Avian Medicine: Princilpes and Application - IVIS · tious diseases are less of a problem in birds...

BRANSON W. RITCHIE, DVM, PhD Assistant Professor, Avian and Zoologic Medicine Department of Small Animal Medicine College of Veterinary Medicine University of Georgia Athens, Georgia

GREG J. HARRISON, DVM Director, The Bird Hospital Lake Worth, Florida President, Harrison’s Bird Diets Omaha, Nebraska

LINDA R. HARRISON, BS President, Wingers Publishing, Inc. Former Editor, Journal of the Association of Avian Veterinarians Lake Worth, Florida

AVIAN MEDICINE: PRINCIPLES AND APPLICATION

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he mycoplasmatales constitutes one orderwithin the class Mollicutes that replicatesmainly by binary fission. Strains that pro-duce mycelia-like forms may propagate by

dissociation of these “mycelia.” Morphologically, colo-nies and single organisms can exist in multiple forms(coccoid, rods, ring-forms), depending on the physicalproperties of the media in which they are growing. Inmost cases, morphology is unsuitable for species dif-ferentiation. In contrast to bacteria, mycoplas-matales have no cell wall and are bound by a three-layer membrane. Thus, they are resistant toantibiotics that inhibit cell wall development (eg,penicillins, cephalosporins, bacitracin) and sul-fonamides. Mycoplasmatales are fastidious and mustobtain most of the nutrient requirements from thegrowth media because of their relatively smallgenome. They grow on agar media in small, friedegg-shaped colonies, which in many instances, can berecognized only under the microscope. Generally,specialized laboratories are necessary for isolationand identification. The lack of a cell wall makes theorganism sensitive to inactivation outside the host (itsurvives only hours on dry surfaces, two to four daysin water); therefore, transport media are necessaryfor shipping infected tissues intended for isolationattempts. Mycoplasmatales that are free in the envi-ronment are susceptible to all commonly used disin-fectants. Organisms within host excretions are pro-tected from contact with the disinfectant. Secretionsand excretions must be removed before disinfectingprocedures are effective.

The mycoplasmatales consist of three genera, whichcan be distinguished roughly by the following proper-ties:

Mycoplasma need cholesterol for growth (productionof the cellular membrane).

Acholeplasma do not need cholesterol for growth, butmany strains can be inhibited by the thallium ace-tate that is commonly used for inhibiting gram-nega-tive bacteria in media used for the isolation of myco-plasma.

Ureaplasma were formerly called T-strains becauseof their tiny colony sizes. They require urea for theirenergy metabolism and also cholesterol for growth.

T C H A P T E R

38MYCOPLASMA AND

RICKETTSIA

Helga Gerlach

Published in IVIS with the permission of the editorClose window to return to IVIS www.ivis.org

Avian Medecine : Principles and Application, B.W. Ritchie, G.J Harrison and L.R. Harrison (Eds.)

Many isolates from companion and aviary birds havenot yet been fully identified and have no valid name.In addition, the pathogenicity and epizootiology ofthese strains have not been defined to date.

Mycoplasmatales are distributed worldwide in con-nection with the poultry industry. There is little in-formation on the prevalence of mycoplasmatales incaptive or free-ranging Psittaciformes or othergroups of birds. Isolations have been rare, and theimportance of the majority of the strains is unknown.With intensified aviculture, increased farm sizes andpopulation densities on these farms, more problemswith mycoplasmatales can be expected.

Mycoplasmatales

The host spectrum of the mycoplasmatales is rathernarrow (see Table 38.1), with the exception of Myco-plasma cloacale and the genus Acholeplasma. Re-ports suggesting isolates of well known species fromunusual hosts should be met with skepticism. Thevarious mycoplasmatales have similar biochemicalproperties and serologically cross-react with otherspecies of the order, creating a high number of false-positive results (low specificity). The reason for thesecross reactions is that the lack of a cell wall diminishesthe antigenicity, which is probably governed by en-zymes within the microorganisms. Because these en-zymes are phylogenetically old and highly conserved,they do not vary much between genera. Physical meth-ods such as electrophoresis (combined with blot meth-ods) are more reliable than serologic methods for differ-entiating between species or strains.28

Transmission

Mycoplasmatales are relatively low in infectivity.Close contact between individuals is necessary fortransmission, and infections are most common indense populations (Figure 38.1). The respiratory andgenital tracts are the primary portals of entrance.The organism is spread by respiratory excretions andby the gonads of both sexes as well as hematologicallythrough the body. Infected air sacs can lead to contacttransmission of the ovary (and developing follicle).Transovarian transmission is epornitically impor-tant, although in clinically healthy breeders, the egg

transmission rate is low (between 0.1 and 1.0 %);however, there are some exceptions. The egg trans-mission rate of M. meleagridis in turkeys can be ashigh as 25%. This species causes predominantly avenereal disease. Infected breeders may be asympto-matic. Close contact is the primary mode of transmis-sion in neonates. Offspring feeding on contaminatedcrop regurgitations (eg, crop milk in pigeons) mayalso become infected.

Pathogenesis

Primary pathogenic strains, ie, strains that can dam-age epithelial cells and cause disease without addi-tional factors, have to be distinguished from secon-dary pathogenic strains that need predamagedepithelium, and from strains that are assumed to beapathogenic. Mycoplasmatales preferably colonizethe mucosa of the respiratory and the genital tracts.Strains capable of inducing systemic infections canbe found in the brain and joints. Infections start withthe adsorption of the organism to the surface of hostcells (including erythrocytes with hemagglutinatingstrains). Multiplication takes place on the cell sur-face, and both the membrane integrity as well as thefunction of the host cell can be altered. Because theagent may be hidden in the recesses of the host cellmembrane, it can remain rather inaccessible bytherapeutics and the host defense mechanisms. As aconsequence, only negligible amounts of humoral an-tibodies, if any, are produced. M. gallisepticum (and

FIG 38.1 High density, confined, indoor breeding operations in-crease the exposure of individual birds to a mixture of microorgan-isms that may include Mycoplasma spp. Damage to the respiratorytract caused by increased dust, dry-heated air and respiratory viralinfections predispose birds to mycoplasma infections. Most infec-tious diseases are less of a problem in birds maintained in lowdensity outdoor breeding facilities (reprinted with permission JAssoc Avian Vet).

SECTION FIVE DISEASE ETIOLOGIES

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TABLE 38.1 Avian Host Spectrum of Mycoplasmatales13,17

Species Host Spectrum Signs of Disease

M. gallisepticum Chicken, turkey, guineafowl, peafowl, pheasants,partridge, rock partridge, Red-legged Partridge,Japanese Quail, Bobwhite Quail, House Sparrow,domestic duck and goose,5 Canada Goose5

Rhinitis, sinusitis, tracheitis, air sacculitis,pneumonia, arthritis, encephalitis, ophthalmitis

M. gallinarum Chicken, pheasant, Chinese Bamboo Partridge, HouseSparrow, Demoiselle Crane, Domestic Goose, Bewick’sSwan

Mild respiratory signs in geese also infected withparvovirus

M. pullorum Chicken, Turkey, Pheasant, Partridge Asymptomatic

M. gallinaceum Chicken, Turkey, Pheasant, Hoopoe Asymptomatic: complicated with PMV1

M. iners Chicken, Turkey, Domestic Goose,2 Golden Pheasant31 Asymptomatic

M. gallopavonis Turkey, (Chicken?) Mild respiratory signs only in turkeys

M. meleagridis Turkey Sinusitis, air sacculitis, infection of the genital tract

M. iowae Chicken, Turkey, Yellow-crowned Amazon Parrot Mild air sacculitis, in turkeys veneral transmissionand reduced hatchability

M. columbinasale Pigeon Rhinitis, pharyngitis

M. columborale Pigeon (Chicken ?) Rhinitis, pharyngitis

M. columbinum Pigeon Asymptomatic

M. synoviae Chicken, Turkey,23 Guineafowl, Red-legged Partridge,Japanese Quail,2 House Sparrow,31 Tree Sparrow,31

Domestic Duck and Goose

Sinusitis, synovitis, air sacculitis, hepatitis,splenomegaly

M. anatis Domestic Duck, Greater Scaup, Common Teal and otherTeals,31 Domestic Goose,2 Coot,31 Common Shoveler31

Rhinitis, sinusitis, air sacculitis only if triggered byinfluenza virus

M. glycophilum Chicken

M. lipofaciens Chicken

M. cloacale Turkey,2 Domestic Duck and Goose,2 Tufted Duck,European Pochard, Muscovy Duck, Skylark, Starling,Cockatiel, Lesser Spotted Woodpecker4

M. anseris Domestic Goose Together with M. cloacale lesions in geese

M. spp. n.n.(7 different types)

Domestic Duck Mild respiratory signs

M. sp. n.n. Budgerigar Air sacculitis

A. laidlawii B(var. inocuum)

Chicken, Pigeon, Greater Adjutant Stork, Night Heron Asymptomatic

A. laidlawii A Domestic Duck and Goose Air sacculitis, conjunctivitis, cloacitis

A. axanthum Domestic GooseDomestic Duck

Embryonal death, peritonitis, salpingitis, air sacculitisConjunctivitis, cloacitis

A. equifetale Chicken

A. spp. n.n.(2 different types)

Pigeon Rhinitis, pharyngitis

U. gallorale Chicken, Red Junglefowl Air sacculitis, pneumonia

U. spp. n.n. Turkey, Jungle Bush Quail Respiratory signs

Unidentified(3 different types)

Severe Macaw, Cockatoo spp., Cockatiel, Canary Chronic conjunctivitis, rhinitis, sneezing, sinusitis,dyspnea, arthritis

Several types ? Saker Falcon, Peregrine Falcon, Prairie Falcon, Rough-legged Buzzard, Common Buzzard, Griffon Vulture,31

Common Kestrel

Synovitis, air sacculitis, catarrhal tracheitis,serofibrinous pneumonia, sitting on paralyzed hocks,

One type Phasianinae See text

Several types ? Black-headed Gull, Brown-eared Bulbul, Phasianinae,White-fronted Goose

Asymptomatic

CHAPTER 38 MYCOPLASMA AND RICKETTSIA

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probably other strains of avian mycoplasmatales)has a special organelle for attaching to the host cell.

Depending on the virulence of the strain in question,cellular damage may be caused at the site of coloni-zation. The host reacts with a serofibrinous inflam-mation and activation of the cell-mediated defensesystem. The excessive response of the latter (which isgenetically determined) governs the type and magni-tude of pathologic changes.

Many mycoplasmatales cause transformation of thehost lymphoblasts (mainly T-cells) by excreting amutagenic substance. Affected cells function improp-erly and there is a severe proliferation of immaturelymphocytes in local lymph follicles with invasion ofthe lymphoid cells into the infected area. These al-tered lymph follicles can appear similar to thosedescribed for lymphoma. Other pathogenicity factorsare cytotoxins (exotoxins, H2O2) and polysaccharides.Triggering factors for mycoplasmatales are imma-ture epithelial membranes, environmentally induceddyspnea (heat, dry air) and damage to the respira-tory epithelium (excess NH3, paramyxovirus, re-ovirus, adenovirus, infectious bronchitis virus and E.coli). The involvement of several different factors ina flock outbreak creates a high variability in clinicaland pathologic changes.

Incubation PeriodIncubation periods for M. gallisepticum are 6-21 daysin chickens and 7-10 days in turkeys.43 In other avianspecies and with other mycoplasmatales, long la-tency periods, egg transmission and the involvementof environmental factors make the determination ofan incubation period difficult.

Clinical Disease and Pathology

Red JunglefowlU. gallorale has been isolated from the pharynx ofthis species. Although the strain is serologically iden-tical with isolates from the chicken, experimentalinfections did not cause disease in chickens.

PhasianinaeThe Common Pheasant and its subspecies, Crossop-tilon spp., the Golden Pheasant and probably otherpheasants are susceptible. The main host is the Com-mon Pheasant, which is typically maintained in largeflocks. The strains of mycoplasma that are infectiousto pheasants have been incompletely studied anddocumentations in the literature provide conflictinginformation.13 The author’s experience suggests that

the Phasianinae have host-adapted strains, one thatis apathogenic and another that experimentally re-produces typically defined signs of the disease. Clini-cal signs are most common in large groups of chicksat the age of two to eight weeks. Adults are rarelyaffected. A seasonal peak can be observed betweenJune and August. The disease spreads slowly and notall aviaries are always affected. Morbidity is high.Mortality depends on secondary factors and canrange from 30 to 90%. Blinking the eyes and scratch-ing at the eyelids are the first clinical signs. Deterio-ration of the general condition, photophobia andswelling of the eyelids are followed by exudation,blepharoconjunctivitis and sometimes keratitis; ap-proximately 25% of the corneal surface is affected.Death can be caused by cachexia as a result of blind-ness. Voluminous expansion of the infraorbital sinus,which contains only a small amount of exudate, maybe observed. Birds are frequently dyspneic, particu-larly when agitated. At postmortem, the air sacs maybe mildly inflamed or grossly normal.13

Japanese QuailExperimental infections indicate that JapaneseQuail are less susceptible to M. gallisepticum thanare chickens. Isolation of the organism is possiblefrom the trachea, lung and brain for weeks post-in-fection. The course is subclinical. Infections derivedfrom contact with infected chickens or egg transmis-sion have been documented.

Bobwhite QuailThis species is raised in large numbers in the southernparts of the United States. Dyspnea and anorexia havebeen observed. An isolate assumed to be M. gallisep-ticum from Bobwhite Quail caused typical lesions inturkey poults. However, a strain of M. gallisepticumexperimentally given to Bobwhite Quail chicks did notresult in lesions, and no antibodies were produced.

CLINICAL APPL ICAT IONSMycoplasmatales are most important in dense populationsof birds where direct transmission can easily occur. Controlcan be enhanced through sound hygiene.

Mycoplasma are resistant to antibiotics that inhibit cell walldevelopment (eg, penicillins, cephalosporins, bacitracin andsulfonamides).

Mycoplasmatales that are free in the environment are sus-ceptible to all commonly used disinfectants. Organismswithin host excretions are protected from contact with thedisinfectant. Secretions and excretions must be removedbefore disinfecting procedures are effective.

With intensified aviculture, increased farm sizes and popula-tion densities on these farms, more problems with mycoplas-matales are to be expected.


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PartridgeIt is assumed that the same strains as in Phasiani-nae cause disease in the partridge, although this hasnot been proven. Affected birds develop infectionsthat are similar to those seen in pheasants, but thereis no defined seasonal peak. Birds up to 11 weeks ofage show a swelling of the infraorbital sinuses which,in contrast to pheasants, are filled with a fibrinous,cheesy exudate (Figure 38.2). Free-ranging Red-legged Partridge usually develop clinical disease inAugust to December. Isolates are assumed to be iden-tical to strains removed from pheasants and otherpartridges.

Rock PartridgeDisease has been described only in chicks and not inthe respective breeding flock. Emaciation and swol-len sinuses are the main clinical signs. Isolates as-sumed to be M. gallisepticum were experimentallyapathogenic for chickens and turkeys.

PeafowlAffected birds are lethargic, shake their heads toremove sticky nasal exudates, have swollen infraor-bital sinuses and make gurgling respiratory sounds.Latent infections are thought to occur.

GuineafowlAn outbreak of infectious synovitis caused by M.synoviae in guineafowl could not be distinguishedclinically or pathologically from the lesions that oc-cur in chickens and turkeys.26 In contrast to chickensand turkeys, affected guineafowl (including experi-mentally infected birds) developed severe amyloi-dosis and did not develop sinusitis.29 Strains isolatedfrom guineafowl are more virulent for guineafowlthan for chickens.

Domestic DuckA variety of Mycoplasma and Acholeplasma strainscan be isolated from domestic ducks. In the few iso-lates that have been evaluated experimentally,pathogenicity is limited to mild respiratory lesions,conjunctivitis and cloacitis. M. anatis may cause en-zootic rhinitis, sinusitis, conjunctivitis and lacrima-tion in association with concommitant influenzavirusA infections. Morbidity may be as high as 50-80%, butmortality remains below 5%. As a rule, affected ducksrecover spontaneously without therapy. Experimen-tally, the clinical disease can be produced using M.anatis and influenzavirus A,32 although both infec-tious agents alone have been proven to be apatho-genic. The role of M. cloacale as the cause of a cloaci-tis in ducks has not been fully studied. Most of theMycoplasma and Acholeplasma strains described arecapable of causing increased embryonic mortality.

Domestic GooseGeese suffering from cloacitis and necrosis of thephallus were found to be infected with mixed culturesof M. spp. (mostly M. cloacale, but also M. anseris andstrain 1220). Phallus lesions are characterized byserofibrinous inflammation of the mucous membraneof the lymph sinus, the glandular part of the phallus,and occasionally the cloaca and the peritoneum. Ne-crosis of the affected phallus can be severe if secon-dary pathogens are present. Mortality is less than1%. M. spp. can be isolated from the phallic lymphsecretion as well as the spleen, testes, air sacs, peri-toneum and liver. The incidence of affected gandersin some flocks can be as high as 40-100%. Highnumbers of infertile eggs and a high incidence ofembryonic death are common in affected flocks.36,40

A. axanthum may cause embryonic mortality (up to60%) around the 13th day of incubation. The organ-ism can be isolated from the respiratory tract andfeces of breeding birds showing embryonic mortality.Infected adults develop fibrinous peritonitis, salpin-gitis and air sacculitis. Goslings from infected flocksand experimentally infected neonates can suffer from

FIG 38.2 A young partridge was presented with a five-day historyof progressive ocular irritation, photophobia, dyspnea and periocu-lar swelling. Large, bilateral, periocular masses were noted onphysical examination. Large quantities of necrotic debris weresurgically removed from both intraorbital sinuses. Mycoplasma sp.was isolated from culture samples taken from the sinus cavities.The bird responded to postsurgical therapy with tylosin (courtesyof Helga Gerlach).


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mild to severe air sacculitis (depending on the viru-lence of the strain in question).37 The pathogenicityof A. axanthum can be potentiated by concommitantinfection with parvovirus, even if antibody titers arehigh enough to prevent the parvovirus from causingclinical disease.22

Domestic PigeonAt least six different species of mycoplasmataleshave been isolated from domestic pigeons.13,15 All ofthem apparently are incapable of causing primarydisease. Following natural or experimental infectionwith all except M. columbinum and A. laidlawii(which have not been tested), the agents can be iso-lated from various organs including brain, eye andjoints of asymptomatic birds. The frequent coloniza-tion of the pharyngeal mucosa is epizootiologicallyimportant because pigeons feed their offspring cropmilk. During the act of regurgitation the crop milkpasses over the infected mucosa and may be contami-nated. Although egg transmission has been proven,this means of transmission might play the most im-portant role.

Clinical signs of rhinitis, sinusitis, tracheitis andconjunctivitis are generally chronic in nature andvary with secondary factors such as concomittantinfections with Salmonella spp. or Chlamydia psit-taci. Under these conditions, mycoplasmatales canbe isolated from the lower third of the trachea, airsacs and occasionally lung, and birds frequently havepersistent respiratory sounds and serofibrinous in-flammation of these organs. The association betweenthe colonization of the meninges and synovial struc-tures by mycoplasmatales and the frequency of ar-thritis and meningoencephalitis caused by salmonel-losis has not been determined. Further evidence forthe apathogenicity of uncomplicated mycoplasmatalinfections in pigeons is the fact that humoral anti-bodies only occasionally develop following natural orexperimental infections. In contrast to some olderreports, experimental infection of chickens with pi-geon mycoplasma strains does not lead to clinicaldisease. M. gallisepticum does not affect pigeons. M.columborale was recovered from a pigeon flock withrespiratory signs that responded to treatment withtylosin. Experimentally infected three-week-oldchicks developed mild to severe air sacculitis, butwere clinically asymptomatic. There is no report ofnatural infection in chickens with M. columorale.25

Other PigeonsIn addition to domestic pigeons, infections with my-coplasmatales have also been described in the Wood

Pigeon,20 Collared Dove14 and Crowned Pigeon.13 Theisolation of M. columbinum and M. columborale hasalso been recovered from healthy “feral pigeons.”21

Saker FalconMycoplasma was isolated from the trachea of a SakerFalcon with insufflation of the soft tissues around theeye and between the rami mandibulares followingeach expiration. Similar respiratory signs occured intwo contact birds.10 A definitive connection betweenthe clinical signs and the M. isolate was not estab-lished. Mycoplasma-induced synovitis has also beendescribed in this species.19

Peregrine FalconA Mycoplasma sp. was isolated from the trachea oftwo Peregrine Falcons with anorexia, vomiting, res-piratory sounds and tachypnea (60-70 beats per min-ute).10 The animals responded to treatment with ty-losin.

Budgerigar A Mycoplasma sp. was isolated from a budgerigarwith air sacculitis.1 The serum of five contact birdsrevealed humoral antibodies against the homologousstrain with titers between 1:160 and 1:640. Antibod-ies were not detected against M. gallisepticum andM. meleagridis. The budgerigar strain propagated inthe embryonated chicken egg and showed no em-bryonal pathogenicity. Budgerigars experimentallyinfected with M. gallisepticum6 and M. synoviae3 de-veloped clinical signs. Budgerigar are not consideredto be a natural host of M. gallisepticum and M.synoviae.

CockatielIt has been assumed that conjunctivitis in cockatielscan be caused by mycoplasmatales, (see Color 26) aswet sneezes and sinusitis are common in those birds.Although mycoplasmatales can be isolated fromsome of these cases, their importance in the diseaseprocess has not been determined. From the clinicalcourse and response to treatment it can be concludedthat chlamydiosis and infections with polyomavirusare the main pathogens in these conditions.9,12 Manycockatiels in Florida with symptoms of mycoplas-mosis respond to tylosin (as an eyewash) or lincocin-spectinomycin (Harrison GJ, unpublished).

Severe MacawAn epornitic of mycoplasma was described in SevereMacaws with clinical and pathologic lesions in therespiratory tract. Although mycoplasmas were iso-


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lated, no causal relationship between the agent andthe disease could be established.11

Yellow-crowned AmazonA flock of Yellow-crowned Amazon Parrots experi-enced high mortality (200 of 1100 birds) with anupper respiratory tract disease. Lesions were compli-cated by the presence of many bacteria and also somefungi. A mycoplasma strain (assumed to be M. gal-lisepticum) was isolated and used for experimentalinfections in budgerigars and chickens. Mild air saclesions were induced in budgerigars, but the strainwas apathogenic for chickens. The serologic evidencefor M. gallisepticum was questionable.3

CockatooAn unidentified strain of mycoplasma was isolatedfrom a group of cockatoos with severe air sacculitis.12

CanaryAn unidentified strain of mycoplasma was recoveredfrom a flock of canaries with a high incidence ofwheezing and “tail-bobbing.” The affected flock hadsuffered from canarypox.

Pathology

At necropsy, lesions caused by various mycoplas-matales in respective hosts vary in degree but not inpresentation. Serous to serofibrinous conjunctivitis,rhinitis, sinusitis, tracheitis, air sacculitis and focalbronchopneumonia have all been described.18 Thenasal cavity and the infraorbital sinus frequentlydisplay a unilateral, seromucoid (later fibrinous)exudate that also fills the choanal fissure. In ducksand turkeys, the exudate is often semigelatinous,fibrinous or caseous, and leads to distension of theinfraorbital sinus. The mucous membranes are swol-len and may show petechiation. The tenaciousexudate can be mixed with fibrinous debris.

Histopathologically, the disease is initially charac-terized by severe distension of the mucous glands,the swollen cells of which have particularly largenuclei. Subsequent proliferation of epithelial cellsleads to multilayered, glandular epithelium, pres-sure on the glands themselves and mucoid degenera-tion. The superficial mucosal epithelia lose theircilia, proliferate to 10-15 cellular layers and finallyshow vacuolization and karyorrhexis. In contrast tovarious viral diseases, desquamation and necrosis ofthe epithelium are mild.

From the second week after infection, an infiltrationof the lamina propria with lymphocytes and histio-cytes is seen. Round-to-oval, up to 400 µm in diame-ter nodules that consist mainly of lymphocytes ap-pear in the submucosa. The proliferation of thelymph follicles persists in the lower part of the tra-chea and the syrinx from the 2nd to the 12th weekafter infection. This is generally a longer course ofreactions than are noted with other infections of therespiratory tract. As healing occurs, there is a prolif-eration of connective tissue in the submucosa. Thehistopathomorphologic changes vary depending onthe presence of secondary bacterial or viral infec-tions. Secondary fungal infections are rather rare.

Lesions of the air sacs start with edema between theinner and outer epithelial layers. The cellular reac-tion consists initially of subepithelial infiltrates ofheterophils. The capillaries are engorged. Progres-sion is marked by increased edema and growingnumbers of heterophils followed by lymphocytes,macrophages and plasma cells. The normally flatepithelium becomes cubic, loses its cilia and is finallydesquamated. Inflammatory exudate, mainly in theform of fibrin, appears on both sides of the air sacmembrane. The host responds with proliferation ofthe endodermal epithelial layer and necrotic foci ofepithelial cells and proliferation of fibrocytes andmononuclear cells (80% can be lymphocytes). Agranulomatous reaction characterized by the forma-tion of multinucleated giant cells occurs by the thirdweek post-infection. Lymph follicles and diffusemononuclear infiltrations govern the pathologic pic-ture. Pneumonia is a rare complication in avian my-coplasmosis, and in most instances is caused by sec-ondary infections with E. coli.

Mycoplasma colonization of the mucosa of the uro-genital tract can cause pathologic lesions, althoughcolonization of the phallus may be inconspicuous.Histologically, lesions are mainly seen in the part ofthe mucosa where the majority of the glands aresituated (species-specific differences). Submucosalproliferation of lymph follicles and disseminated in-filtration of lymphocytes into the tissue are the mainlesions. In ganders, the phallus is enlarged and cov-ered with fibrinous exudate, and may finally becomenecrotic if secondary infections occur. Synovial mem-brane lesions have been rarely reported in compan-ion and aviary birds.23


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Differential Diagnosis

The rule-out list includes many viral, bacterial andfungal diseases. In Psittaciformes, pigeons, ducksand geese, chlamydiosis is the main rule-out. Thegenital tract can be infected by other microorganismsas well. Embryopathologic lesions and embryonaldeath are suggestive. With mycoplasmatales, in-fected embryos generally die late in incubation. Em-broyos that die after pipping frequently have airsacculitis of the left thoracic air sac group (exceptionsare geese that have air sacculitis bilaterally). Afterhatching, chronic lymphofollicular proliferation canbe so severe that lymphoma must be considered inthe rule-out list.

Diagnosis

A tentative diagnosis can be made by histopathologicexamination. Isolation of the agent is necessary foridentification and biologic assays. Because of thefastidious nature of this organism and the difficultiesin identifying the agent, specialized laboratories arenecessary to isolate Mycoplasma. In addition, themycoplasmatales need to be differentiated from bac-terial L-forms. Swabs from the upper respiratorytract, or the phallus in males, can be taken from livebirds. Endoscopic biopsies of affected air sacs areuseful diagnostic aids. Samples from air sacs, sal-pinx, lungs and spleen should be collected for post-mortem evaluation. Transport media are necessaryfor shipping samples. They should contain heart in-fusion broth, mycoplasma broth or another similarmedium with penicillin (2000 IU/ml). Because theorganisms are primarily to be protected from dryingduring transport and are not supposed to grow, thepH is not of particular importance (however, it shouldbe around 7 or slightly above). Penicillin does notaffect acholeplasma, but thallium-acetate does;therefore, mycoplasma broth that contains thallium-acetate should not be used for material from geese.

Indirect diagnosis of mycoplasmatales by serology ishampered by false-positive (cross-reactions) andfalse-negative tests. The presence of a mycoplas-matales on a mucosal surface usually does not stimu-late production of humoral antibodies. Antigen-rec-ognizing cells become active only after the mucosahas been penetrated. The most frequently used testsare: serum slide agglutination (SSA), HI test (for thehemagglutinating species), growth- or metabolic-in-hibition test, immunodiffusion test, immunofluores-cence test, ELISA, and recently the polymerase chainreaction.

Treatment

Clinical infections can be treated with tylosin, spi-ramycin and erythromycin or spectinomycin in com-bination with clindamycin or pleuromutilin. The effi-cacy of tetracyclines against avian mycoplasmataleshas yet to be proven. However, in Psittaciformes, thetetracyclines are recommended because of the clini-cal similarities between mycoplasmosis and chlamy-diosis. The pigeon strains are highly resistant toerythromycin and, to a lesser extent, tylosin. Onlypleuromutilin was able to inhibit 57 of 65 strainsrecovered from pigeons.15 The LD50 for pleuromutilinin pigeons is 440 mg/kg, considerably less than forchickens and turkeys. This drug must be carefullyused when treating pigeons that are feeding off-spring or if used in the water during hot weather.

Spiramycin given parenterally to Ploceidae,Estrildae and even canaries may lead to suddendeath from unknown causes. The same dose given viadrinking water is well tolerated. Spiramycin is one ofthe macrolid antibiotics and is given at a dose of 100mg/kg body weight IM, or 100-200 mg/kg body weightorally. Since the primary patent has expired, severalmanufacturers produce it. Enrofloxacin has beenused to treat mycoplasmosis in poultry. There havebeen no reports of success in treating mycoplasmosiswith enrofloxacin in other birds. Treatment is de-signed to allow clinically affected birds to recover.The organism is difficult to eliminate.

Control

Recovery from mycoplasmosis results in the produc-tion of a low antibody level and a persistent infection.In vitro, mycoplasmatales can propagate in the pres-ence of homologous antibodies, indicating that hu-moral antibodies are not correlated with immunity.The disease is governed by the excessive reactions ofthe cell-mediated immune system. Therefore, vacci-nations might sensitize a bird to the organism andcause a severe reaction to a field exposure. Theoreti-cally, vaccines that prevent mycoplasmatales fromattaching to the mucus cells might give protection.


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Rickettsia

Little information is available on rickettsial infec-tions in birds. Rickettsia form a group of microorgan-isms, the taxonomy of which has still not been fullydetermined. They are obligatory cellular parasites,and can be differentiated from chlamydia by theabsence of a developmental cycle and the capacity tosynthesize energy-rich compounds (ATP). Rickettsiaare small rods or coccoids with an average size of 0.3to 0.5 µm in diameter and 0.8 to 2.0 µm in length.They may also be pleomorphic, and are generallynonmotile. Multiplication takes place by binary fis-sion. The organism parasitizes reticuloendothelialcells, vascular endothelial cells or erythrocytes. In-fections may occur in arthropods, which can serve asvectors or as primary hosts. The mutualistic forms ininsects are considered to be essential for develop-ment and reproduction of the host.47 Rickettsia maycause disease in humans, many vertebrates and in-sects. The organisms can be cultured in embryonatedchicken eggs or metazoan cells. The chlamydialstaining procedures may be used, although somechanges are made particularly in fixation.

The rickettsia have historically been divided intothree families:47 Rickettsiaceae, Bartonellaceae andAnaplasmataceae. The latter two families are nolonger considered to be Rickettsia and are “phylo-genically unaffiliated bacteria” (Gothe, unpub-lished).

Rocky Mountain Spotted Fever (RMSF)

RMSF is a mammalian disease caused by Rickettsiarickettsii. Prior to the availability of antibiotics(tetracyclines), infections were characterized by highmortality rates. The pathogenicity (for many otherrickettsia as well) involves a toxin-like action thatdamages endothelial cells, inducing increased capil-lary permeability, plasma flow into the tissue, hemo-concentration and eventual circulatory collapse. Cy-totoxicity is thought to be the mechanism by whichrickettsia gain access to the cytoplasm of the hostcells.41

RMSF is transmitted by ticks (mainly Dermacentorspp.) and rodents. Dogs and opossum are thought tobe reservoirs. Birds, including chickens, several Co-lumbiformes, pheasants, Falconiformes and the

Magpie, are susceptible to experimental infectionand may also serve as reservoirs. Pigeons may beparticularly important reservoirs.24 Clinical abnor-malities have not been described in infected birds.

Q-Fever

Q-fever, caused by Coxiella burnetti, is an aerosol-borne disease in humans with worldwide distribu-tion. Direct and indirect transmission (arthropods)can occur in humans and other host species. Thisagent differs considerably from other members of theRickettsiaceae. The cells are smaller (0.2 to 0.4 µmby 0.4 to 1.0 µm) and replicate in vacuoles of the hostcells. In contrast to chlamydia, C. burnetti infectionsresult in the formation of phagolysosomes. Two dif-ferent phase variations are distinguished. Phase I ismore virulent, presumably because of the highamount of lipopolysaccharides, and occurs naturally.Phase II appears following repeated passages in theyolk sac of embryonated chickens. The occurrence ofplasmids has been established. The replication cycleincludes production of endospore-like bodies, whichare probably the cause of the high tenacity of C.burnetti. This organism is resistant to chemical dis-infectants and high temperatures that might killother rickettsia. Metabolic activity in these en-dospore-like bodies is extremely low outside the host.Survival, which can be several years in the feces ofticks, is augmented by dryness. Chlorine-containingpreparations are recommended for disinfection. Theeffects of phenol and formaldehyde products are de-batable.35

The host spectrum of C. burnetti is wide, and includesarthropods (particularly ticks), birds and mammals.An infectious cycle in free-ranging animals, arthro-pods, birds and mammals is differentiated from aninfectious cycle in domesticated animals (sheep,goats, cattle, dogs). Cross transmission, including inman, is possible from both cycles.35

Avian susceptibility to C. burnetti seems to be high,and this organism has been demonstrated in at least49 avian species.8,30,39 The type of feeding behavior,breeding areas and seasonal migrations are the mainfactors for the spread of the organism. Carrion-eatingbirds may be infected by ingesting the infected pla-centa of ruminants from endemic areas. The feedinglocation of granivores and insectivores is more impor-tant than their food preferences. Birds that live inclose contact with humans (synanthrops) are exposedmore frequently to C. burnetti than birds that avoidcivilization (exanthrops). The susceptibility to infec-


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tion in domesticated birds is variable.8,38,39 Chickensare apparently most susceptible and may shed theagent in the feces for 7 to 40 days post-infection.Vertical transmission via the egg may also occur.Domesticated pigeons are next in susceptibility. Do-mesticated turkeys, ducks and geese are rarely in-fected. All the free-ranging urban pigeons that wereexamined (125) in the Netherlands showed CF titersof ≤1:20 against C. burnetti.7 In experimentally in-fected domesticated pigeons, the agent could be iso-lated from the spleen and lung 58 days post-infec-tion.34 No clinical disease has been observed in anysusceptible avian species.

C. burnetti can be identified in tissues using variousstaining methods (Giemsa, Macchiavello, Cas-tañeda) as for chlamydia. The organism replicates inembryonated chicken eggs and various cell cultures.Antibodies can be demonstrated using the CF test orthe ELISA. Not all exposed domestic pigeons havebeen found to produce CF antibodies.34 Antibody pro-duction is not related to immunity.

Therapy with tetracyclines is effective for the clinicaldisease, but elimination of the organism is not possi-ble. Treatment for infected birds is not encouragedbecause of the high immunosuppressive side effect ofthe tetracyclines.

Aegyptianella (Ae.)

Ae. pullorum is the causative agent of anemia andhepatitis in chickens and other birds. It is an erythro-cytic parasite that produces endocytoplasmic inclu-sions, which stain using the Giemsa or Pappenheimprocedures. The inclusions measure 0.3 by 4.0 µm,and each can contain up to 26 initial bodies (repro-ducing form up to 0.8 µm in diameter). In manyinstances, the inclusions are polymorphic (round,oval, ring- or horseshoe-shaped) and are separatedfrom the plasma by a single-layered membrane. In-fection of the cell starts with an endocytosis-likeprocess followed by vesiculation in the erythrocyte.Exocytosis is one way in which the organism is re-leased from the infected host cell. However, the hosterythrocytes are usually damaged by the parasite,leading to lysis and release of the parasite into theplasma. Arthropods, mainly ticks of the genus Ar-gas,16 are essential for transmission.

The organism is most common in tropical and sub-tropical regions including the Mediterranean. Thehost spectrum is probably incomplete but certainlyincludes chickens, quail, Columbiformes, Strigidae,

Falconidae, Accipitridae, crows, canaries, ostriches,ducks, geese and some Psittaciformes such asAgapornis spp.16,17,33

Clinical signs in young birds are characterized by anacute onset of anemia, anorexia, weakness, weightloss, greenish diarrhea and death.33 Chronic infec-tions in older birds are characterized by icterus,which may not be clinically recognizable. The post-

FIG 38.3 a) A group of Gouldian Finches died following an onsetof clinical signs that included depression, dyspnea and coughing.Electron microscopy of tracheal epithelial cells revealed intracyto-plasmic inclusion bodies with organisms morphologically sugges-tive of rickettsia. Magnification: x 22,250. b) Same cell type withrickettsia in binary fission; magnification: x 52,500; (courtesy E.Göbel).


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mortem examination reveals anemia as well as aconsiderable enlargement of the liver and spleen.

Small inclusion bodies were demonstrated in themajority of the mature erythrocytes in two domesti-cally raised Eclectus Parrot neonates (six weeks old)with heterophilia (toxic heterophils) and anemia.These intracellular parasites resembled those identi-fied as Aegyptianella in several imported AfricanGrey Parrots. Following a long-term course of doxy-cycline therapy, the parasites were no longer identi-fiable in the erythrocytes. The parents that producedthese neonates were also positive for aegyptianellaand responded to long-term doxycycline therapy.33

In gallinaceous birds, the age and condition of the hostgovern the pathogenesis and outcome of the infection.Up to 60% of the erythrocytes may be infected inone-day-old chicks, while by one year of age less than1% of the erythrocytes may be infected.16 Mortality ishigher in chicks less than two days of age.

The rule-out list includes internal bleeding, chlamy-diosis and chronic diseases of various etiology. Fordiagnosis, a blood smear stained according to Giemsaor Pappenheim shows the parasites in the erythro-cytes. Tetracyclines are effective for treatment. Tickcontrol is mandatory to prevent reinfection and epi-zootics.

Unclassified

There are indications that diseases caused by rick-ettsia other than aegyptianella may occur. Trachealepithelial cells in Gouldian Finches with severe res-piratory disease were filled with cytoplasmic “inclu-sions.” Many of the affected birds died. Electron mi-croscopy of the epithelial cells revealed particles thatwere morphologically compatible with rickettsia(Figure 38.3). Treatment with tetracyclines was suc-cessful. Isolation was not possible because the mate-rial had been prepared for histopathology.27

References and Suggested Reading

1.Adler HE: Isolation of a pleuropneu-monia-like organism from the air sacof a parakeet. J Am Vet Med Assoc130:408-409, 1957.

2.Bencina D, et al: Mycoplasma speciesisolated from six avian species. AvianPathol 16:653-664, 1987.

3.Bozeman LH, et al: Mycoplasma chal-lenge studies in budgerigars (Melop-sittcus undulatus) and chickens.Avian Pathol 13:426-434, 1984.

4.Bradbury JM, et al: Isolation of Myco-plasma cloacale from a number of dif-ferent avian hosts in Great Britainand France. Avian Pathol 16:183-186,1987.

5.Buntz B, et al: Isolation of Myco-plasma gallisepticum from geese.Avian Pathol 15:615-617, 1986.

6.Butcher GD, et al: Reductions of clini-cal signs in budgerigars experimen-tally infected with Mycoplasma gal-lisepticum. J Assoc Avian Vet 4:227-230, 1990.

7.Dorrestein GM, et al: Some observa-tions on the health status of free-liv-ing urban pigeons (Columba livia).VII DVG-Tagung Vogelkrht,München, 1990, pp 77-87.

8.Enright JB, et al: Q-fever antibodies inbirds. J Wildlife Dis 7:14-21, 1971.

9.Fudge AM: Common medical condi-tions seen in the cockatiel Nym-phicus hollandicus. Conf Europ ComAssoc Avian Vet, Vienna, 1991, pp351-356.

10.Furr PM, et al: Isolation of mycoplas-mas from three falcons (Falco spp.).Vet Rec 100:72-73, 1977.

11.Gaskin JM, et al: A mycoplasma-asso-ciated epornitic in severe macaws(Ara severa severa). Proc Am AssocZoo Vets pp. 59-61, 1979.

12.Gaskin JM: Mycoplasmosis in cagedbirds. Proc 1st Intl Conf Zoological &Avian Med, 1987, pp 57-60.

13.Gerlach H: Infektionen durch Molli-cutes bei Vögeln. In Gylstorff I: Infec-tionen durch Mycoplasmatales. Infek-tionskrankheiten und ihre Erreger.Band 21 VEB Verlag Gustav Fischer,Jena, 1985, pp 448-491.

14.Gerlach H: Über das Vorkommen vonMycoplasmen bei Tauben. BerlMünch Tierärztl Wschr 90:140-143,1977.

15.Gerlach H: Untersuchungen überMollicutes bei Tauben. Habil.-SchriftTierärztliche Fakultät München,1978.

16.Gothe R, et al: Genus II. Aegyptian-ella (Carpano, 1929). In Bergey’sManual of Systematic Bacteriologyvol 2. Baltimore, London, Los Ange-les, Sydney, Williams and Wilkins,1986, pp 722-723.

17.Gylstorff I: Rickettsiales. In GylstorffI, Grimm F: Vogelkrankheiten.Stuttgart, Verlag Eugen Ulmer,1987, p 317.

18.Gylstorff I: Entzündung der Naseund Nebenhöhlen. In Joest: Hand-buch der speziellen pathologischenAnatomie der Haustiere. Band 7, 3.Auflage Berlin und HamburgVerlagPaul Parey.

19.Gylstorff I: Mollicutes. In Gylstorff I,Grimm F: Vogelkrankheiten.Stuttgart, Verlag Eugen Ulmer,1987, pp 323-325.

20.Jordan FTW, et al: The recovery ofMycoplasma fron the avian oesopha-gus. Vet Rec 102:403-404, 1978.

21.Jordan FTW, et al: The isolation ofMycoplasma columbinum and M. co-lumborale from feral pigeons. Vet Rec109:450, 1981.

22.Kisary J, et al: Mycoplasma infectionin geese II. Studies on pathogenicityof mycoplasmas in goslings and gooseand chicken embryos. Avian Pathol5:15-20, 1976.

23.Kleven SH, et al: Mycoplasma syno-ciae infection. In Calnek BW, et al(eds): Diseases of Poultry 9th ed.Wolfe Publishing, 1991, pp 223-231.

24.Lundgren DL, et al: Infectious dis-eases of wild animals in Utah. VI. Ex-perimental infection of birds withRickettsia rickettsii. J Bact 91:963,1966.

25.Macowan KJ, et al: Mycoplasma co-lumborale in a respiratory conditionand experimental airsacculitis ofchickens. Vet Rec 109:562, 1981.

26.Maestrini N, et al: Amiloidosi nellagallina faraona. Veterinaria 24:378-380, 1967.

27.Merkus: Dr. med vet., Landesunter-suchungsamt, Krefeld Germany

28.Müllegger P-H: Die Differenzierungaviärer Mycoplasmen mit Hilfe derDisk-Elektrophorese. Zbl Bakt Hyg IAbt Orig A 226:119-136, 1974.

29.Pascucci S, et al: Mycoplasmasynoviae in the Guinea-Fowl. AvianPathol 5:291-297, 1976.

30.Pospisil R, et al: Q-Fieber in derOstslowakei (CSSR). 2. Internat. Ar-beits-koll. Naturherde von Infek-tionskrankheiten in Zentraleuropa.Graz, 1976.

31.Poveda JB, et al: An epizootiologicalstudy of avian mycoplasmas in south-ern Spain. Avian Pathol 19:627-633,1990.

32.Roberts DH: The isolation of an influ-enza A virus and a mycoplasma asso-ciated with duck sinusitis. Vet Rec76:470-473, 1964.

33.Rosskopf WJ, et al: Ageyptiennella-like parasites seen in eclectus parrotsin southern California. Proc AssocAvian Vet, 1992, pp 129-133.

34.Schmatz H-D, et al: Zum Verhaltendes Q-Fieber-Erregers, Coxiellaburnetii, in Vögeln. 2.Mitteilung: Ex-

perimentelle Infektion von Tauben.Dtsch Tierärztl Wschr 84:60-63, 1977.

35.Selbitz H-J: Lehrbuch der veterinär-medizinischen Bakteriologie. GustavFischer Verlag Jena, Stuttgart pp.257-268, 1992.

36.Stipkovits L, et al: Mycoplasma infec-tion of geese. I. Incidence of Myco-plasma and Acholeplasma in geese.Avian Pathol 4: 35-43, 1975.

37.Stipkovits L, et al: Occurrence of myco-plasmas in geese affected with in-flammation of the cloaca and phallus.Avian Pathol 15:289-299, 1986.

38.Syrucek L, et al: Q-fever in domesticand wild birds. Bull Wld Hlth Org15:329-337, 1956.

39.Thiel N, et al: Übersichtsreferat: ZumVerhalten des Q-Fiebererregers,Coxiella burnetii, in Vögeln. 1. Mittei-lung: Natürliche Infektionen beiVögeln. Dtsch Tierärztl Wschr 84:63-68, 1977.

40.Varga ZS, et al: Biochemical and sero-logical study of two Mycoplasmastrains isolated fromn geese. Arch ex-per Vet med Leipzig 43:733-736, 1989.

41.Weiss E, et al: Order I Rickettsiales(Gieszczkiewicz, 1939). In: Bergey’sManual of Systematic BacteriologyVol 2. Baltimore, London, Los Ange-les, Sydney, Williams and Wilkins,1986, pp 687-722.

42.Yamamoto R: Mycoplasma melea-gridis infection. In Calnek BW, et al(eds): Diseases of Poultry 9th ed.Wolfe Publishing, 1991, pp 212-223.

43.Yoder H Jr: Mycoplasma gallisep-ticum infection. In Calnek BW, et al(eds): Diseases of Poultry 9th ed.Wolfe Publishing, 1991, pp 198-212.


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