of Agricultural Sciences, 34 (11):10761–10770.

7. Kok C.J., Papert A., and Hok A Hin C.H. 2001. Microflora of Meloidogyne egg masses: species composition, population density and effect on the biocontrol agent Verticillium chlamydosporium. Nematology, 3(8): 729–734.

8. Liu T., Wang L., Duan Y. X., and Wang X. 2008. Nematicidal activity of culture filtrate of Beauveria bassiana against Meloidogyne hapla. World Journal of Microbiology and Biotechnology, 24: 113–118.

9. Manzanilla Lopez R.H., Esteves I., Fintti Sialer M.M., Hirsch P.R., Ward E., Devonshire J., and Hidalgo Diaz L. 2013. Pochonia chlamydosporia: Advances and challenges to improve its performance as a biological control agent of sedentary endo-parasitic nematodes. Journal of Nematology, 45(1): 1–7.

10. Monreal J., and Reese E.T. 1969. The chitinase of Sarratia marcescens. Canadian Journal Microbiology, 15:689–696.

11. Nicol J.M., Turner S.J., Coyne D.L., den Nijs L., Hockland S., and Tahna Maafi Z. 2011. Current nematode threats to world agriculture. In Jones, J., Gheysen, G., and Fenoll, C (Eds.), Genomics and molecular genetics of plant-nematode interactions. Springer Science + Business Media, Pp. 21–43.

12. Pereira L., Noronha E.F., Miller R.N. G., and Franco O.L. 2007. Novel insights in the use of hydrolytic enzymes secreted by fungi with biotechnological potential. Letters of Applied Microbiology, 44: 573–581.

13. Ruanpanun P., Tangchitsomkid N., Hyde KD., and Lumyong S. 2010. Actinomycetes and fungi isolated from plant-parasitic nematode infested soils: screening of the effective biocontrol potential, indole-3-acetic acid and siderophore production. World Journal of Microbiology and Biotechnology, 26: 1569–1578.

14. Safari Motlagh M.R., and Samimi Z. 2013. Evaluation of Trichoderma spp.as biological agents in some of plant pathogens. Annals of Biological Research, 4(3): 173–179.

15. Saifullah and Thomas B.J. 1996. Studies on the parasitism of Globodera rostochiensis by Trichoderma harzianum using low temperature scaning electron microscopy. International Journal of Nematology, 6: 117–122.

16. Saifullah., and Naqibullah khan. 2014. Low temperature scanning electron microscopic studies on the interaction of Globodera rostochiensis woll and Trichoderma harzianum rifai. Pakistan Journal of Botany, 46(1): 357–361.

17. Sankaranarayanan C., Hussaini S. S., Kumar P. S., and Rangeswaran R. 2002. Parasitism of Meloidogyne incognita eggs by Fusarium oxysporum and other fungi. Indian Journal of Nematology, 32: 1 (Abstract).

18. Santos MC., Esteves I., Kerry B., and Abrantes I. 2013. Biology, growth parameters and enzymatic activity of Pochonia chlamydosporia isolates from potato cyst and root-knot nematodes. Nematology, 15: 493–504.

19. Seifert K. 1996. FusKey: Fusarium interactive key. Agriculture and Agri-Food Canada, Research Branch, Eastern Cereal & Oilseed Research Centre.

20. Seyedasli N., Zamani M., Motallebi M., and Harghighi M.J. 2004. The study of chitinase production in Trichoderma fungus. Iran Biology, 17(3): 227–246.

21. Sharon E., Bar Eyal M., Chet I., Herrera E., Strella A., Klelfeld O., and Splege, Y. 2001. Biological control of the root-knot nematode Meloidogyne javanica by Trichoderma harzianum. Phytopathology, 91: 687–693.

22. Sharon E., Chet I., and Spiegel Y. 2009. Improved attachment and parasitism of Trichoderma on Meloidogyne javanica in vitro. European Journal of Plant Pathology, 123: 291–299.

23. Szabo M., Csepregi K., Galber M., Viranyi F., and Fekete C. 2012. Control plant-parasitic nematodes with Trichoderma species and nematode-trapping fungi: the role of chi18-5 and chi18-12 genes in nematode egg-parasitism. Biological Control, 63: 121–128.

24. Tikhonov V.E., Lopez Llorca L.V., Salinas J., and Jansson H.B. 2002. Purification and characterization of chitinases from the nematophagous fungi Verticillium chlamydosporium and V. suchlasporium. Fungal Genetics and Biology, 35: 67–78.

25. Verdejo Lucas S., Ornat C., Sorribas F.J., and Stchiegel A. 2002. Species of root-knot nematodes and fungal egg parasites recovered from vegetables in Almeria and Barcelona, Spain. Journal of Nematology, 34 (4): 405–408.

26. White T.J., Bruns T.D., Lee S., and Taylor J. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis MA., Gelfand D.H., Sninsky J.J., and White T. J (eds) PCR protocols, a guide to methods and applications. San Diego, California: Academic Press, 315–322.

27. Zeilinger S., Galhaup C., Payer K., Woo S.L., Mach R.L., Fekete C., Lorito M., and Kubicek C.P. 1999. Chitinase gene expression during mycoparasitic interaction of Trichoderma harzianum with its host. Fungal Genetics and Biology, 26: 131–140.