G.J. UNDERWOOD,
C. BUTTER,
T.F. DAVISON
Institute for Animal Health,
Compton, Newbury, Berkshire,
UK
Natural infection with Marek's disease virus (MDV) occurs after inhalation of infective fragments from feathers and sloughed skin flakes (Beasley et al., 1970). Adldinger & Calnek (1973) suggested that MDV could be taken up by lung phagocytic cells and transported to the lymphoid organs. However, the actual mechanism of viral uptake and transportation remains unknown, as most studies of MD pathogenesis have involved inoculating chickens with cell-associated MDV by a parenteral route.
Studies on natural infection with MDV have relied on generating an airborne suspension of infective dust in an enclosed space (Beasley et al., 1970; Calnek et al., 1970; Jurajda & Klimes, 1970; Adldinger & Calnek, 1973) or direct contact exposure with infected individuals (Gildow et al., 1940; Sevoian et al., 1963; Schmittle & Eidson, 1968; Kenzy & Cho, 1969) and did not aim to elucidate early events after infection. These methods lack reproducibility as MD clinical signs were not always detected (Jurajda & Klimes, 1970). Furthermore, timing of infection could not easily by standardised. Calnek et al. (1970) demonstrated infection with cell-free MDV by the intra-tracheal route but details were not provided. Davidson and Borenshtain (2004) extracted cell-free MDV from sonicated feather-shaft tips that were 'dripped' into the beak, though this extract did not cause clinical MD in all birds. Here, we describe a method to infect individual chickens with MDV by the natural route using a standard dose of MDV in its 'native' state in infective dust.
Specified-pathogen free Rhode Island Red (RIR) chicks were obtained from the Institute Animal Health (IAH). Experimental birds were housed in wire cages in isolation rooms provided with high efficiency particle air (HEPA)-filtered air under positive pressure. Rooms were hosed daily to prevent build-up of dust on the floor and walls. Food and water were available ad libitum and experiments met with UK Home Office ethical guidelines.
The characteristics of fractionated MDV infective dust samples were studied using an Andersen filter. Infective dust was obtained by inoculation of week-old chicks with 1,000 plaque forming units (pfu) MDV, strain HPRS16. A similar group of uninfected controls was housed in a separate room. Three weeks later, an Andersen impact air sampler (Graseby Andersen, Orpington, UK) and air compressor were placed in each room (for details see Underwood, 2003). The sampler had filters to separate the dust into various particle diameters (9.0, 5.8, 4.7, 3.3, 2.1, 1.1, 0.65 and 0.43μm) representing the size of particles that could be deposited in the airways (Hayter and Besch, 1974). The flow rate was 28.3 L/min and air turnover through the room was minimised to increase airborne particulates. The collection period was 3wk. Dust and dander also were collected from the room extract filters, coarsely filtered (300μm-steel mesh) and desiccated.
Most particles were present in the 9.0, 5.8 and 4.7 μm fractions and smaller-sized fractions yielded less, with the 0.65 μm fraction being barely visible. Light microscopy revealed the fractions ranging from 1.1 to 9.0 μm consisted of a mixture of feather barbules and intact or fragmented squamous epithelial cells (Figure 1). Particle dimensions varied in the fractions as aerodynamic properties are determined by the smallest two dimensions and it was not uncommon to see particles as large as 60μm in the 1.1μm diameter fraction. The particle composition of the coarsely-filtered dust used for infecting chicks resembled that of the 9.0μm fraction. The amount of MDV in the coarsely-filtered dust was determined by extracting MDV and using quantitative PCR assay to compare the amounts in the different-sized fractions and in coarsely filtered dust collected at the same time (Figure 2). No significant difference between any of the fractions and the coarsely-filtered dust was detected.
In a separate experiment the ability of dust to induce clinical signs of MD was tested. Two-week-old RIR chicks were infected with 1000 pfu MDV, strain RB1B, and dust collected, coarsely-filtered as described above, and stored at -70°C. Chickens were infected by insufflating dust into the trachea using a microaerosolizer™ (PennCentury, New Jersey, USA). The optimal dose of RB1B-infective dust required to induce clinical signs of MD was determined by insufflating 2.5, 10 and 20 mg amounts into groups of 10 birds. Blood samples were collected at various times. Citrated blood (50μl) was mixed with 500μl 1% saponin and incubated (10 min) and centrifuged (300xg). Pellets were washed and resuspended in phosphate buffered-saline (PBS: pH 7.6) for DNA isolation using a DNeasy© 96 tissue kit (Qiagen, Crawley, UK). Samples were resuspended in 200μl SuperQ water (Quantum, Orpington, UK) and stored at -20ºC pending analysis. The numbers of copies of MDV present in the DNA isolated from the blood was quantified using Taqman™, essentially as described by Kaiser et al. (2003).
Chickens were monitored for clinical signs for 30 days. Liver, heart, proventriculus, kidney and lung were removed post mortem and processed for scoring lymphocyte infiltration as described by Burgess et al. (2001).
After insufflation of infective dust chicks developed clinical signs of MD but there was no evidence of a dose-response relationship (Figure 3). There was evidence that all the chicks became viraemic (Figure 4) and that there was significant lymphocyte infiltration in the tissues with most of the doses used (Figure 5).
We conclude that this method for administering MDV more closely represents the natural process for infection and delivers MDV in its 'native' form, associated with feather and squamous epithelial cell debris. Our method is reproducible and allows delivery of a precisely-timed, quantifiable dose of MDV. This provides the opportunity to investigate early events in the infection process: determining the constitution of the infecting virus (cell-free or keratin-coated virus); identifying those cells that are resident in or line the airways and take up MDV-infected particles to translocate them into the pulmonary tissues. This method also allows investigation of the location (pulmonary tissue or airways) and types of phagocytic cells (macrophage, dendritic cell or epithelial cells lining the airways) that take up MDV and transport it to the lymphoid tissues.
References
Adldinger, H. & Calnek, B. (1973). Pathogenesis of Marek's disease: Early distribution of virus and viral antigens in infected chickens. Journal of the National Cancer Institute 50, 1287-1298.
Beasley, J. N., Patterson, L. T. & McWade, D. H. (1970). Transmission of Marek's disease by poultry house dust and chicken dander. American Journal of Veterinary Research 31, 339-344.
Burgess, S. C., Basaran, B. H. & Davison, T. F. (2001). Resistance to Marek's disease herpesvirus-induced lymphoma is multiphasic and dependent on host genotype. Veterinary Pathology 38, 129-142.
Calnek, B. W., Adldinger, H. K. & Kahn, D. E. (1970). Feather follicle epithelium: a source of enveloped and infectious cell-free herpesvirus from Marek's disease. Avian Diseases 14, 219-233.
Davidson, I. & Borenshtain, R. (2003) Novel applications of feather tip extracts from MDV-infected chicken; diagnosis of commercial broilers, whole genome separation by PFGE and synchronic mucosal infection. FEMS Immunology and Medical Microbiology 38: 199-203.
Gildow, E. M., Williams, J. K. & Lampman, C. E. (1940). The transmission of and resistance to fowl paralysis (lymphomatosis). University of Idaho Agricultural Experiment Station Bulletin 235, 1-22.
Hayter, R.B. 7 Besch, E.L. (1974). Airborne-particle deposition in the respiratory tract of chickens. Poultry Science 53, 1507-1511.
Jurajda, V. & Klimes, B. (1970). Presence and survival of Marek's disease agent in dust. Avian Diseases 14, 188-190.
Kaiser, P., Underwood, G. & Davison, F. (2003). Differential cytokine responses following Marek's disease virus infection of chickens differing in resistance to Marek's disease. Journal of Virology 77, 762-768.
Kenzy, S. G. & Cho, B. R. (1969). Transmission of classical Marek's disease by affected and carrier birds. Avian Diseases 13, 211-214.
Schmittle, S. C. & Eidson, C. S. (1968). Studies on acute Marek's disease IV. Relative susceptibility of different lines and crosses of chickens to GA isolate. Avian Diseases 12, 571-576.
Sevoian, M., Chamberlain, D. M. & Larose, R. N. (1963). Avian lymphomatosis. V. Air-borne transmission. Avian Diseases 7, 102-105.
Underwood, G.J. (2003). An investigation of early events in the pathogenesis of Marek's disease. PhD thesis, University of Bristol.
From Proceedings of the "7th International Symposium on Marek's disease", St. Catherine's College, Oxford, United Kingdom.
