Livestock Research for Rural Development 31 (2) 2019 Guide for preparation of papers LRRD Newsletter

Citation of this paper

Parasites infesting Nile tilapia grown in aquaculture systems in Kenya

D M Mukwabi, S O Otieno1, P O Okemo2, R O Odour2 and B Agwanda3

Kenya Fisheries Service, PO Box 48511-00100, Nairobi, Kenya
dmmakilla@yahoo.com
1 Kenyatta University, Department of Zoological Sciences, PO Box 43844-00100, Nairobi, Kenya
2 Kenyatta University, Department of Biochemistry, Microbiology and Biotechnology, PO Box 43844-00100, Nairobi, Kenya
3 National Museums of Kenya, PO Box 40658, Nairobi, Kenya

Abstract

Aquaculture production faces a number of challenges, including fish diseases and pollution. However, most studies on fish diseases in Kenya have been done on fish in the wild. This study, therefore, focused on characterization of parasites present in Nile tilapia farmed in aquaculture systems in Bungoma County. Fish were harvested from ponds using a seine net. Fins, scales and skin were sampled from each fish and placed in sterile Bijou bottles.The harvested fish were opened dorso-ventrally under aseptic conditions and mouth, gills, stomach, liver, kidney and intestinal contents washed into the sterile Bijou bottles. Physiological saline was then added and stirred using a mounted pin. One hundred (100) ml of fish pond water samples were also collected in sterile bijou bottles and 3ml of 0.9% physiological saline added. The bijou bottles containing sampled water were then packed in a cooler box. Commercial fish feeds were purchased from fish feed millers operating in the County. The sampled fish, pond water and fish feeds were transferred to National Museums of Kenya Parasitology laboratory in Nairobi where they were subjected to parasitilogical analysis using hand lens and microscopy. At every fish pond, five water quality parameters were assessed using a portable auto sampler. The parasites recovered werePhilometroides spp., Acanthocephalus spp. and Procamallanus spp. from Nile tilapia and Cleidodiscus spp. from pond water. It is recommended that aquaculture systems management in the county should put in place measures to increase turbidity levels to minimize stress on fish thereby reducing infection levels from parasites.

Key words: diseases, farms, sub county, water quality


Introduction

In Kenya, fish farming is practiced in regions endowed with a lot of water resources including springs, wetlands, rivers, streams and reservoirs, in earthen ponds, cages, raceways, tanks, lockable ponds and recirculating systems (Nyonje et al 2011; Munguti et al 2014;). The main fish species cultured are Nile tilapia (Oreochromis niloticus), African catfish (Clarias gariepinus), Common carp (Cyprinus carpio) and Rainbow trout (Oncorhynchus mykiss) (Ngugi et al 2007; MFD 2008; 2012). The capita fish consumption in Kenya and Africa is 4.5 kg and 10 kg respectively (APRM 2015). At the global level, the per capita fish consumption is 20 kg per person due to investments and production in aquaculture. China, India, Viet Nam, Bangladesh, Egypt, Greece, Czech Republic, Hungary, Laos and Nepal produce more farmed than wild-caught fish (FAO 2016).

Fish farming faces challenges of diseases (Munguti et al 2014) which constitute the largest cause of economic loss with a global estimate at approximately US$ 3 billion per annum (Subasinghe et al 2001). Among the fish disease- causing parasites namely platyhelminthes, helminthes, protozoa, acanthocephalans, arthropods, nematodes, gastropods and trematodes (Al-Harbi 1994; Al-Harbi and Uddin 2003; Faruk et al 2004; Mastan et al 2009; Orina et al 2014; Emere and Dibal 2014;). The cultured fish are usually infected by parasites present either in fish pond water, introduced via source of pond water supply, fish feeds or surface run-offs. Fish faming is becoming an important undertaking in many of the African resource poor settings. Yet there are few articles targeting fish health and production. In Kenya, endoparasites such asPolyacanthorhynchus, Amirthalingamia, Cyclustera and Proteocephalus occur in wild fish in Lake Naivasha and Oloiden Bay while Contracaecum and Procamallanus occur in River Tana (Aloo 2002; Gichohi 2010). The prasitesProcamallanus, Contacaecum, Camallanus, Acanthocephala and Trypanosoma spp. are prevalent in fish from Lake Victoria (Kamundia 2011).

Environmental factors such as temperature, oygen content and pH influence parasitic fish infections (Ali et al 2004; State and State 2009; Lagrue et al 2011). An increase in stream temperature by 5C enhances growth rate and adhesion capacity of parasites to fish tissues. Decrease in dissolved oxygen due to temperature rise above 25C jeopardises the immune system of fish, increasing the potential of parasitic infections. Most warm water fish prefer dissolved oxygen in excess of 2mg/l (Suomalainen et al 2005). Suspended solids affect fish by physically abrading the skin, clogging gills, and impairing visual feeding (Wedemeyer 1996). An increase in ventilation rate due to reduced water quality will increase the volume of water (and number of water-borne pathogens) passing through the opercular cavity and damage the gills epithelium that increase the risk of uptake of pathogens (Graig 2008).


Materials and methods

The study was carried out in Bungoma County where a total of 78 farms spread in the six sub counties were sampled to determine the pathogenic parasites infecting Nile tilapia which is the major fish species grown.

Sampling of fish

The sampling procedures were carried out according to techniques described bu Emere and Egbe (2006). The fish were harvested using a seine net. Fish skin and fins were sampled from each fish and placed in sterile Bijou bottles. Fish were opened dorso-ventrally in aseptic conditions and gills, mouth, stomach and intestinal contents were washed into the sterile Bijou bottles containing 3ml of 0.9% physiological saline and stirred using a mounted pin. The Bijou bottles were then packed in a cooler box with ice and transferred to National Museums of Kenya Parasitology laboratory in Nairobi. The study involved nine (9) sampling sessions, each session taking place after every three months that ran from August 2015 to December 2017 with 394 pieces of different sizes of fish sampled. In the laboratory, external parasites (from skin and fins) were examined using hand lens. To examine internal parasites, 0.5 ml of the sampled internal organs solutions were collected using a dropper, placed on a slide, and then covered with a cover slip after which, observations were made using a compound light microscope at 100X - 400X magnification. Parasites present were identified using the fact sheets on pictorial guide on fish parasites and counted (Pouder et al 2005).

Sampling of pond water

For each fish pond that was sampled, 100 ml of water was collected in sterile bijou bottles, after which 3ml of 0.9% physiological saline solution was added. The Bijou bottles containing sampled water were then packed in a cooler box with ice and transferred to National Museums of Kenya Parasitology laboratory in Nairobi. In the laboratory, parasites were examined by placing 0.5 ml of the mixed solution using a dropper on a slide. The drop was covered with a cover slip after which, observations were made using a compound light microscope at 100X - 400X magnification. Parasites present were identified and counted.

Sampling of fish feeds

Commercial fish feeds that farmers use to feed their cultured fish were purchased from two fish feed millers in Bungoma County. The purchased fish feeds were transported to National Museums of Kenya Parasitology laboratory in Nairobi for parasitological analysis. In the laboratory, 1 g of fish feeds were washed into the sterile bijou bottles with 3ml of 0.9% physiological saline and stirred using a mounted pin. Then 1ml of the mixed solution was collected using a dropper, placed on a slide, covered with a cover slip, and observations made using a compound light microscope at 100X - 400X magnification. Parasites present were identified and counted.

Sampling of water quality parameters

Water quality parameters were assessed using a portable auto sampler in every pond from which fish had been sampled. The parameters assessed were temperature, dissolved oxygen, conductivity, pH and turbidity.

Data management and analysis

Kruskal-Wallis test was used to determine differences in parasite infestation rates. Regression analysis was used to determine the effect of temperature, dissolved oxygen, electrical conductivity, pH and turbidity on the number of isolated parasites.


Results

During the study, some of the sampled fish showed symptoms of infection. The observed symptoms were dark eyes, deformed scales, macerated fins and stuffy growth (Photo 1).

Photo 1. Nile tilpia sampled from Bungoma West Sub County
with dark eyes, macerated fins and stuffy growth

Parasites were recovered from Nile tilapia and pond water. Nile tilapia sampled from Bumula, Mt. Elgon and Bungoma East Sub Counties recorded no parasites in any of the organs that were observed. However, pathogenic parasites of different species were recorded (Table 1) from Bungoma South, Bungoma West and Bungoma North sub counties. Further, only pond water sampled from Bungoma North Sub County had a positive result for parasites whereas the other five sub counties had zero parasites in water collected for analysis. Further, there were no parasites recovered in fish feeds sampled.

Table 1. Results for parasites present in Nile tilapia

Sub County

Farms

Organ

Parasitic Species

Mean

Bungoma South

5

Intestine

Acanthocephalus spp.

2.5

Bungoma West

5

Gills

Philometroides spp.

2.5

3

Stomach

Philometroides spp.

1.33

Bungoma North

3

Stomach

Procamallanus spp.

3.33

In Bungoma South, parasites isolated from intestines ranged from 0 to 3 ( =2.5). The species of parasites recovered were Acanthocephalus spp. (Photo 2) from 5 (28%) of (n=18) of the farms sampled in the Sub County. There was no difference (p=0.35) in the distribution of the total parasites in the intestines among the sub counties.

Photo 2. Acanthocephalus spp. isolated from intestines in Bungoma South Sub County

In Bungoma West, there were parasites recovered from gills and stomach. Those recovered from gills ranged from 0 to 4 ( =2.5) while those from the stomach ranged from 0 to 2 ( =1.33). The recovered parasites from gills were Philometroides spp. (Photo 3) in 5 (20%) of the farms (n-25) sampled with no significance difference (p= 0.484) in the distribution of the parasite in gills. In the stomach (Photo 4), Philometroides spp. were detected from 3 (12%) of the farms (n=25) sampled in the Sub County. There was no difference (p=0.47) in the distribution of parasites across the sub counties.

Photo 3. Philometroides spp isolated from gills in Bungoma West Sub County Photo 4. Philometroides spp isolated from stomach in Bungoma West Sub County

Finally in Bungoma North Sub County, the highest number of parasites were 5 ( =3.33) from 3 (25%) of the farms (n=12); all were recovered from the stomach. The recovered species were Procamallanus spp. (Plate 5). There was no difference (p= 0.47) in the distributions of the parasites.

Photo 5. Procamallanus spp isolated from the stomach in Bungoma North Sub County
Parasites in pond water

Parasites isolated from pond water had the lowest total number at 0 counts and the highest was 9 counts ( =8.74), recovered from 33.3% of farms sampled. The parasites recovered were flatworms (Photo 6) of the genus Cleidodiscus spp.

Photo 6. Cleidodiscus spp isolated from pond water in Bungoma North Sub County
Water quality parameters

There were no difference in water quality parameters of fish ponds (Table 2. ).

Table 2. Mean values for water quality parameters of fish ponds

Farms

Temperature
(C)

Dissolved Oxygen
(mg/l)

Conductivity
(s/ cm)

pH

Turbidity
 (ppm)

Bumula

10

30.4

2.8

88.2

6.72

15

Bungoma South

18

29.1

2.5

103

6.73

18.6

Bungoma West

25

26.7

3.04

83.6

6.63

15.4

Bungoma North

12

21.7

2.71

97.2

6.83

14.9

Mt. Elgon

5

23.6

2.85

91

6.71

17.4

Bungoma East

8

29.9

2.94

87.4

6.76

14.8


Discussion

The recovery of Philometroides spp., Procamallanus spp. and Acanthocephalus spp. from the gills, intestines and stomachs indicates that the three species (nematodes) prefer to inhabit digestive systems. This could be attributed to the nematodes’ parasitic nature where they rely on the hosts for their food requirements (Yanong 2006). The occurrence of Cleidodiscus species in farmed fish could be related to contamination of the fish farms and hatcheries via runoffs.

The presence of Acanthocephalus spp. could be attributed to the fact that the parasite has a wide geographical range (Gichohi 2010; Kamundia 2011; Maina et al 2017; Mavuti et al 2017). The observed Acanthocephalus spp. in the intestines mirrors what has been documented in other regions within the country. The Acanthocephalus spp. was found in the intestines of farmed Nile tilapia in Tetu Sub County in Nyeri (Mavuti et al 2017). In addition, Acanthocephalus spp. was observed in farmed Nile tilapia Kikuyu Sub County in Kiambu County (Maina et al 2017). Furthermore, infections of Oreochromis niloticus sampled from Lake Victoria in Homa Bay County (Kamundia 2011) and River Tana (Gichohi 2010) withAcanthocephalus spp. have been recorded. Regionally, Acanthosentis tilapiae was reported in Bahr Youssef and Fayoum fish farms in Egypt (Dayhoum 2003).

The presence of nematodes Philometroides spp. and Procamallanus spp. in farmed tilapia implies that intermediate hosts for the nematodes exist in the farms in which fish were harvested. This is because nematodes require at least one intermediate host before it infects fish (Paperna 1980). Alternatively, it could be that the fish that were stocked already had larval stages of the nematodes and therefore, the source of contamination was the fish hatcheries and farms where fish had been sourced. The occurrence of nematodes has been documented in Sangoro and Sagana fish farms that are located in Lake Victoria basin and Mt Kenya highland regions, respectively (BOMOSA 2009). Philometroides spp. is known to cause hemorrhage, fibrosis and chronic active ulcerative dermatitis in farmed fish. Therefore, their presence in Bungoma West Sub County requires improved farm husbandry measures to avoid its spread to other sub counties.

The findings of the current study concur with that of Adikwu and Ibrahim (2004) and Ayanda (2009) who reported the presence of Procamallanus spp. in catfish from Kano region and in farmed catfish as well as in the intestine of freshwater Synodontis zambezensis in lower Congo River, Democratic Republic of Congo (Boomker 1993). Fish is infected by Procamallanus spp. while feeding on the copepod harbouring the infective-stage larvae (De et al 2000).

Cleidodiscus spp. recovered from water sampled from Bungoma North Sub County could have originated from contaminated run offs where the parasites found their way into rivers passing through the sub county. The rivers in the County are the main source of water supply for fish ponds. The presence of Cleidodiscus spp. in pond water is in concurrence with other reports showing that it is a fresh water parasite in tropical regions (Noga 2000). Though Cleidodiscus spp. is endoparasitic, they also infect gills and nasal cavities (Hoffman 1998).

The water parameters tested during the period of this study other than turbidity were found suitable for Nile tilapia survival without predisposing it to opportunistic pathogens recovered. This is in line with the Kenyan national standards for physical and chemical parameters for production of fresh water fish culture (MALF 2015). The required standards include; temperature, 22-32C for warm water and 14-17C for cold water fish; dissolved oxygen, ≥3 mg/l in warm water and ≥5 mg/l in cold water. The conductivity standards are 2-10 s/ cm for warm water as well as cold water; pH of 6.5-8.5 for both cold and warm water fish cultures. Finally, turbidity standards are 30-45ppm.

The current study recorded temperature ranges of 21.7-30.4C that are within the recommended Kenyan national standards (14-32C) for Nile tilapia production in both cold and warm water areas (MALF 2015). Further, the ranges fall within the acceptable temperature ranges of 15C-35C for tilapia fish production (Anita and Pooja 2013). High temperatures are known to accelerate life cycles of parasites in infected fish (Hassan 1999).

The mean ( =2.8mg/l) of dissolved oxygen recorded in Bumula could be linked to levels of organic manure used to fertilize fish ponds that takes up a lot of dissolved oxygen for decomposition. The low levels of dissolved oxygen in Bungoma North ( =2.71mg/l) and Bungoma South ( =2.52mg/l) could be attributed to inorganic fertilizer being used in the two Sub Counties for crop production that are washed into ponds and that used in fertilizing fish ponds. The study recorded dissolved oxygen levels that fall within recommended levels of 1 mg/ l to 8 mg/ l for fish ponds (Anita and Pooja 2013; Ekubo and Abowei 2011).

The high conductivity recorded in Bungoma South ( =103s/cm) could be attributed to the economic activities of sugarcane and maize production in the sub county. The inorganic fertilizer used to grow sugarcane and maize could release ions that contributed to conductitvity levels going up. All the conductivity levels among the six sub counties were found to be within recommended ranges for fish production. This implies that there was no stress exerted on farmed fish arising from conductitvity.

The highest pH (6.83) in Bungoma North could be due to its rocky terrain and soil texture. In Niger delta in Nigeria, pH recorded ranged from 6.5 to 9.5 for all ponds sampled (Njoku et al 2015). The recorded mean ranges (6.63 - 6.83) in this study are within allowable ranges for tilapia culture production. The appropriate pH for fish production is 6-9 (Ntengwe and Edema 2008; Mohamed 2005).

Among the six sub counties, Bungoma South had the highest ( = 18.6ppm) turbidity levels which could be due to high levels of sediment depositions in fish ponds from organic manure and inorganic fertilizers which are used to fertilize fish ponds. Mt. Elgon Sub County was found to have the second highest ( = 17.4ppm) turbidity probably because of runoffs that depost sediments into fish ponds especially during rainy season. These ranges are lower than recommended ranges (15-80 ppm) for fish health (Santhosh and Singh 2007; Bhatnagar et al 2004).


Conclusions


Acknowledgement

We acknowledge the National Commission for Science, Technology and Innovation-Kenya for the Post-Graduate Students Funding in the financial year 2015/2016 grant for the financing of the study.


References

Adikwu I A and Ibrahim E A 2004 Studies on the endoparasites in the gastro - intestinal tract of Clarias gariepinus (Tugels) in Wase dam, Kano State, Nigeria. African Journal of Applied Zoology and Enviromental Biology, 6:35-40. http://ajoI.info/index.php/ajazeb/ article/view/41173 on 31/12/2009.

African Peer Review Mechanism-Kenya (APRM) 2015 The Second Kenya Self-Assessment Report. February; pp 872

Al-Harbi A H 1994 First isolation of Streptococcus spp. from hybrid tilapia (Oreochromis niloticus and O. aureus) in Saudi Arabia. Aquaculture, 128: 195-201.

Al-Harbi A H and Uddin M N 2003 Quantitative and qualitative studies on bacterial flora of hybrid tilapia ( Oreochromis niloticus O. aureus) cultured in earthen ponds in Saudi Arabia. Aquaculture Research, 34 (1):43-48.

Ali F M, Abdus–Salam B A, Ahmad K S, Qamar M and Umer K 2004 Seasonal variations of physico – chemical characteristics of River Soan water at Dhoak Pathan Bridge (Chakwal), Pakistan. International Journal of Agriculture and Biology, 6(1): 89 – 92. https://www.researchgate.net/publication/237694196.

Aloo P A 2002 A comparative study of helminth parasites from the fish Tilapia zillii and Oreochromis leucostictus in Lake Naivasha and Oloidien Bay, Kenya. Journal of Helminthology, 76: 95-102.

Anita B and Pooja D 2013 Water quality guidelines for the management of pond fish culture. International Journal of Envinmental Science, 3(6): 1980-2009.

Ayanda O I 2009 Comparative parasitic helminth infection between cultured and wild species of Clarias gariepinus in Ilorin, North - Central Nigeria. Scientific Research and Essay, 4:018-021. http: /Avww.academicjournal.org/SRE

Bhatnagar A, Jana S N, Garg S K, Patra B C, Singh G and Barman U K 2004 Water quality management in aquaculture, In: Course Manual of summer school on development of sustainable aquaculture technology in fresh and saline waters, CCS Haryana Agricultural, Hisar (India):203- 210.

BOMOSA 2009 Integrating BOMOSA cage fish farming system in reservoirs, ponds and temporary water bodies in Eastern Africa: Final activity report of veterinary and public health aspects (results of surveys and recommendations). https://cordis.europa.eu/docs/publications/1228/122807391-6-en.pdf.

Boomker J 1993 Parasites of South African freshwater fish. IV. Description ofSpirocamallanus daleneae n. sp. (Nematoda: camallanidae) fromSynodontis zambezensis Peters 1852 (Mochokidae) with comments on Spirocamallanus spiralis (Baylis, 1923). Onderstepoort Journal of Veterinary Research, 60: 131-137.

Dayhoum A H M A 2003 A general survey of the helminth parasites of fish from inland waters in the Fayoum Govemorate, Egypt. Parasitological Research, http://www.springer.com/content on 9/7/2009 .

De Buron I and Nickol B B 1994 Histopathological effects of the acanthocephalanLeptorhynchoides thecatus in the ceca of the green sunfish, Lepomis cyanellus. Transactions of the American Microscopical Society, 133:161-168.

Ekubo A A and Abowei J F N 2011 Review of some water quality management principles in culture fisheries. Res. J. Appl. Sci., Engineering and Technology, 3(2): 1342-1357.

Emere M C and Dibal D M 2014 Diseases of a (Clarias gariepinus) Freshwater Fish from River Kaduna, Nigeria. World Rural Observation, 6(2):77-81.

Emere M C and Egbe E L N 2006 Protozoan parasites of Synodonits clarias (a fresh water fish in river Kaduna). BEST Journal. 3(3):58–64.

FAO 2016 The State of World Fisheries and Aquaculture. Contributing to food security and nutrition for all. Rome, Pp 200.

Faruk M A R, Sarker M M R, Alam M J and Kabir M B 2004 Economic loss from Fish Diseases on Rural Fresh water aquaculture of Bangladesh. Pakistan Journal of Biological Sciences, 7(12): 2086-2091

Gichohi C M 2010 Prevalence, Intensity and Pathological lesions associated with Helminth infections in farmed and wild fish in Upper Tana River basin, Kenya. MSc.Thesis, University of Nairobi, Kenya

Graig M M 2008 Water quality and welfare assessment on United Kingdom trout farms, Ph.D Thesis. Institute of Aquaculture, University of Stirling, Pp 203

Hassan A H M 1999 Trichodiniasis in Farmed Freshwater Tilapia in Eastern Saudi Arabia. Journal of KAU: Marine Science, 10:157-168. https://www.kau.edu.sa/Files/320/Researches/51774_21907.pdf

Hoffman G L 1998 Parasites of North American Freshwater Fishes (2nd Ed.). Ithaca, New York: Cornell University Press, Pp 576.

Kamundia P W 2011 Pathological changes associated with pollution and endoparasites in Nile perch and tilapia in Lake Victoria, Kenya. MSc Thesis, University of Nairobi, Kenya.

Lagrue C, Kelly D W, Hicks A and Poulin R 2011 Factors influencing infection patterns of trophically transmitted parasites among a fish community: host Diet, host –parasite compatibility or both? Journal of Fish Biology, 79:466-85.

Maina K W, Mbuthia P G, Waruiru R M, Nzalawahe J, Murugami J W, Njagi L W, Mdegela R H and Mavuti S K 2017 Risk factors associated with parasites of farmed fish in Kiambu County, Kenya. International Journal of Fisheries and Aquatic Studies, 5(4): 217-223.

Mastan S, Priya G L and Babu E G 2009 Hematological profile of Clarias batrachus L. exposed to subhethal doses of lead nitrate of lead nitrate. The Internet Journal of Hematology, 6 (1): 1-7. http:// ispub.com/IJHE/6/1/10239#

Mavuti S K, Waruiru R M, Mbuthia P G, Maina J G, Mbaria J M and Otieno R O 2017 Prevalence of ecto- and endoparasitic infections of farmed tilapia and catfish in Nyeri County, Kenya. Livestock Research for Rural Development, 29, Article #122. http:// www.lrrd.org/ lrrd29/6/ stek29122.htm l

Ministry of Agriculture, Livestock and Fisheries (MALF) 2015 State Department of Fisheries: Manual of Standard Operating Procedures for Fish Inspection and Quality Assurance in Kenya. Government Press, Pp 260.

Ministry of Fisheries Development (MFD) 2008 Strategic plan 2008-2012.

Ministry of Fisheries Development (MFD) 2012 Annual Statistical Bulletin.

Mohammed H A 2005 Physico -chemical characteristic of Abuzabaal pond, Egypt. Egypt Journal of Aquaculture Resources, 316:1-13.

Munguti J M, Kim J D and Ogello E O 2014 An Overview of Kenyan Aquaculture: Current Status, Challenges, and Opportunities for Future Development. Fish and Aquatic Sciences, 17: 1-11.

Ngugi C N, Bowman J R and Omolo B O 2007 A new guide to fish farming in Kenya. Aquaculture CRSP Management Office, Oregon USA, Pp 95.

Njoku O E, Agwa O K and Ibiene A A 2015 An investigation of the microbiological and physicochemical profile of some fish pond water within the Niger Delta region of Nigeria. European Journal of Food Science and Techlogy, 3 (4): 20-31.

Noga E J 2000 Fish disease: diagnosis and treatment. Iowa: Iowa state university press, USA. Pp 367. https://trove.nla.gov.au/version /46521894

Ntengwe F N and Edema M O 2008 Physico-chemical and microbiological characteristics of water for fish production using small ponds. Journal of Physics and Chemistry of Earth, 33:701-707.

Nyonje B M, Charo-Karisa H, Macharia S K and Mbugua M 2001 Aquaculture Development in Kenya: Status, Potential and Challenges. In Samaki News: Aquaculture Development in Kenya in Kenya towards Food Security, Poverty Alleviation and Wealth Creation, 7 (1): 8-11.

Orina P S, Maina J G, Wangia S M, Karuri E G, Mbuthia P G and Omolo B 2014 Situational analysis of Nile tilapia and African catfish hatcheries management: a case study of Kisii and Kirinyaga counties in Kenya. Livestock Research for Rural Development, 26 (Article #87). http://www.lrrd.org/lrrd26/5/orin26087.htm

Paperna I 1980 Parasites, infections and Diseases of Fish in Africa, CIFA Technical Paper, 7: Pp 216.

Pouder D B, Yanong R P and Curtis E W 2005 Common Freshwater Fish Parasites Pictorial Guide-EDIS (UF/IFAS) Fact sheets. http://edis.ifas.ufl.edu/fa11400.pdf and 11500.pdf

Santhosh B and Singh N P 2007 Guidelines for water quality management for fish culture in Tripura, ICAR Research Complex for NEH Region, Tripura Center, Publication No. 29.

State E and State A I 2009 Parasite Assemblages in Fish Hosts. Bio-Research, 7(2):561-70.

Subasinghe R P, Bondad-Reontase M G and McGladdery S E 2001 Aquaculture development, health and wealth. In: Subasinghe R. P.,P. Bueno, M J Philips, C Hogh, S E McGladdery and J R Arthur (Eds). Aquaculture in the Third Millennium Technical Proceedings of the Conference on Aquaculture in the Third Millennium, Bangkok, Thailand, 20-25 February 2000. NACA, Bangkok and FAO, Rome, Pp 167-191.

Suomalainen L R, Tiirola M and Valtonen E T 2005 Influence of rearing conditions on Flavobacterium columnare infection of rainbow trout, Oncorhynchus mykiss (Walbaum). Journal of Fish Diseases, 28(5): 271-277.

Wedemeyer G A 1996 Physiology of fish in intensive culture systems. Chapman & Hall, London, UK, Pp 201.

Yanong E P R 2006 Nematode (Roundworm) infections in fish. Circular 91, Department of Fisheries and Aquatic Sciences, Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida.


Received 27 December 2018; Accepted 6 January 2019; Published 1 February 2019

Go to top