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1 Department of Microbiology and Immunology, Box 44, College of Medicine, State University of New York, 450 Clarkson Avenue, Brooklyn, NY 11203, USA
2 Animal Health Laboratory, Laboratory Service Division, University of Guelph, Guelph, Ontario, Canada NIH 6R8
3 Clinic for Pigs, Small Ruminants, Forensic Medicine and Ambulatory Service, School of Veterinary Medicine, D-30173 Hannover, Germany
Correspondence
Harold Neimark
neimah25{at}hscbklyn.edu
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ABSTRACT |
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The GenBank/EMBL/DDBJ accession number for the 16S rRNA gene sequence of the Hannover strain of Mycoplasma ovis is AF338268.
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; Kreier & Ristic, 1963; Daddow, 1979; Mason et al., 1989; Mason & Statham, 1991).
As E. ovis has not been cultivated, diagnosis is made by detection of organisms on erythrocytes in Romanowsky-type or acridine orange-stained blood smears and by serological methods. Parasitism of erythrocytes often occurs at a low level and is transient; detection requires examination of repeated blood smears. However, erythrocyte parasitaemia can be as high as 100 % in our experience; a high level of parasitaemia can occur even in subclinical infection and can persist for months (Overas, 1969; Brun-Hansen et al., 1997). E. ovis is completely resistant to penicillin and other antimicrobial agents that target the cell wall. Also, tetracycline treatment does not eliminate this agent from chronically infected animals.
This organism has been thought to be a rickettsia because of its obligate parasitism, erythrocyte localization, small size, staining properties and transmission by arthropod vectors. E. ovis and a number of similar haemotrophic bacteria have been classified in the order Rickettsiales, family Anaplasmataceae, in the genera Haemobartonella and Eperythrozoon (Kreier & Ristic, 1984; Kreier et al., 1992). Recently, phylogenetic analysis of 16S rRNA gene sequences from four of these bacteria, Haemobartonella felis, Haemobartonella muris, Eperythrozoon suis and Eperythrozoon wenyonii, has demonstrated that these wall-less bacteria are not rickettsiae, but that they are actually mycoplasmas (Neimark & Kocan, 1997; Rikihisa et al., 1997; Neimark et al., 2001, 2002b). Here, the results of a phylogenetic study of E. ovis are reported.
Electron microscopy (EM)
Blood samples from E. ovis-infected sheep in Canada were used. For transmission EM, whole blood was fixed overnight in 2 % glutaraldehyde (v/v) in 0·1 M HEPES buffer, pH 7·0, at 4 °C. All solutions were used at a minimum of 10x blood volume. After fixation, samples were centrifuged gently to form a cell pellet, washed in HEPES buffer, post-fixed in 1 % osmium tetroxide (w/v) for 1 h at 4 °C, washed again in buffer and dehydrated in a graded acetone series. Samples were embedded in Spurr's resin, polymerized at 60 °C overnight and ultrathin sections (70–90 nm) were cut. Sections were mounted on grids, stained with uranyl acetate and lead citrate and examined by using a Hitachi 7100 transmission electron microscope.
Samples for scanning EM were prepared by the following method. Fresh heparin-treated blood was placed on poly-lysine-coated slides that had been moistened with distilled water and allowed to stand for 30 min in a covered Petri dish. The material was then fixed in place on the slides by immersion in 2 % glutaraldehyde (v/v) in 0·1 M HEPES buffer, pH 6·8, for 30 min. This was followed by rinsing in HEPES buffer, post-fixing with 2 % osmium tetroxide (w/v) in HEPES buffer for 30 min, rinsing again with HEPES buffer and dehydration in a graded ethanol series. Slides were critical-point-dried from 100 % ethanol by using liquid CO2 in a critical point drier, sputter-coated with 60 : 40 gold : palladium and examined with a Hitachi S-4500 field emission scanning electron microscope.
The micro-organisms, as described previously by a number of investigators (e.g. Kreier & Ristic, 1984; Zachary & Basgall, 1985), are observed on the surface of erythrocytes in Giemsa-stained blood films as small bodies of approximately 0·5–1·0 µm diameter. Morphology and epierythrocytic location of the organisms were also observed by using transmission and scanning EM (Figs 1 and 2). Transmission EM of thin sections confirmed that this coccoid organism is a prokaryote that lacks a cell wall and that these bacteria are located on the surface of the sheep erythrocyte membrane (Fig. 1). The organisms are 0·3–0·4 µm in diameter and some have a 20–30 nm wide electron-dense layer that surrounds much of the cell. Both scanning and transmission EM showed the presence of several short fibrils that formed bridges between cells and the erythrocyte surface (Figs 1 and 2); the origin of these fibrils and whether they have a function in attachment is currently not known. These fibrils are similar to fibrils observed in thin sections of mycoplasma-infected erythrocytes from squirrel monkeys (Saimiri sciureus) (Aikawa & Nussenzweig, 1972; Adams et al., 1984; Neimark et al., 2002a), pigs (Zachary & Basgall, 1985), an opossum (Messick et al., 2002), putative Haemobartonella-infected or Eperythrozoon-like-infected erythrocytes from an owl monkey (Aotus trivirgatus) (Peters et al., 1974) and cynomolgus monkeys (Macaca fascicularis) (Dillberger et al., 1994).
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Preserved blood from the sheep used for EM was not available for comparison with the samples from Germany, but stained blood smears on microscope slides were available, from which DNA was extracted by the following method. Slides were washed with 300–400 µl methanol by allowing a few drops to fall on the slide and pouring off until little colour was visible in the wash, and then air-dried. The dry blood smear was scraped with an alcohol-flamed, UV light-irradiated razor blade and the scrapings were transferred to a 1·5 ml microfuge tube that contained 100 µl TES extraction buffer (100 mM Tris, 10 mM disodium EDTA, 100 mM NaCl, pH 8) that contained 100 µg proteinase K ml-1 and 0·5 % SDS (w/v). A filter pipette with the tip melted closed was used to aid transfer and a drop of extraction buffer was added if static electricity posed problems. The volume of extraction buffer was brought to 200–400 µl and an equal volume of buffer AL (QIAamp DNA blood mini kit; Qiagen) was added. The mixture was pulse-vortexed for 15 s and placed in heat block at 55 °C for 1 h or longer until it was completely transparent (some sediment and blue colour will remain). A volume of ethanol equal to the volume of extraction buffer was added and the mixture was pulse-vortexed; it was then centrifuged (6000 g for 1 min) to pellet the sediment and the supernatant was transferred to a blood mini kit spin column in a 2 ml collection tube (Qiagen). If more than one pipette transfer was required, the sample tube was centrifuged after each withdrawal to prevent transfer of sediment. The column was centrifuged (6000 g for 1 min) and transferred to a clean 2 ml collection tube; the extraction continued with step 8 as described in the Qiagen protocol. DNA was eluted from the column with 50 µl TE buffer (pH 9).
Rigorous precautions were taken to avoid PCR contamination. 16S rRNA genes were amplified (31 cycles: 94 °C, 1 min; 55 °C, 1 min; 72 °C, 3 min; with a 10 min extension for the last cycle) by using two conserved primers: 8F (5'-AGAGTTTGATCMTGGCTCAG-3') and 1492R (5'-CGGTTACCTTGTTACGACTT-3'). PCRs (50 µl) contained: 10 mM Tris/HCl, pH 8·3; 50 mM KCl; 2·5 mM MgCl2; 200 µM each of dATP, dCTP, dGTP and dTTP; 50 pM each primer; and 2·5 U Taq polymerase (Perkin Elmer). Positive and negative controls were included in all amplifications.
PCR products were purified with a spin column (Qiagen) and DNA was sequenced in both directions with a sequencing kit in an automated sequencer (Applied Biosystems). Sequencing primers were 8F, 339R, 360F, 519R, 482F, 797F, 907R, 1083R, 1088F and 1492R; primers 340F (5'-CCATATTCCTACGGGAAGCA-3') and 543R (5'-CCGGATAATGCTTGTGACCT-3') were also used for amplification and sequencing (see results for the nucleotide position numbering of these two primers).
Mycoplasma 16S rRNA gene sequences were obtained from GenBank. Alignments were made with the program CLUSTALW (Thompson et al., 1994) and adjusted manually. Phylogenetic analysis of the sequence alignment was carried out with the phylogenetic program package PHYLIP 3.51c (Felsenstein, 1993). The phylogenetic tree was constructed by using the neighbour-joining method, corrected for nucleotide substitutions by the Kimura two-parameter option with the transition/transversion ratio set at 2. The dataset was resampled 1000 times and the resulting bootstrap percentage values are indicated at nodes of the tree (Fig. 3).
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Comparison of the 16S rRNA gene sequence of E. ovis Hannover to sequences in GenBank by using BLASTN showed that nearly all matches were to mycoplasmas; highest similarity scores were obtained with haemotrophic mycoplasmas. Phylogenetic analysis produced an evolutionary tree that showed the relationship between E. ovis, other haemotrophic mycoplasmas, representative species of the pneumoniae group and representatives of other mycoplasma groups (Fig. 3
). The tree shows that the E. ovis 16S rRNA gene sequence clusters with those of other previously identified haemotrophic mycoplasmas (Neimark & Kocan, 1997; Rikihisa et al., 1997) on a branch within the pneumoniae group of the genus Mycoplasma (Neimark & Kocan, 1997; Johansson et al., 1999; Neimark et al., 2001). The phylogenetic tree shows that, among known haemotrophic mycoplasmas, E. ovis is related most closely to Mycoplasma wenyonii (95 % similarity). Reported cross-reactions between these two blood parasites with fluorescein-labelled sera from E. ovis- or M. wenyonii-infected animals (Kreier & Ristic, 1963) might be a reflection of this relationship.
High bootstrap values at most of the branch-points in the phylogenetic tree (Fig. 3) indicate that this is probably a robust tree. However, the few low bootstrap values (<50 %) at some branch-points suggest that these branches may be due to chance.
Over the years, bodies observed on erythrocytes from nearly 30 different vertebrate hosts (ranging from fish to primates) and presumed to be Haemobartonella or Eperythrozoon species have been described and given species names on the sole basis of observing them in Giemsa-stained blood smears (Tyzzer, 1942; Weinman, 1944; Kreier et al., 1981). Although a few of these bodies may have been staining artefacts or misidentified erythrocytic structures, such as Pappenheimer bodies, Heinz bodies or Howell–Jolly bodies, it seems likely that the majority represent authentic haemotrophic bacteria that remain to be characterized. More recent observations of haemotrophic bacteria have been documented by experimental infection and EM studies, which have demonstrated wall-less bacteria on the surface of host erythrocytes (e.g. on raccoon erythrocytes; Frerichs & Holbrook, 1971). Recently, haemotrophic organisms observed on erythrocytes from an opossum (Messick et al., 2002), llamas (Reagan et al., 1990; McLaughlin et al., 1991; Messick et al., 2002) and South American squirrel monkeys (Neimark et al., 2002b) have been identified as Mycoplasma species by molecular methods. It is proposed that classification of these haemotrophic bacteria should be changed, in order to reflect their actual phylogenetic position as mycoplasmas (Neimark et al., 2001, 2002b). These results support the suggestion that the remaining species of Haemobartonella (‘Haemobartonella procyoni’) and Eperythrozoon (Eperythrozoon parvum), both of which are wall-less and share identical properties with the haemotrophic mycoplasmas, will probably be found to be mycoplasmas (Neimark et al., 2001).
Mycoplasmas (class Mollicutes) comprise a large group of >110 species of wall-less bacteria that occur as pathogens or commensals in a wide variety of vertebrate animals, insects and plants. Mycoplasmas are notable for their low DNA G+C content (Neimark, 1970) and small chromosomes (Pyle et al., 1988; Neimark & Lange, 1990). They have evolved through large losses of chromosomal DNA from low-G+C-containing, Gram-positive bacteria (Neimark, 1979; Woese, 1980) and are related most closely to streptococci (Neimark, 1979; Neimark & London, 1982; Maniloff, 1996). All known animal mycoplasmas are parasites on mucous membranes of vertebrates; thus, the haemotrophic mycoplasmas display a novel mode of parasitism that has not been recognized previously in the mollicutes and have enlarged our understanding of the pathogenic capacity of mycoplasmas.
Identification of E. ovis as a mycoplasma provides a new perspective for clinicians and investigators in dealing with this organism, as it immediately allows information about mycoplasmas to be applied to this bacterium. Similarly, this identification augments our knowledge of this novel group of blood-parasitic mycoplasmas.
Whether the natural host range of E. ovis extends beyond sheep and goats is unclear. E. ovis has not been reported to infect cattle and is not infectious in calves, even when they are splenectomized (under the unnatural condition of splenectomy, the parasite was found for 9 days in the blood of a calf) (Hoyte, 1971). A splenectomized deer (Dama virginiana) infected with sheep blood that contained E. ovis was reported to yield positive blood films (Kreier & Ristic, 1963). Also, an eland (Taurotragus oryx) inoculated with sheep blood that contained both Anaplasma ovis and E. ovis did not show any clinical symptoms, and neither organism was detected in daily blood smears after 50 days, but a sheep inoculated with day 51 blood from the eland eventually displayed both organisms in blood smears and developed fever, lethargy and loss of appetite (Enigk, 1942). In retrospect, the possibility that this sheep was infected from another source cannot be ruled out.
It is proposed that E. ovis should be transferred to the genus Mycoplasma as Mycoplasma ovis comb. nov., with the specific epithet ‘ovis’ retained for continuity. This name-change involves the transfer of a bacterial species that was previously recognized on the Approved Lists of Bacterial Species (Skerman et al., 1980); consequently, the Candidatus designation is not appropriate for this bacterium, as this designation is for novel, incompletely described taxa, to give them a provisional status (Neimark et al., 2001a).
Description of Mycoplasma ovis comb. nov. (basonym Eperythrozoon ovis Neitz, Alexander and du Toit 1934)
Mycoplasma ovis (o'vis. L. fem. n. ovis a sheep).
Epierythrocytic parasite of sheep and goats. Giemsa stain shows small, round bodies attached to the surface of erythrocytes. Erythrocytes can reach 100 % infection. Organisms also are observed free in the blood during periods of fever in heavily parasitized animals. By using phase-contrast microscopy, M. ovis appears as a round cell. Cells seen in thin sections by EM are round or oval bodies of 0·3–0·4 µm diameter and are applied closely to the erythrocyte membrane. A 20–30 nm wide electron-dense layer partially surrounds many of the organisms. M. ovis can produce mild to severe anaemia in normal sheep. Infections are frequently unapparent. Mortality from infection is rare, but anaemia and failure to gain weight occur in lambs. M. ovis is reported to share antigens with M. wenyonii (Kreier & Ristic, 1963). Organisms in blood can be preserved by freezing untreated blood at -18 °C (M. Ganter, unpublished results) or in 10 % DMSO (v/v) at -70 °C (Brun-Hansen et al., 1997); 10 % horse serum/10 % glycerol, which is used for freezing other mycoplasmas at -70 °C, should also be suitable. Transmission is by blood-feeding arthropods, including ticks [Haemaphysalis plumbeum and Rhipicephalus bursa (Nikol'skii & Slipchenko, 1969)] and mosquitoes [Aedes camptorhynchus (Howard, 1975) and Culex annulirostris (Daddow, 1980)]. The stable fly (Stomoxis calcitrans) would appear to be a potential vector and subcutaneous injection of sheep with a suspension of fly material has resulted in infection, but transmission under natural conditions has not been demonstrated (Overas, 1969); similarly, injection of head lice (Linognathus ovillas) has also produced infection in sheep (Foogie & Nisbet, 1964). An attempt to transmit the disease by removing sheep keds (the blood-sucking wingless fly Melophagus ovinus) from carrier sheep and injecting them intravenously into a splenectomized sheep (Neitz, 1937) or a normal susceptible sheep (Foogie & Nisbet, 1964) did not produce visible infection. Infections can be transmitted by reusing needles during herd immunization and as a very small number of infected erythrocytes is sufficient to transmit disease (Mason & Statham, 1991), it has been suggested that blood on tools during shearing or ear-tagging may contribute to spread in flocks (Brun-Hansen et al., 1997). Oral transmission has been shown experimentally (Overas, 1969). M. ovis occurs in sheep-rearing regions throughout the world.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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