Division of methods for counting helminths’ eggs and the problem of efficiency of these methods

Jacek Karamon 4,  
Jacek Malicki 3,  
Institute of Agrophysics, Polish Academy of Sciences, Lublin, Poland
Department of Biological Health Hazards and Parasitology, Institute of Rural Health, Lublin, Poland
Department of Water Supply and Wastewater Disposal, Faculty of Environmental Engineering, Lublin University of Technology, Lublin, Poland
Department of Parasitology and Invasive Diseases, National Veterinary Research Institute, Pulawy, Poland
Ann Agric Environ Med 2017;24(1):1–7
From the sanitary and epidemiological aspects, information concerning the developmental forms of intestinal parasites, especially the eggs of helminths present in our environment in: water, soil, sandpits, sewage sludge, crops watered with wastewater are very important. The methods described in the relevant literature may be classified in various ways, primarily according to the methodology of the preparation of samples from environmental matrices prepared for analysis, and the sole methods of counting and chambers/instruments used for this purpose. In addition, there is a possibility to perform the classification of the research methods analyzed from the aspect of the method and time of identification of the individuals counted, or the necessity for staining them. Standard methods for identification of helminths’ eggs from environmental matrices are usually characterized by low efficiency, i.e. from 30% to approximately 80%. The efficiency of the method applied may be measured in a dual way, either by using the method of internal standard or the ‘Split/Spike’ method. While measuring simultaneously in an examined object the efficiency of the method and the number of eggs, the ‘actual’ number of eggs may be calculated by multiplying the obtained value of the discovered eggs of helminths by inverse efficiency.
Katarzyna Jaromin-Gleń   
Institute of Agrophysics, Polish Academy of Sciences, Lublin, Poland
1. de Silva NR, Brooker S, Hotez PJ, Montresor A, Engels D, Savioli L. Soil-transmitted helminth infections: updating the global picture. Trends Parasitol. 2003; 19(12): 547–551.
2. Bethony J, Brooker S, Albonico M, Geiger SM, Loukas A, Diemert D, Hotez PJ. Soil-transmitted helminth infections: ascariasis, trichuriasis, and hookworm. Lancet. 2006; 367(9521): 1521–1532.
3. Pullan RL, Smith JL, Jasrasaria R, Brooker SJ. Global numbers of infection and disease burden of soil transmitted helminth infections in 2010. Parasites Vector. 2014; 7(37): 1–19. doi: 10.1186/1756-3305-7-37.
4. Palmer LJ, Celedon JC, Weiss ST, Wang B, Fang Z, Xu X. Ascaris lumbricoides infection is associated with increased risk of childhood asthma and atopy in rural China. Am J Resp Crit Care. 2002; 165(11): 1489–1493.
5. Ojha SC, Jaide C, Jinawath N, Rotjanapan P, Baral P. Geohelminths’: public health significance. J Infect Dev Countr. 2014; 8(1): 5–16.
6. van Riet E, Hartgers FC, Yazdanbakhsh M. Chronic helminth infections induce immunomodulation: consequences and mechanisms. Immunobiology. 2007; 212(6): 475–490.
7. World Health Organization. Research Priorities for Helminth Infections. Technical Report of the TDR Disease Reference Group on Helminth Infections.Geneva, 2013.
8. Zeukeng F, Tchinda VHM, Bigoga JD, Seumen CHT, Ndzi ES, Abonweh G, Makoge V, Motsebo A, Moyou RS. Co-infections of malaria and geohelminthiasis in two rural communities of Nkassomo and Vian in the Mfou Health District, Cameroon. PLoS Neglected Tropical Diseases. 2014; 8(10): e3236. doi:10.1371/journal.pntd.0003236.
9. Mantovi P, Baldoni G, Toderi G. Reuse of liquid, dewatered, and composted sewage sludge on agricultural land: effects of long-term application on soil and crop. Water Res. 2005; 39(2–3): 289–296. doi: 10.1016/j.watres. 2004.10.003.
10. Frąc M, Oszust K, Lipiec J, Jezierska-Tys S, Oluchi Nwaichi E. Soil microbial functional and fungal diversity as influenced by municipal sewage sludge accumulation. Int J Environ Res Public Health. 2014; 11: 8891–8908; doi: 10.3390/ijerph110908891.
11. Werle S. Sewage sludge-to-energy management in eastern Europe: a Polish perspective. Ecol Chem Eng S. 2015; 22(3): 459–469. doi: 10.1515/eces-2015–0027.
12. Franus M, Barnat-Hunek D, Wdowin M. Utilization of sewage sludge in the manufacture of lightweight aggregate. Environ Monit Assess. 2016; 188 (1): e10. doi: 10.1007/s10661–015–5010–8.
13. Kozan E, Gonenc B, Sarimehmetoglu O, Aycicek H. Prevalence of helminth eggs on raw vegetables used for salads. Food Control. 2005; 16(3): 239–242.
14. Mahvi AH, Kia EB. Helminth eggs in raw and treated wastewater in the Islamic Republic of Iran. East. Mediterr. Health J. 2006; 12(1–2): 137–143.
15. Koné D, Cofie O, Zurbrügg C, Gallizzi K, Moser D, Drescher S, Strauss M. Helminth eggs inactivation efficiency by fecal sludge dewatering and co-composting in tropical climates. Water Res. 2007; 41(19): 4397–4402.
16. Abougrain AK, Nahaisi MH, Madi NS, Saied MM, Ghenghesh KS. Parasitological contamination in salad vegetables in Tripoli-Libya. Food Control. 2010; 21(5): 760–762.
17. Stoll NR. Investigations on the control of hookworm disease. XV. An effective method of counting hookworm eggs in feces. Amer J Hygiene. 1923; 3: 59–70.
18. Kochanowski M, Karamon J, Dąbrowska J, Cencek T. Coproscopical quantitative methods in the parasitological diagnosis – the use and problems with estimation of their efficiency (in Polish). Postep Mikrobiol. 2013; 52(1): 111–118.
19. World Health Organization. Basic Laboratory Methods in Medical Parasitology. Geneva, 1991.
20. Santos FLN, Cerqueira EJL, Soares NM. Comparison of the thick smear and Kato-Katz techniques for diagnosis of intestinal helminth infections. Rev Soc Bras Med Tro. 2005; 38(2): 196–198.
21. Goodman D, Haji HJ, Bickle QD, Stoltzfus RJ, Tielsch JM, Ramsan M, Savioli L, Albonico M. A comparison of methods for detecting the eggs of Ascaris, Trichuris, and hookworm in infant stool, and the epidemiology of infection in Zanzibari infants.Am J Trop Med Hyg. 2007; 76(4): 725–731.
22. Quinn R, Smith HV, Bruce RG, Gidwood RWA. Studies on the incidence of Toxocra and Toxocara spp. ova in the environment. I. A comparision of flotation procedures for recovering Toxocara spp. ova from soil. J Hyg. 1980; 84(1): 83–89.
23. PN-Z-19000-4 2001. Polish Standard. Soil quality – Assessment of the soil sanitary conditions – Detection of eggs of the intestinal parasites Ascaris lumbricoides and Trichuris trichiura.
24. Overgaauw PAM, van Knapen F. Toxocarosis, an important zoonosis. EJCAP. 2008; 18(3): 259–266.
25. Kochanowski M, Dąbrowska J, Karamon J, Cencek T, Osiński Z. Analysis of the accuracy and precision of the McMaster method in detection of the eggs of Toxocara and Trichuris species (Nematoda) in dog faeces. Folia Parasitol. 2013; 60(3): 264–272.
26. Zdybel J, Cencek T, Karamon J, Kłapeć T. Effectiveness of selected stages of wastewater treatment in elimination of eggs of intestinal parasites. Bull Vet Inst Pulawy. 2015; 59: 51–57.
27. Cox DD, Todd AC. Survey of gastrointestinal parasitism in Wisconsin dairy cattle. J Am Vet Med Assoc. 1962; 141: 706–709.
28. Varga I, Sreter T, Békési L. Quantitative method to assess Cryptosporydium oocyst shedding in the chicken model. Parasitol Res. 1995; 81(3): 262–264.
29. Malicki J, Montusiewicz A, Bieganowski A. Improvement of counting helminth eggs with internal standard. Water Res. 2001; 35(9): 2333–2335.
30. Gallizzi K. Co-composting reduces helminth eggs in fecal sludge, a field study in Kumasi, Ghana, SANDEC Switzerland http://www.eawag.ch/fileadmin/Domain1/Abteilungen/sandec/publikationen/SWM/Co-composting/Gallizzi_2003.pdf (access:2015.11.03).
31. Cobbina SJ, Kotochi MC, Korese JK, Akrong MO. Microbiological contamination in vegetables at the farm Gate Due to irrigation with wastewater in the Tamale Metropolis of Northern Ghana. JEP. 2013; 4(7): 676–682.
32. Marley CL, Cook R, Barrett J, Keatinge R, Lampkin NH, McBride SD. The effect of dietary forage on the development and survival of helminth parasites in ovine faeces, Vet Parasitol. 2003; 118(1–2): 93–107.
33. Pereckiene A, Petkevicius S, Vysniauskas A. Comparative evaluation of efficiency of traditional McMaster chamber and newly designed chamber for the enumeration of nematode eggs. Acta Vet Scand. 2010; 52(Suppl 1): S20.
34. Cringoli G, Rinaldi L, Maurelli MP, Utzinger J. FLOTAC: new multivalent techniques for quantitative copromicroscopic diagnosis of parasites in animals and humans. Nat Protoc. 2010; 5(3): 503–515.
35. Presland SL, Morgan ER, Coles GC. Counting nematode eggs in equine faecal samples. Vet Rec. 2005; 156(7): 208–210.
36. Plitt A, Imarom S, Joachim A, Daugschies A. Interactive classification of porcine Eimeria spp. by computer-assisted image analysis. Vet Parasitol. 1999; 86(2): 105–112.
37. Katz N, Chavez A, Pellegrino J. A simple device for quantitative stool thick–smear technique in schistosomiasis mansoni. Rev Inst Med Trop São Paulo. 1972; 14(6): 397–400.
38. Dąbrowska J, Zdybel J, Karamon J, Kochanowski M, Stojecki K, Cencek T, Kłapeć T. Assessment of the nematode eggs (genera: Ascaris, Toxocara, Trichuris) in sewage sludge with the use of LIVE/DEAD Bacterial Viability Kit. Ann Agr Env Med. 2014; 21(1): 35–41.
39. Karkashan A, Khallaf B, Morris J, Thurbon N, Rouch D, Smith SR, Deighton M. Comparison of methodologies for enumerating and detecting the viability of Ascaris eggs in sewage sludge by standard incubation-microscopy, the BacLight Live/Dead viability assay and other vital dyes. Water Res. 2015; 68: 533–544.
40. Dendukuri N, Joseph L. Bayesian approaches to modeling the conditional dependence between multiple diagnostic tests. Biometrics. 2001; 57(1): 158–167.
41. Nikolay B, Brooker SJ, Pullan RL. Sensitivity of diagnostic tests for human soil–transmitted helminth infections: a meta-analysis in the absence of a true gold standard. Int J Parasitol. 2014; 44(11): 765–774.
42. Kochanowski M, Karamon J, Dąbrowska J, Cencek T. Experimental estimation of the efficacy of the FLOTAC basic technique. J Parasitol. 2014; 100(5): 633–639.
43. Karamon J, Sroka J, Cencek T. Limit of detection of sedimentation and counting technique (SCT) for Echinococcus multilocularis diagnosis, estimated under experimental conditions. Exp Parasitol. 2010; 124(2): 244–246.
Copy url