Print version ISSN 1019-9128
J. S. Afr. Vet. Assoc. vol.82 no.3 Pretoria Sept. 2011
A J LeslieI,*; C J LovelyI; J M PittmanII
IDepartment of Conservation Ecology & Entomology, University of Stellenbosch, Private Bag X1, Matieland, 7600 South Africa
IIJohannesburg Zoo, Private Bag X13, Parkview, 2122 South Africa
The objective of this study was to conduct a preliminary survey of diseases that might be present in the wild Nile crocodile population in the Okavango Delta, Botswana. Blood samples were collected from crocodiles ranging in size from 34.0 cm to 463.0 cm total length. Samples were examined for blood parasites and underwent a haematological analysis. Before release the crocodiles were examined for various clinical abnormalities. Of the 144 crocodiles examined, none were visibly sick or displayed any signs of disease. No antibodies to Mycoplasma crocodyli were detected. Hepatozoon pettiti was present in 55.3 % of blood smears examined, but there was no significant difference in any of the haematological values between the infected and uninfected crocodiles, and a high prevalence of Hepatozoon infection is not uncommon in other species. Only 7.6 % of the examined crocodiles were infested with leeches. Further research is required for several of the crocodilian diseases, in particular to elucidate the role of wild crocodilians as reservoirs of infection.
Keywords: Nile crocodile, Crocodylus niloticus, Okavango Delta, disease survey.
Since the development of the crocodile farming industry in the 1980s, a number of studies have been carried out worldwide on diseases in farmed crocodiles6,7,12,13,20,27,28. Very little, however, is known about the diseases of wild crocodilians, particularly wild Nile crocodiles. Wild crocodiles are often difficult to study owing to the remoteness of the areas in which they occur.
The initial objective of this study was to determine disease prevalence in the wild Nile crocodile population in the Okavango Delta. However, several factors made this difficult:
(i) Very few serological tests have been developed for crocodile diseases. Consequently a serological survey is of very limited value.
(ii) Virus isolation is unavailable due to the lack of crocodile cell lines in veterinary diagnostic laboratories.
(iii) Sick crocodiles are difficult to identify by observation. Under farmed conditions they often stop eating but show no other clinical signs and therefore are not noticed to be unhealthy until they are found dead. Diseases causing visible external lesions, for example pox virus infection, are the obvious exception.
(iv) Clinical examination has several limitations: cardiac and respiratory rates are variable according to conditions, and are strongly influenced by capture stress. Because crocodiles are poikilothermic, body temperature cannot be used as a diagnostic tool.
(v) Sick/weak crocodiles are unlikely to survive for long due to predation. Therefore, the likelihood of encountering these individuals in a capture and release survey is low.
(vi) Owing to research permit limitations no crocodiles could be sacrificed for histopathological and parasitological examination.
In view of these limitations, it was postulated that a low incidence of diseases would be found in the Okavango population during a capture and release survey. However, it was possible to do a more accurate survey for mycoplasmosis and haemogregarine infection thanks to the availability of diagnostic tests. In addition, ectoparasites, in the form of leeches, were easy to find, remove and count per individual.
MATERIALS AND METHODS
Botswana's Okavango Delta, the world's biggest Ramsar site (a wetland of international importance), is a large wetland within the Kalahari Desert, covering an area of approximately 16 000 km2 in the dry season and increasing to over 22 000 km2 with the annual flood. The 111 250 km2 active catchment area falls entirely within Angola. Owing to the geology of the catchment, the incoming water is low in nutrients and sediment32.
The Okavango River flows through Namibia briefly before entering Botswana and forming a broad floodplain, the Panhandle. An estimated 40 % of incoming water leaks into the surrounding swamps by the time the river leaves the Panhandle. The remaining 60 % is distributed down 3 main channels, which fan out to form the Delta. The Okavango Delta consists of permanent and seasonal swamp, which is inundated during the annual flood32.
The northern part of the Delta is characterised by shallow water, flooded grasslands, ox-bow lakes and lagoons mostly interconnected by narrow waterways. Only a few main channels lined by tall reeds (mainly Phragmites australis) carry the remainder of the Okavango's water southwards through the Delta. The permanent and seasonal swamp together form a unique ecosystem and provide a high-quality habitat for a great many species. As a keystone species, the Nile crocodile, helps maintain the fragile balance within this ecosystem. This includes selective predation on various fish species,10,42 recycling of nutrients and maintenance of wet refugia in times of drought48. Crocodiles are unevenly distributed throughout the Delta with the majority of the breeding population occurring in the 120 km long Panhandle17.
The Nile crocodile, Crocodylus niloticus, is the most widespread and abundant of the 3 crocodile species that occur in Africa. It occurs throughout the continent south of the Sahara in a variety of wetland habitats, including coastal areas41,46. Historically its distribution in southern Africa extended as far south as the Eastern Cape Province during the past 100-200 years42.
Nile crocodiles are ectothermic and regulate their body temperature behaviourally by moving between sun-exposed sandbanks and the water. Typical adult lengths are around 3.5 m, but the males can grow up to 5 m18. Sexual maturity is reached from 2.9 m total length for males, and 2.2 m for females10. Nesting occurs in a hole in the ground and on average 50 eggs are laid. Nile crocodiles exhibit temperature-dependent sex determination31. Hatchlings emerge after an incubation period of approximately 90 days in early to mid summer, and parental protection occurs39,40,41. As with other crocodilian species, a high mortality rate is experienced in their 1st year of life due primarily to predation45.
One hundred and forty-four crocodiles were captured in the Panhandle of the Okavango during summer (February 2005). Capture was carried out using 2 methods. At night, using a 4.8 m flat-bottomed aluminium motor boat, crocodiles were located with a spotlight which revealed the eyes glowing red. The beam of light was then kept focused on the crocodile's eyes, making it possible to approach the animal by boat. Crocodiles estimated to be smaller than 1.2 m total length were captured by hand. Crocodiles between 1.2 m and 2.3 m were captured using a swivelling noose (Animal Handling Co., SA) which was placed over the snout and pulled tight in the neck region. Crocodiles were then brought onto the boat, jaws were taped shut and the animals were physically restrained. Animals larger than 2.3 m were captured using a noose attached to a climbing rope, which was secured to the boat. The crocodile was allowed to swim so as to tire it out before it was brought onto the boat. In addition to night capture, baited box and Pitman traps were strategically placed on river banks. Traps were baited at sunset and checked at 1st light the next morning. Captured animals were immediately restrained, measured, examined and samples collected.
Each crocodile was blindfolded and restrained in ventral recumbency. Fifty-three animals were randomly selected for blood collection. Blood was collected from the post-occipital sinus9 on the dorsal midline and just caudal to the base of the head. A 21 G (0.80 × 25-90 mm) or 23 G (0.65×25mm) needle and a 3, 5 or 10 m syringe was used, depending on the size of the crocodile, and the blood was transferred directly into a lithium heparin tube. Blood smears were made from whole blood using the cover slip method24. Following blood collection, each crocodile was measured (total length (TL) and snout-to-vent length (SVL) using a flexible measuring tape, ±1 mm). It was then weighed using a harness placed around the forelimbs and a Pesola spring balance. Each crocodile was sexed by cloacal examination of the cliteropenis23,31 and the entire body was examined for clinical abnormalities including bite wounds, skin lesions, conjunctivitis, joint swelling and poor condition. Dorsal and lateral body surfaces were also examined for the presence of leeches. Leeches were removed by means of a pair of tweezers, counted and stored in 70 % ethanol for later identification.
On return to the field laboratory 1.0 m of blood was transferred to an Eppendorf tube for haematological analysis. The remaining blood was centrifuged using a manual desktop centrifuge and the plasma frozen for serology. If the volume of the blood sample was small it was allocated for either Hepatozoon examination or serology.
Thirty-eight samples were examined for blood parasites and underwent haematological analysis. Blood smears were stained with Diff-Quick Stain (American Scientific Products, Illinois, USA)8. The presence of Hepatozoon gametocytes was determined by microscopic examination of the Diff-Quick stained blood smears. A minimum of 3 slides per animal and in some cases more were examined including a minimum of 1000 HPF's (×100 oil immersion) per slide. The degree of erythrocyte regeneration was estimated by examining the red cell series and scoring each slide on a scale of 1 to 4.
On this scale, a score of 2 represented normal erythrocyte regeneration with 10-20 % orthochromic erythroblasts but no earlier stages. A score of 2.5 represented a moderate increase in erythrocyte regeneration, with late polychromatic erythroblasts through to mature and aging erythrocytes, while a score of 3 represented strong regeneration with basophilic erythroblasts and early poly-chromatic erythroblasts through to mature and aging erythrocytes, but no proerythroblasts. Finally, a score of 4 included all the other stages plus proerythroblasts.
Packed cell volumes (PCV) were determined using a Statspin MP microhaematocrit centrifuge: blood was drawn into standard microhaematocrit tubes and centrifuged for 5 min at 12 000 g.
Total red cell counts (RCC) were performed both manually and automatically using an electronic particle counter. The automated counts were made using a Beckman Coulter Ac*T Series haematology analyser (Coulter SA). The manual counts were made using Natt and Herrick's solution. A 1:200 dilution was made by drawing blood up to the 0.5 mark on a red blood cell diluting pipette, then filling the pipette to the 101 mark with Natt and Herrick's solution9. The diluted blood was then used to charge both counting chambers of an improved Neubauer haemocytometer (Hawksley and Sons, Lancing, UK). After 5 mins in a humidity chamber the red cells were counted in the 4 corner cells and central cell of the central large square of the counting chamber. This was repeated on the second chamber and the average multiplied by 10 000 to obtain the total red cell count per microlitre.
Haemoglobin concentrations (Hb) were determined using a Beckman Coulter Ac*T Series haematology analyser (Coulter SA).
Red blood cell indices were calculated using standard equations24
Mean cell volume:
MCV (fl) = PCV/RCC
Mean cell haemoglobin:
MCH (pg) = Hb (g/d ) × 10 / RCC
Mean cell haemoglobin concentration:
MCHC (g/d ) = Hb (g/d ) / PCV
Total white cell counts (WBC) were obtained indirectly using the Unopette 5877 system (Becton-Dickinson, USA). The Unopette pipette was filled with blood (25 µ ) and mixed with the phloxine B diluent in the reservoir. From this, both counting chambers of an improved Neubauer haemocytometer were charged. After 5 min in a humidity chamber all the pink-staining granulocytes were counted in both chambers.
Differential counts were done on the Diff-Quick stained smears. The percentage of heterophils and eosinophils was calculated and used to calculate total WBC9.
The separated plasma was stored at -10 ºC in a domestic gas freezer for up to 1 month. After return from the research site, 30 samples were submitted to the laboratory (Faculty of Veterinary Science, Department of Veterinary Tropical Diseases, University of Pretoria) for serological testing for mycoplasmosis, using a plate agglutination test.
Haematological values were analysed for significant differences (P < 0.05) between Hepatozoon-infected and uninfected crocodiles by 1-way analysis of variance (ANOVA). The residuals were checked for normality of distribution with normal probability plots. Where data were not normally distributed, significance was tested by means of a Mann-Whitney test.
The crocodiles ranged in TL from 34.0 cm to 46.3 cm, with a mean of 59.7 cm, and ranged in SVL from 25.5 cm to 101.5 cm with a mean of 28.4 cm.
Of the 144 crocodiles caught and examined, none were visibly sick or displayed any clinical signs of disease. The body condition of all the crocodiles was good. The only external lesions observed were old healed bite injuries in one case, and a recent puncture injury in another case.
The only ectoparasite found on the Nile crocodiles examined was the leech Placobdelloides multistriatus. The leeches were identified with the aid of a key37,38. Eleven crocodiles were infested with the leech, a prevalence of 7.6 %. One crocodile had 7 leeches, and another had 2. The remaining 9 parasitised crocodiles each had a single leech. Leeches were found in various places on the crocodiles, both dorsally and ventrally, for example on the tail, neck, belly, armpits, between webbing of back legs, and other sites. There was no obvious pattern of leech distribution on the crocodiles. No correlation between current leech infestation and H. pettiti infection was found.
No antibodies to Mycoplasma crocodyli were detected in any of the sera tested.
Hepatozoon pettiti parasites were present in 21 of 38 blood smears examined (55.3 %). The mean SVL of the 38 crocodiles tested was 42.6 cm. The mean SVL of the infected crocodiles was 46.3 cm compared with 38.2 cm for the uninfected crocodiles. The youngest infected crocodile in this study had a SVL of 25.5 cm.
Six of 8 females (75 %) and 13 of 27 males (48 %) were infected. The sex of 2 crocodiles was undetermined due to their small size.
Two (9.5 %) of the infected crocodiles were infested with leeches, as were 4 (23.5 %) of crocodiles negative for H. pettiti.
Of the 21 infected crocodiles, 10 (47.6 %) showed an increased rate of red cell regeneration, with a mean score of 2.4 on the erythrocyte regeneration index. In contrast, 6 of 17 (35.3 %) uninfected crocodiles displayed an increased rate of red cell regeneration, with a mean score of 2.3 on the erythrocyte regeneration index. This was not a significant difference.
There was no significant difference between any of the haematological values of the infected and uninfected crocodiles (Table 1). The mean PCV and RCC of the infected group were 18.2 % and 0.62 × 106/µ, respectively, compared with 17.6 % and 0.56 × 106 /µ in the uninfected group.
Owing to the limitations mentioned in the introduction it is probable that a capture and release survey does not provide a conclusive reflection of disease prevalence in a crocodile population. On the other hand, a very low disease incidence in wild crocodiles is probable because stress is a very important predisposing factor to disease in crocodiles. Crocodilians have been found to respond to non-specific stress with a chronic increase in corticosterone secretion, inhibition of growth, inhibition of the reproductive system and suppression of the immune system30. Crocodiles living in a relatively pristine natural environment, with a low population density, will not be exposed to the stress experienced by farmed crocodiles in an artificial, intensive environment. Furthermore, transmission of infectious diseases under natural conditions is usually far slower due to less host-pathogen exposure compared with an intensive situation.
The 1st recorded outbreak of mycoplasmosis in crocodiles occurred in Zimbabwe on 5 farms simultaneously34. Rearing stock 1-3 years of age developed swollen limb joints and lameness. Morbidity was 10 % and mortality even lower. A new species of Mycoplasma was cultured from the joints of affected animals and named Mycoplasma crocodyli26. The disease was then reproduced in healthy crocodiles by experimental infection with this isolate.
Also in 1995 a highly fatal outbreak of disease, characterised by arthritis and pneumonia, occurred in farmed alligators (Alligator mississippiensis) in Florida2. Nine out of 74 adult animals died over a 10-day period. A new species of Mycoplasma was isolated and later named M. alligatoris3. Losses continued over the next 6 months, despite tetracycline treatment, until only 14 alligators remained. An enzyme-linked immunosorbent assay (ELISA) for the detection of antibodies produced by alligators in response to M. alligatoris exposure has since been developed4.
Following the initial outbreak in Zimbabwe, cases of mycoplasmosis were reported annually, from about 1/3 of Zimbabwean crocodile farms. An autogenous vaccine was developed which, in an experimental trial, afforded limited protection35. A subsequent severe outbreak occurred in crocodiles following translocation, in which the morbidity was over 50 % and the mortality rate over 20 %33. Subsequently mycoplasmosis has become an important disease on South African crocodile farms, with several severe outbreaks having occurred (J Picard, University of Pretoria, pers. comm., 2006).
The epidemiology of this disease is not well understood. The source of the original Zimbabwean outbreak remains elusive. It was suggested that wild crocodiles may act as reservoirs of infection, with vertical transmission being the main mode of transmission35. However, unidentified mycoplasmas have been found in the faeces of farmed Nile crocodiles, indicating the possibility of horizontal transmission22. The concept of wild crocodilian reservoirs of infection has more recently been confirmed in the case of M. alligatoris. In a seroprevalence study, 5.4 % of wild American alligators were found to be positive for M. alligatoris antibodies, at 12 of 20 sites (60 %)5. Further elucidation of the epidemiology of mycoplasmosis in Nile crocodiles requires that the status of wild crocodiles be established.
Many crocodile farms in southern Africa (excluding South Africa) collect eggs from wild crocodile nests. If the wild population concerned is infected, this is a very risky practice. Often these farms are obliged to reintroduce a certain number of juvenile individuals into the wild. If, on the other hand, the wild population is free from mycoplasmosis, it could be disastrous to reintroduce infected juveniles.
At the time these samples were tested, the M. crocodyli plate agglutination test had just been developed. It was believed that the test could identify recently infected animals, but not old infections. Subsequent testing on a positive farm revealed that the test only identified very recent infection, as only animals that still had lesions tested positive (J Picard, pers. comm., 2006). The negative results obtained here do not, therefore, exclude historical infection with mycoplasms. The Mycoplasma infection status of the Okavango crocodiles therefore remains uncertain. An indirect enzyme-linked immunosorbent assay (iELISA) test for M. crocodyli was recently developed11. This test has a high sensitivity and specificity (86 % and 100 %, respectively) and will be ideal for further testing of the Okavango population.
The Samochima crocodile farm in Botswana collects a maximum of 2000 eggs per nesting season from nests in the Panhandle region of the Okavango Delta. In return the farm is obliged to release 5 % of juvenile crocodiles back into the wild. Testing of these juveniles should be undertaken before release to avoid the risk of infecting the wild population.
Haemogregarines are protozoal blood parasites. Members of the genus Hepatozoon occur in various crocodilians. Asexual schizonts are found in the liver of infected crocodiles and gametocytes are found in erythrocytes or free in the blood. Sexual multiplication occurs in the intermediate host, usually haematophagous insects and leeches25,29.
Hepatozoon pettiti was described in Nile crocodiles from Senegal and Uganda47 and H. sheppardi was described from Nile crocodiles in Mozambique44. Hoare19 was the first to show transmission of H. pettiti by the tsetse fly, Glossina palpalis. A later study14 suggested that the leech P. multilineata was a vector of haemogregarines in alligators and this was supported by a further study a few years later15. Fairly recently it was confirmed that the species found in crocodiles in the Okavango is H. pettiti, although the vector is unknown at this stage16. Hepatozoon parasites are thought to be apathogenic in their crocodilian hosts. In this survey, the aim was to determine the prevalence of H. pettiti and its possible effect on the crocodile host.
In a study which ran concurrently to this one16, the authors found H. pettiti in 61 out of 186 Nile crocodiles (32.8 %). As in this study, no significant difference in the PCV of infected and uninfected groups was found. It is clear that there is a high prevalence of H. pettiti infection in the Okavango crocodile population, and that it appears not to be pathogenic to the crocodile host. The mean SVL of the Hepatozoon-infected crocodiles was greater than that of the uninfected group. It would appear that crocodiles do not lose the infection with increasing age.
A very high prevalence of Hepatozoon infection is not uncommon in other species. Haemogregarina crocodilinorum was found to be widely distributed in the southern United States and was found in 77 (59 %) of 130 American alligators examined25. In another study1 a Haemogregarina sp. was found in all of 9 wild alligators captured. They were anaemic compared with captive-bred controls. It is not clear whether the haemogregarine played a role in the anaemia, or whether it was caused solely by leech infestation and a lower nutritional plane of the wild alligators. A prevalence of 71.4 % was reported for H. caimani in caimans (Caiman crocodilus) in Western Brazil49 and very recently a prevalence of 76 % was reported in C. yacare in Central Brazil50. A haemogregarine was found in 16 of 25 (64 %) salt-and fresh-water crocodiles (C. porosus and C. novaeguineae)28.
The ectoparasites of crocodilians have not been well studied and most reports to date refer primarily to ticks and leeches. However, only 7 publications have addressed tick parasitism of crocodilians43. Leeches are a more common crocodilian ectoparasite and have been found on a few species including C. porosus52, Alligator mississippiensis14,15,25, C. johnstoni51 and C. niloticus36,37. However, little is known about the possible pathogenicity of the various species, although leeches could possibly play a role in the transmission of crocodile-specific viral and bacterial infections21 and as vectors of blood protozoans25.
Placobdelloides multistriatus has been recorded as a parasite on crocodiles in Africa 10,36,37 although there is no previous record of its occurrence on crocodiles in the Okavango region. A next step would be to determine whether P. multistriatus is a potential vector for H. pettiti infection in the Okavango's Nile crocodiles.
Further research is required on several of the crocodilian diseases to elucidate their epidemiology, and particularly the role of wild crocodilians as reservoirs of infection. Pox viruses, Adenovirus, West Nile virus, chlamydiosis and mycoplasmosis are all of particular interest.
This research was supported by a National Research Foundation and an Earthwatch Institute (USA) grant awarded to A J Leslie. The South African Veterinary Foundation and the Johannesburg Zoological Gardens are acknowledged for additional financial support. We thank Prof. Daan Nel (University of Stellenbosch) for helping with the statistical analysis and the Government of Botswana for providing the necessary research permits.
1. Barnett J D, Cardeilhac P T, Barr B, Wolff W, Bass O L 1999 Differences between captive-raised and wild-caught Everglades National Park alligators in serum chemistry values, serum protein electrophoresis, thyroid hormone levels, and complete blood counts. Proceedings of the International Association of Aquatic Animal Medicine, Boston, Massachusetts, USA 30: 136-139 [ Links ]
2. Brown D R, Clippinger T L, Helmick K E, Schumacher I M, Bennett R A, Johnson C M, Vliet K A, Jacobson E R, Brown M B 1996 Mycoplasma isolation during a fatal epizootic of captive alligators (Alligator mississippiensis) in Florida. International Organisation of Mycoplasmology Letters 4: 42-43 [ Links ]
3. Brown D R, Farley J M, Zache L A, Carlton J M-R, Clippinger T L, Tully J G, Brown M B 2001 Mycoplasma alligatoris sp. nov., from American alligators. International Journal of Systematic and Evolutionary Microbiology 51: 419-424 [ Links ]
4. Brown D R, Schumacher I M, Nogueira M F, Richey L J, Zacher L A, Schoeb T R, Vliet K A, Bennett R A, Jacobson E R, Brown M B 2001 Detection of antibodies to a pathogenic mycoplasma in American alligators (Alligator mississippiensis), broad-nosed caimans (Caiman latirostris), and Siamese crocodiles (Crocodylus siamensis). Journal of Clinical Microbiology 39: 285-292 [ Links ]
5. Brown D R, Zacher L A Carbonneau D A 2005 Seroprevalence of Mycoplasma alligatoris among free-ranging alligators (Alligator mississippiensis) in Florida-2003. Journal of Zoo and Wildlife Medicine 36: 340-341 [ Links ]
6. Buenviaje G N, Ladds P W, Martin Y 1998 Pathology of skin diseases in crocodiles. Australian Veterinary Journal 76: 357-363 [ Links ]
7. Buenviaje G N, Ladds P W, Melville L, Manolis S C 1994 Disease-husbandry associations in farmed crocodiles in Queens-land and the Northern Territory. Australian Veterinary Journal 71: 165-173 [ Links ]
8. Campbell T W 1995 Avian hematology and cytology (2nd edn). Iowa State University Press, Ames [ Links ]
9. Campbell T W 1996 Clinical pathology. In Mader D R (ed.) Reptile medicine and surgery, W B Saunders, Philadelphia: 248-257 [ Links ]
10. Cott H B 1961 Scientific results of an enquiry into the ecology and economic status of the Nile crocodile (Crocodylus niloticus)in Uganda and Northern Rhodesia. Transactions of the Zoological Society of London 29: 211-356 [ Links ]
11. Dawo F, Mohan K 2007 Development and application of an indirect ELISA test for the detection of antibodies to Mycoplasma crocodyli infection in crocodiles (Crocodylus niloticus). Veterinary Microbiology 119: 283-289 [ Links ]
12. Foggin C M 1987 Diseases and disease control on crocodile farms in Zimbabwe. In Webb,GJW, Manolis S C, Whitehead P J (eds) Wildlife management: crocodiles and alligators. Surrey Beattie & Sons, Chipping Norton: 351-356 [ Links ]
13. Foggin C M 1992 Diseases of farmed crocodiles. In: Smith G A, Marais J (eds) Conservation and utilization of the Nile crocodile in South Africa. Handbook on crocodile farming. The Crocodile Study Group of Southern Africa, Pretoria: 107-140 [ Links ]
14. Forrester D J, Sawyer R T 1974 Placobdella multilineata (Hirudinea) from the American alligator in Florida. Journal of Parasitology 60: 673 [ Links ]
15. Glassman A B, Holbrook T W, Bennett C E 1979 Correlation of leech infestation and eosinophilia in alligators. Journal of Parasitology 65: 323-324 [ Links ]
16. Gomersall D P, Smit N J, Leslie A J 2006 Prevalence, distribution and possible vector of Hepatozoon pettiti, blood parasite of Nile crocodiles in the Okavango Delta. Journal of the South African Veterinary Association 77: 93 [ Links ]
17. Graham A, Simbotwe M, Hutton J M 1992 Monitoring of an exploited crocodile population on the Okavango River, Botswana. In Hutton J M, Games I (eds) The CITES Nile Crocodile Project. Secretariat of the Convention on International Trade in Endangered Species of Wild Fauna and Flora, Lausanne: 53-69 [ Links ]
18. Groombridge B 1987 The distribution and status of world crocodilians. In WebbGJW, Manolis S C, Whitehead P J (eds), Wildlife management: crocodiles and alligators. Surrey Beatty & Sons, Chipping Norton: 9-21 [ Links ]
19. Hoare C A 1932 On protozoal blood para sites collected in Uganda. Parasitology 24: 210-224 [ Links ]
20. Huchzermeyer F W 2002 Diseases of farmed crocodiles and ostriches. Revue scientifique et technique (International Office of Epizootics) 21: 265-276 [ Links ]
21. Huchzermeyer F W 2003 Crocodiles: biology, husbandry and diseases. CABI Publishing, Wallingford [ Links ]
22. Huchzermeyer F W, Gerdes G H, Putterill J F 1994 Viruses and mycoplasmas from faeces of farmed Nile crocodiles. In Proceedings of the 12th Working Meeting of the Crocodile Specialist Group, Pattaya, Thailand (unedited), 2-4 May 1994, Vol. 2. IUCN - The World Conservation Union, Gland: 303-308 [ Links ]
23. Hutton J M 1987 Incubation temperatures, sex ratios, and sex determination in a population of Nile crocodiles (Crocodylus niloticus). Journal of Zoology 211: 143-155 [ Links ]
24. Jain N C 1986 Hematological techniques. Schalm's Veterinary hematology. Lea and Febiger, Philadelphia: 36-66 [ Links ]
25. Khan R A, Forrester D J, Goodwin T M, Ross C A 1980 A haemogregarine from the American alligator (Alligator mississippiensis). Journal of Parasitology 66: 324-328 [ Links ]
26. Kirchhoff H, Mohan K, Schmidt R, Runge M, Brown D R, Brown M B, Foggin C M, Muvavarirwa P, Lehmann H, Flossdorf J 1997 Mycoplasma crocodyli sp. nov., a new species from crocodiles. International Journal of Systematic Bacteriology 47: 742-746 [ Links ]
27. Ladds P W, Mangunwirjo H, Sebayang D, Daniels P W 1995 Diseases of young farmed crocodiles in Irian Jaya. Veterinary Record 136: 121-124 [ Links ]
28. Ladds P W, Simms L D 1990 Diseases of young captive crocodiles in Papua New Guinea. Australian Veterinary Journal 67: 323-330 [ Links ]
29. Lainson R, Paperna I, Naiff R D 2003 Development of Hepatozoon caimani (Carini, 1909) Pessoa, De Biasi & De Souza, 1972 in the caiman Caiman c. crocodilus, the frog Rana catesbeiana and the mosquito Culex fatigans. Memorias do Instituto Oswaldo Cruz 98: 103-113. [ Links ]
30. Lance V A 1990 Stress in reptiles. In Epple A, Scanes C G, Stetson M H (eds) Progress in comparative endocrinology. Wiley-Liss, New York: 461-466. [ Links ]
31. Leslie A J 1997 Ecology and physiology of the Nile crocodile, Crocodylus niloticus, in Lake St. Lucia, KwaZulu/Natal, South Africa. PhD thesis, Drexel University, Philadelphia [ Links ]
32. Mendelsohn J, el Obeid S 2004 Okavango River, the flow of a lifeline. Struik Publishers, Cape Town [ Links ]
33. Mohan K, Foggin C M, Dziva F, Muvavarirwa P 2001 Vaccination to control an outbreak of Mycoplasm crocodyli infection. Onderstepoort Journal of Veterinary Research 68: 149-150 [ Links ]
34. Mohan K, Foggin C M, Muvavarirwa P, Honywill J, Pawandiwa A 1995 Mycoplasma-associated polyarthritis in farmed crocodiles (Crocodylus niloticus) in Zimbabwe. Onderstepoort Journal of Veterinary Research 62: 45-49 [ Links ]
35. Mohan K, Foggin C M, Muvavarirwa R, Honywill J 1997 Vaccination of farmed crocodiles (Crocodylus niloticus) against Mycoplasma crocodyli infection. Veterinary Record 141: 476 [ Links ]
36. Moore J P 1938 Additions to our knowledge of African leeches (Hirudinea). Proceedings of the Academy of Natural Sciences of Philadelphia 90: 297-360 [ Links ]
37 Oosthuizen J H 1979 Redescription of Placobdella multistriata (Johansson, 1909) (Hirudinoidea: Glossiphoniidae). Koedoe 22: 61-79 [ Links ]
38. Oosthuizen J H 1991 An annotated check list of the leeches (Annelida: Hirudinea) of the Kruger National Park with a key to the species. Koedoe 34: 25-38 [ Links ]
39. Pooley A C 1977 Nest opening response of the Nile crocodile Crocodylus niloticus. Journal of Zoology (London) 182: 17-26 [ Links ]
40. Pooley A C, Gans C 1976 The Nile crocodile. Scientific American 234: 114-124 [ Links ]
41. Pooley A C 1982 The ecology of the Nile crocodile, Crocodylus niloticus, in Zululand. MSc thesis, University of Natal, Pietermaritzburg [ Links ]
42. Pooley AC 1982 Discoveries of a crocodile man. William Collins & Sons, Johannesburg [ Links ]
43. Rainwater R R, Platt S G, Robbins R G, McMurry S T 2001 Ticks from a Morelet's crocodile in Belize. Journal of Wildlife Diseases 37: 836-839 [ Links ]
44. Santos DiasJAT 1952 Acerca de uma nova espécie de Haemogregarina, parasita dos eritrocitos do Crocodilus niloticus em Moçambique: Haemogregarina sheppardi n.sp. Anais do Instituto de Medicina Tropical 9: 181 [ Links ]
45 Richardson K C, Webb G J W, Manolis S C 2002 Crocodiles: inside out. Surrey Beatty & Sons, Chipping Norton [ Links ]
46. Taplin L E, Loveridge J P 1988 Nile crocodiles, Crocodylus niloticus, and estuarine crocodiles, Crocodylus porosus, show similar osmoregulatory responses on exposure to seawater. Comparative Biochemistry and Physiology 89A: 443-448 [ Links ]
47. Thiroux A 1910 Une haemogregarine de Crocodylus niloticus. Comptes Rendus des Seances de la Société de Biologie 69: 577-578 [ Links ]
48. Thorbjarnarson, J B 1992 Crocodiles: an action plan for their conservation. IUCN - The World Conservation Union, Gland [ Links ]
49. Viana L A, Marques E J 2005 Haemogregarine parasites (Apicomplexa: Hepatozoidae) in Caiman crocodilus yacare (Crocodilia: Alligatoridae) from Pantanal, Corumba, MS, Brazil. Revista Brasileira de Parasitologia Veterinaria 14: 173-175 [ Links ]
50. Viana L A, Paiva F, Coutinho, M E, Lourenço-de-Oliveira R 2010 Hepatozoon caimani (Apicomplexa: Hepatozoidae) in wild caiman, Caiman yacare, from the pantenal region, Brazil. Journal of Parasitology 96: 83-88 [ Links ]
51, Webb G J W, Manolis C 1983 Crocodylus johnstoni in the McKinley River area, N.T.V. Abnormalities and injuries. Australian Wildlife Research 10: 407-420 [ Links ]
52. Yang T, Davies R W 1985 Parasitism by Placobdella multilineata (Hirudinoidea; Glossiphoniidae) and its first record from Asia. Journal of Parasitology 71: 86-88 [ Links ]
Received: July 2011.
Accepted: August 2011.