Department of Pediatrics, Box B140, Section of Infectious Diseases1, Department of Pediatrics and Immunology2, University of Colorado Health Sciences Center, 4200 East 9th Avenue, Denver, CO 80262, USA
Department of Virology and Immunology, Christian Medical College Hospital, Vellore, 632004, India3
Department of Pathology, Providence Memorial Hospital, El Paso, TX 79902, USA4
Author for correspondence: Esther Ponnuraj. Fax +1 303 315 4124. e-mail esther.ponnuraj{at}uchsc.edu
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
A wealth of information about RSV pathogenesis has been derived from the mouse model. RSV-specific T helper (Th) lymphocytes are implicated in the inflammatory response to RSV that occurs in naive and vaccine-primed mice and humans (Graham, 1995 ; Openshaw et al., 1988
). Primary and secondary infections of mice with RSV induce a predominant Th1 lymphocyte response detected in spleen (Openshaw et al., 1988
), lung (Graham et al., 1993
) and bronchioalveolar lavage (BAL) specimens (Waris et al., 1996
). Conversely, in animals pre-immunized with FI-RSV vaccine high levels of interleukin-4 (IL-4) mRNA are detected, suggesting that a Th2 lymphocyte response contributes to the enhanced lung pathology (Graham et al., 1993
). In mice, depletion of CD4 cells (Connors et al., 1992
) or depletion of Th2 cytokines IL-4 and IL-10 (Connors et al., 1994
; Tang & Graham, 1994
; Waris et al., 1996
) reduces the severity of enhanced disease. Transfer of CD4 Th2-type cells secreting IL-4 into RSV-infected mice is associated with loss of body weight and with increased infiltration of eosinophils in BAL (Alwan et al., 1994
). These mice experiments suggest that primary infection with RSV elicits a Th1 response, while an RSV infection following FI-RSV vaccine induces a Th2 response and enhanced pathology.
The pathogenesis of enhanced disease has not been fully studied in the primate models. The current work shows that FI-RSV vaccine-induced enhanced disease in bonnet monkeys (Macaca radiata) is associated with virus replication in the inflammatory cells around blood vessels.
![]() |
Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Virus stock preparation.
The RSV Long strain was selected because of its human origin. It was grown in HEp-2 cells by infecting 150 cm2 plates at an m.o.i. of 0·01. When the CPE reached 50%, the medium was replaced by serum-free DMEM. When the CPE was approximately 90%, the cell monolayer was harvested with a rubber policeman into DMEM, snap-frozen in liquid nitrogen, thawed, clarified and snap-frozen in 1 ml aliquots. An aliquot of the virus stock was titrated using an enzyme-linked immuno-plaque assay, as described later.
Infection of monkeys.
Monkeys were anaesthetized with 5 mg/kg ketamine intravenously and intubated using a 2·0 or 2·5 Fr gauge endotracheal tube. The virus was instilled into the bronchi of the left and right lungs. Infected and uninfected animals were housed in separate rooms in individual cages.
Bronchoalveolar lavage (BAL).
Monkeys were anaesthetized with ketamine, intubated and a BAL was performed using three 5 ml/kg aliquots of normal saline through a cuffed 2·53 Fr gauge endotracheal tube. Lavage fluid was obtained using an 8 Fr gauge feeding tube passed through the endotracheal tube. Aliquots of aspirated BAL fluid were frozen for virus culture and antibody activity. BAL samples were collected from two monkeys from each of the groups on days 1, 2 and 3 post-inoculation (p.i.), and from all monkeys on day 7 p.i.
Collection of serum samples.
Blood samples were collected on the day of RSV inoculation and at necropsy for antibody measurements. Serum samples from the blood were frozen at -70 °C prior to assay.
Specimen collection at necropsy.
At necropsy the chest was opened and sections of the upper and lower lobes of the left lung were placed in M4 medium (Bio Whittaker) for virus culture. Perihilar lymph nodes were dissected out and placed in RPMI 1640 (Gibco/BRL) with 10% pooled heat-inactivated monkey serum (RPMI-MS) for lymphocyte culture. The pulmonary artery was identified and a solution of 30% barium sulfate and 10% gelatin in PBS at 60 °C was infused at a pressure of 80 mm Hg. The left bronchus was then clamped and the right lung inflated for at least 2 h with paraformaldehydelysineperiodate infused into the trachea via the endotracheal tube at 40 cm pressure to fix lung tissues in an expanded state. The fixed lungs were transferred to 70% ethanol for histology and in situ hybridization.
Histological studies.
Peripheral and central regions of upper and lower lobes of paraformaldehyde-fixed lungs were processed and embedded in paraffin. Sections of 5 µm were stained with haematoxylin and eosin (H&E). The severity of the pulmonary inflammatory response was scored in a blinded manner according to a histopathological scheme similar to that described by Piedra et al. (1993) . Four sections from the central and peripheral parts of the upper and lower right lung were scored according to the degree of inflammation in interstitium, alveoli and surrounding airways and vessels. A value of 0 (none), 1 (minimal), 2 (mild), 3 (moderate) or 4 (severe) was assigned to each histological site, and the scores were totalled for each animal. The sum of these scores was used as a total lung inflammation score. Every section was scanned entirely to assign an inflammation score with the most inflamed areas evaluated. The highest score for each parameter (peribronchiolar, perivascular, interstitial and alveolar) that involved at least 25% of each section evaluated was used.
Virus culture.
Two parts of each BAL sample were mixed with 1 part of M4 medium. The samples were frozen at -70 °C until they were titrated for virus. Lung samples collected at necropsy were weighed, suspended in M4 medium (Simoes et al., 1999 ), clarified and frozen at -70 °C until they were tested by a plaque assay using immuno-staining for the detection of virus plaques (Piedra et al., 1989
).
In situ hybridization for the detection of RSV in lung tissue.
To localize sites of virus replication in the lung, in situ hybridization was done on all six groups of animals. RSV N gene cDNA (Johnson & Collins, 1989 ) (a kind gift from Peter Collins) was transcribed with SP6 RNA polymerase. An anti-sense probe was used to detect the positive-sense replicative intermediates that are present during RSV replication. RNA probe was generated from N gene cDNA cloned into a pGEM vector. Transcription was performed using 1x Ribomax buffer (Promega) with 40 U RNasin (Promega) and 1 mM each of ATP, GTP, CTP, 0·65 mM UTP and 0·35 mM digoxigenin-11-UTP (Boehringer Mannheim). The optimal concentration of each probe in preliminary trials was determined. In situ hybridization with sense probes generated by transcription with T7 polymerase was also done in order to find out if RSV N gene-specific genomic RNA was present. In situ hybridization was done on sections from the lower central lobe of each monkey according to the methods described earlier (Ponnuraj et al., 1998
; Rotbart et al., 1988
). The specificity of in situ hybridization was checked by testing positive sections with a riboprobe derived from the 5' non-coding region of poliovirus. To quantify in situ labelling of infiltrates, arterioles identified by intravascular barium (size 30300 µm in diameter) with >50% circumferential labelling were examined under 100x magnification and the radial thickness of the area with signals measured using a micrometer eyepiece. Ten measurements were made per slide and the sums of the readings were expressed as a score on a scale of 0100.
Identification of cells in inflammatory infiltrates in FI-RSV-enhanced disease.
Histological sections of lungs from the animals immunized with FI-RSV vaccine and from control animals were used for immuno-staining to identify the specific cell types in the inflammatory infiltrates. Effective cross-reactions were identified with human cytokeratin (an epithelial cell marker), CD45 (a leukocyte common antigen marker), CD68 (a macrophage marker) and myeloperoxidase (a marker for macrophages and polymorphs). These monoclonal antibodies were obtained from Dako Corporation. Monoclonal antibody binding was detected by Vectastain ABC Elite kit (Vector Laboratories) as per the instructions provided by the manufacturer, 3,3'-diaminobenzidine was used as the substrate and the sections were counter-stained with Gills no. 3 haematoxylin.
Anti-RSV antibody assays.
Neutralizing antibody titres were performed using a micro-neutralizing assay as previously described (Simoes et al., 1999 ). Serum IgG and BAL fluid IgA antibodies to RSV were detected by ELISA. UV-irradiated RSV antigen and control antigen for coating ELISA plates were prepared according to a protocol previously used (Vai et al., 1985
). The RSV antigen and cell culture control antigen were diluted 1:20 in PBS and incubated on Immulon II plates overnight. The wells were then blocked with 1% gelatin. Monkey sera and a parallel reference standard positive control serum were diluted 1:101:103 and the BAL samples were diluted 1:11:10. Diluted samples were added to the antigen and control wells and incubated for 30 min. Subsequently the wells were washed with PBS containing 0·1% Tween 20 and incubated with peroxidase-conjugated anti-isotype antibodies (Cappel, Organon Teknika; cat. #55226). The absorbance values (A) were read on a Dynatech reader and the log of the values was regressed against the log of dilution. The IgG and IgA antibody results in monkey serum and BAL samples were expressed as a percentage of the reference positive control.
Lymphocyte proliferation studies.
Lymphocytes were isolated from peripheral blood and perihilar lymph nodes of each monkey. The lymphocyte concentration was adjusted to be 106 cells/ml in RPMI-MS and incubated at 37 °C. Lymphocytes were cultured without stimulation, or were stimulated with UV-irradiated RSV antigen. After 6 days of culture the cells were pulsed with 0·25 mCi (9·25 MBq) [methyl-3H]thymidine (ICN) for 4 h. The cultures were then harvested onto glass fibres in a 96-well harvester and the incorporated thymidine was determined with a Betaplate reader (Wallac). The mean value of triplicate experiments was determined and the preliminary analysis was done on the basis of the stimulation index (SI). A value of SI 3 was taken as evidence of a proliferative response.
Statistical methods.
The significance of differences between groups with respect to inflammation scores, virus replication in the lung, antibody response and lymphocyte proliferation was tested by the two-tailed Student t-test. A Chi-square for trend was used to test for differences in virus replication in the BAL samples for FI-RSV-vaccinated animals and those with primary infections. Correlation between inflammation scores and host factors was done using a correlation matrix, correcting for groups. The significance of Spearmans rank correlation coefficient was obtained using a t-test.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
|
Replication of RSV in the BAL and lung samples is increased in the FI-RSV-vaccinated animals
Virus titres in the BAL samples were high on day 1 p.i. and decreased to undetectable levels by day 3 p.i. in primary infection (Fig. 3a). Animals with tertiary infection had undetectable levels of virus in BAL at each of these time-points. In animals immunized with FI-RSV (full dose), the titre increased with time. Virus titres remained low in animals immunized with low dose FI-RSV. Titres in animals with FI-Vero vaccine resembled those of primary infection. The full dose FI-RSV-immunized animals had about 10-fold more RSV in their lungs than animals with primary infection (Fig. 3b
) and this difference was statistically significant (P=0·001). The animals given the low dose FI-RSV vaccine (Group V) had significantly more virus isolated from their lungs than animals with primary (Group II, P=0·001) or tertiary (Group III, P=0·001) infection. Animals given FI-Vero cell vaccine demonstrated similar levels of virus replication as seen in a primary infection. Animals undergoing tertiary infection did not shed virus in the BAL and had significantly less virus in the lung (P=0·01).
|
|
|
|
Correlation between inflammation scores and virus titres
There was positive correlation between inflammation scores and virus titres. The correlation of virus titres versus total inflammation scores was r=0·66 (P=0·0001) (Fig. 7a). Better correlation was seen with perivascular scores versus virus titre r=0·79 (P=0·0001) (Fig. 7b
) and peribronchial scores versus virus titres r=0·78 (P=0·0001) (Fig. 7c
). Lower correlation was seen with interstitial scores versus virus titre r=0·6201 (P=0·0003) (Fig. 7d
).
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Enhanced pulmonary pathology following FI-RSV vaccine immunization occurs in all animals at all of the evaluated pulmonary sites. Statistically significant differences are identified in total, peribronchial, perivascular, interstitial and alveolar inflammation scores between the FI-RSV vaccine-immunized group as compared to the primary infection group. These differences are greatest in peribronchiolar and perivascular sites, with lesser but consistent differences also present at interstitial and alveolar locations. In all cases, the inflammation in FI-RSV vaccine-immunized animals was greater than that in animals with primary infection.
While overall lung inflammation scores for the FI-Vero cell-immunized animals were not statistically different from the scores of primary infected animals, the peribronchial and perivascular sites had scores that were significantly increased as compared to primary disease. This increased inflammation did not reach the levels seen in the FI-RSV animals, however. In BALB/c mice also mock vaccine produced histopathological changes of a lower magnitude in comparison with those immunized with the FI-RSV vaccine (Bolen et al., 2001 ). Immunization with FI-Vero cell vaccine has already been reported to create enhanced pulmonary pathology upon subsequent RSV infection in cotton rats (Piedra et al., 1993
). The mechanism by which an FI-Vero cell vaccine creates enhanced pathology is not known. One possibility is that the small amounts of foetal calf serum proteins in the Vero cell vaccine sensitize the vaccine recipients and that repeat exposure to these proteins in the RSV virus preparation elicits an inflammatory response. This enhanced inflammatory response might be unrelated to RSV and its localization to central sites (peribronchiolar and perivascular).
Of interest is that the FI-Vero cell vaccine-immunized animals showed greater inflammation centrally and less inflammation peripherally, as compared to animals with primary infection. The FI-RSV vaccine-enhanced inflammation was significantly greater than that of the primary infection at peribronchiolar, perivascular and interstitial sites. In contrast, tertiary infection in this animal model showed a protective effect from the prior RSV exposure. With the tertiary infection there was less inflammation at interstitial and alveolar sites as compared to primary infection. The pattern of protection from pulmonary inflammation in animals with tertiary infection shows a protective effect that is greatest at peripheral sites. Therefore, it appears that RSV-independent mechanisms can cause perivascular and peribronchiolar inflammation. This conclusion would suggest that an effective RSV vaccine should provide the least possible non-specific central pulmonary inflammatory enhancement while creating the greatest possible immune-specific peripheral pulmonary protection.
Augmented virus replication in FI-RSV-immunized animals
The finding of increased RSV replication in FI-RSV vaccine-immunized animals after RSV infection may be fundamental to understanding enhanced disease. The 10-fold increment in virus production in the lungs of the immunized animals is accompanied by an increase in titre in BAL samples with time. Prior studies in the African green monkey model using the A2 strain of RSV have shown similar increases in RSV titres in both lung and BAL samples from animals immunized with FI-RSV vaccine (Kakuk et al., 1993 ). In those animals 510-fold higher virus titres were observed as compared to lungs of animals immunized with an adjuvant alone.
The augmented virus replication documented by virus titres in the lungs of monkeys with FI-RSV vaccine-enhanced disease is further corroborated by the in situ hybridization results. In situ hybridization shows that the increased virus replication occurs at peribronchiolar and perivascular sites where immunostaining identified some of the inflammatory cells to be macrophages. In humans, macrophages have been shown to support virus replication (Becker et al., 1992 ; Midulla et al., 1989
; Panuska et al., 1992
) while other inflammatory cells such as lymphocytes do not seem to support a productive infection by RSV (Domurat et al., 1985
; ODonnell et al., 1998
; Panuska et al., 1992
). In the bonnet monkey the concentration of RSV message is greatest at perivascular and peribronchiolar sites. Comparable sections show that macrophages are the principal population in these sites of increased RSV replication.
The increase in RSV replication in perivascular sites of the FI-Vero cell vaccine group compared with the primary infection group was detected only by mRNA and not by virus culture. This is probably because quantitative results on hybridization for RSV mRNA were possible only in perivascular sites, and it is in these sites that the FI-Vero cell vaccine group animals develop an inflammatory response when challenged by infectious virus.
Previous studies in mice immunized with the FI-RSV vaccine, with the F or G protein of RSV or with vaccinia virus vectors expressing the F or G protein showed no evidence for increased virus replication after challenge with RSV, despite the occurrence of enhanced disease in the lungs (Graham et al., 1995 ; Hussell et al., 1997
; Waris et al., 1996
). Thus, the findings in the mouse model of FI-RSV disease contrast with those in primates. Mouse macrophages do not have a productive RSV infection in vitro (Franke-Ullmann et al., 1995
). If this is true of macrophages in vivo, then FI-RSV vaccine-immunized mice might not show increased virus replication. The mechanisms for enhanced disease in mice and monkeys appear to be different. While there is concordance between the African green and bonnet monkey models, there is discordance between the primate and mouse model in the extent of virus replication following use of the FI-RSV vaccine. The concordance between enhanced virus replication in two separate non-human primate models of RSV disease emphasizes the importance of using primates rather than mice to evaluate vaccines and immunomodulators related to RSV disease.
Immune response following immunization with FI-RSV vaccine
Study of bonnet monkeys with tertiary infection shows that prior RSV respiratory exposure appears to stimulate very good antibody and lymphocyte proliferative responses, with no virus recovered from the lungs. Also the virus titres in the BAL samples remained low, with minimal inflammation seen at perivascular and peribronchiolar sites and the least inflammation at peripheral sites. In animals immunized with the FI-RSV high dose, there were very low amounts of antibody and low levels of proliferative response, with high levels of virus in the lungs and a temporal increase in virus titres in the BAL samples. There was also increased inflammation in all four intrapulmonary sites examined. In animals with low dose FI-RSV, there were moderate amounts of antibody and lymphocyte proliferative response, accompanied by lower virus titres in the lungs. The RSV titre in BAL of low dose FI-RSV animals remained low and inflammation did not extend to the alveoli. The trend in restriction of pathology to peribronchial and perivascular areas in animals with tertiary infection and animals immunized with low dose FI-RSV coupled with the absence of a temporal increase in virus titres in the BAL samples show that there is limited spread of virus from the initial central sites of replication to more peripheral sites. The blocking of virus spread peripherally may be attributed to the presence of anti-RSV antibody. If correct, this hypothesis emphasizes the importance of antibody production in any RSV-specific immunization protocol.
The bonnet monkey model did not reproduce the inverse relationship between susceptibility to enhanced disease and vaccine dose previously shown in mice (Fischer et al., 1997 ) and cotton rats (Murphy et al., 1990
). In bonnet monkeys, it appears that antibody levels and T cell proliferation were greater at the low dose. Antibody and lymphocyte responses were lower in animals immunized with a high dose of FI-RSV vaccine, perhaps due to the induction of regulatory T cells or to some other suppressive mechanism. Our observation of low levels of functional antibody in FI-RSV-vaccinated animals mimics the response to the vaccine in human infants. Following two doses of FI-RSV vaccine neutralizing antibodies were found only in 17% of infants (Kim et al., 1969
).
Enhanced RSV disease in mice has been attributed to alterations in the ratio of -IFN/IL-4-producing (Th1/Th2) cells (Fischer et al., 1997
). In the FI-RSV vaccine-primed animals there is an increase in CD4 cells predominantly producing IL-4 (Graham et al., 1993
; Srikiatkhachorn & Braciale, 1997a
, b
). The results with mice have lead to the speculation that enhanced pathology following FI-RSV vaccine in naive infants might also result from a change in cytokine balance (Chanock et al., 1992
). The results from the bonnet monkey model suggest an alternative explanation. The enhanced pathology following FI-RSV vaccine immunization is in some way a consequence of increased replication of RSV within sites of inflammation. Increased virus load and infection of macrophages might augment the inflammatory response.
Antibody-mediated endocytosis is a likely mechanism by which RSV could enter macrophages. It is known that the IgG1 subclass of antibody binds to Fc receptors on macrophages (Van de Winkel & Capel, 1993 ) and therefore production of this subclass of antibody may be responsible for any augmented replication of RSV that might occur in macrophages of FI-RSV-immunized animals. The absence of augmented replication of RSV in mice, even when IgG1 predominates in the antibody response (as elegantly shown by Bembridge et al., 2001
), argues for monkey studies in the evaluation of experimental RSV vaccines.
Evaluation of RSV vaccines in children could be dangerous. Our results show that the bonnet monkey model offers a reproducible alternative that mimics human disease but permits analysis of variables including immunogen route and attenuation, virus replication and immune response.
![]() |
Acknowledgments |
---|
This work was supported by grant #AI37271 from the National Institute of Allergy and Infectious Diseases and by a grant from the Department of Biotechnology Government of India and the USIndia Vaccine Action Program.
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Becker, S., Soukup, J. & Yankaskas, J. R. (1992). Respiratory syncytial virus infection of human primary nasal and bronchial epithelial cell cultures and bronchoalveolar macrophages. American Journal of Respiratory Cell and Molecular Biology 6, 369-374.[Medline]
Bembridge, G. P., Rodriguez, N., Garcia-Beato, R., Nicolson, C., Melero, A. & Taylor, G. (2001). Respiratory syncytial virus infection of gene gun vaccinated mice induces Th-2 driven pulmonary eosinophilia even in the absence of sensitisation to the fusion (F) or attachment (G) protein. Vaccine 19, 1038-1046.
Bolen, A., Andeweg, A., Kwakkel, J., Lockhorst, W., Bestebroer, T., Dormans, J. & Kimman, T. (2001). Both immunization with a formalin-inactivated respiratory syncytial virus (RSV) vaccine and a mock antigen vaccine induce severe lung pathology and a Th2 cytokine profile in RSV challenged mice. Vaccine 19, 982-991.
Chanock, R. M., Parrott, R. H., Connors, M., Collins, P. L. & Murphy, B. R. (1992). Serious respiratory tract disease caused by respiratory syncytial virus: prospects for improved therapy and effective immunization. Pediatrics 90, 137-143.[Medline]
Collins, P. L., McIntosh, K. & Chanock, R. M. (1996). Respiratory syncytial virus. In Fields Virology , pp. 1313-1351. Edited by B. N. Fields, D. M. Knipe & P. M. Howley. Philadelphia:LippincottRaven.
Connors, M., Kulkarni, A. B., Firestone, C. Y., Holmes, K. L., Morse, H. C.III, Sotnikov, A. V. & Murphy, B. R. (1992). Pulmonary histopathology induced by respiratory syncytial virus (RSV) challenge of formalin-inactivated RSV-immunized BALB/c mice is abrogated by depletion of CD4+ cells. Journal of Virology 66, 7444-7451.[Abstract]
Connors, M., Giese, N. A., Kulkarni, A. B., Firestone, C. Y., Morse, H. C.III & Murphy, B. R. (1994). Enhanced pulmonary histopathology induced by respiratory syncytial virus (RSV) challenge of formalin-inactivated RSV-immunized BALB/c mice is abrogated by depletion of interleukin-4 (IL-4) and IL-10. Journal of Virology 68, 5321-5325.[Abstract]
Domurat, F., Roberts, N. J.Jr, Walsh, E. E. & Dagan, R. (1985). Respiratory syncytial virus infection of human mononuclear leukocytes in vitro and in vivo. Journal of Infectious Diseases 152, 895-902.[Medline]
Fischer, J. E., Johnson, J. E., Kuli-Zade, R. K., Johnson, T. R., Aung, S., Parker, R. A. & Graham, B. S. (1997). Overexpression of interleukin-4 delays virus clearance in mice infected with respiratory syncytial virus. Journal of Virology 71, 8672-8677.[Abstract]
Franke-Ullmann, G., Pfortner, C., Walter, P., Steinmuller, C., Lohmann-Matthes, M. L., Kobzik, L. & Freihorst, J. (1995). Alteration in pulmonary macrophage function by respiratory syncytial virus in vitro. Journal of Immunology 154, 268-280.
Fulginiti, V. A., Eller, J. J., Sieber, O. F., Joyner, J. W., Minamitani, M. & Meiklejohn, G. (1969). Respiratory virus immunization. I. A field trial of two inactivated respiratory virus vaccines: an aqueous trivalent parainfluenza virus vaccine and an alum-precipitated respiratory syncytial virus vaccine. American Journal of Epidemiology 89, 435-448.[Medline]
Graham, B. S. (1995). Pathogenesis of respiratory syncytial virus vaccine-augmented pathology. American Journal of Respiratory and Critical Care Medicine 152, S63-S66.[Medline]
Graham, B. S., Henderson, G. S., Tank, Y. W., Lu, X., Neuzil, K. M. & Colley, D. G. (1993). Priming immunization determines T helper cytokine mRNA expression patterns in lungs of mice challenged with respiratory syncytial virus. Journal of Immunology 151, 2032-2040.
Hussell, T., Khan, U. & Openshaw, P. (1997). IL-12 treatment attenuates T helper cell type 2 and B cell responses but does not improve vaccine-enhanced lung illness. Journal of Immunology 159, 328-334.[Abstract]
Johnson, P. R. & Collins, P. L. (1989). The 1B (NS2), 1C (NS1) and N proteins of human respiratory syncytial virus (RSV) of antigenic subgroups A and B: sequence conservation and divergence within RSV genomic RNA. Journal of General Virology 70, 1539-1547.[Abstract]
Kakuk, T. J., Soike, K. & Brideau, R. J. (1993). A human respiratory syncytial virus (RSV) primate model of enhanced pulmonary pathology induced with a formalin-inactivated RSV vaccine but not a recombinant FG subunit vaccine. Journal of Infectious Diseases 167, 553-561.[Medline]
Kim, H. W., Canchola, J. G, Brandt, C. D., Pyles, G., Chanock, R. M., Jensen, K. & Parrott, R. H. (1969). Respiratory syncytial virus diease in infants despite prior administration of antigenic inactivated vaccine. American Journal of Epidemiology 89, 422-433.[Medline]
Midulla, F., Huang, Y. T., Gilbert, I. A., Cirino, N. M., McFadden, E. R.Jr & Panuska, J. (1989). Respiratory syncytial virus infection of human cord and adult blood monocytes and alveolar macrophages. American Review of Respiratory Disease 140, 771-777.[Medline]
Murphy, B. R., Prince, G. A., Lawrence, L. A., Croen, K. D. & Collins, P. L. (1990). Detection of respiratory syncytial virus (RSV) infected cells by in situ hybridization in the lungs of cotton rats immunized with formalin-inactivated virus or purified RSV F and G glycoprotein subunit vaccine and challenged with RSV. Virus Research 16, 153-162.[Medline]
ODonnell, D. R., McGarvey, M. J., Tully, J. M., Balfour-Lynn, I. M. & Openshaw, P. J. M. (1998). Respiratory syncytial virus RNA in cells from the peripheral blood during acute infection. Journal of Pediatrics 133, 272-274.[Medline]
Openshaw, P. J. M., Pemberton, R. M., Ball, L. A., Wertz, G. W. & Askonas, B. A. (1988). Helper T cell recognition of respiratory syncytial virus in mice. Journal of General Virology 69, 305-312.[Abstract]
Panuska, J. R., Hertz, M. I., Taraf, H., Villani, A. & Cirino, N. M. (1992). Respiratory syncytial virus infection of alveolar macrophages in adult transplant patients. American Review of Respiratory Disease 145, 934-939.[Medline]
Piedra, P. A., Faden, H. S., Camussi, G., Wong, D. T. & Ogra, P. L. (1989). Mechanism of lung injury in cotton rats immunized with formalin-inactivated respiratory syncytial virus. Vaccine 7, 34-41.[Medline]
Piedra, P. A., Wyde, P. R., Castleman, W. L., Ambrose, M. W., Jewell, A. M., Speelman, D. J. & Hildreth, S. W. (1993). Enhanced pulmonary pathology associated with the use of formalin-inactivated respiratory syncytial virus vaccine in cotton rats is not a unique viral phenomenon. Vaccine 11, 1415-1423.[Medline]
Ponnuraj, E. M., John, T. J., Levin, M. J. & Simoes, E. A. F. (1998). Cell-to-cell spread of poliovirus in the spinal cord of bonnet monkeys (Macaca radiata). Journal of General Virology 79, 2393-2403.[Abstract]
Prince, G. A., Jenson, A. B., Horswood, R. L., Camargo, E. & Chanock, R. M. (1978). The pathogenesis of respiratory syncytial virus infection in cotton rats. American Journal of Pathology 93, 771-784.[Abstract]
Prince, G. A., Jenson, A. B. & Hemming, V. G. (1986). Enhancement of respiratory syncytial virus pulmonary pathology in cotton rats by prior intramuscular inoculation of formalin inactivated virus. Journal of Virology 57, 721-728.[Medline]
Rotbart, H. A., Abzug, M. J., Murray, R. S., Murphy, N. L. & Levin, M. J. (1988). Intracellular detection of sense and antisense enteroviral RNA by in situ hybridization. Journal of Virological Methods 22, 295-301.[Medline]
Simoes, E. A. F., Hayward, A. R., Ponnuraj, E. M., Straumanis, J. P., Stenmark, K. R., Wilson, H. & Babu, P. G. (1999). Respiratory syncytial virus (RSV) infects the Bonnet monkey, Macaca radiata. Pediatric Pathology 2, 316-326.
Srikiatkhachorn, A. & Braciale, T. J. (1997a). Virus-specific CD8+ T lymphocytes downregulate T helper cell type 2 cytokine secretion and pulmonary eosinophilia during experimental murine respiratory syncytial virus infection. Journal of Experimental Medicine 71, 421-432.
Srikiatkhachorn, A. & Braciale, T. J. (1997b). Virus-specific memory and effector T lymphocytes exhibit different cytokine responses to antigens during experimental murine respiratory syncytial virus infection. Journal of Virology 71, 678-685.[Abstract]
Tang, Y. W. & Graham, B. S. (1994). Anti-IL-4 treatment at immunization modulates cytokine expression, reduces illness, and increases cytotoxic T lymphocyte activity in mice challenged with respiratory syncytial virus. Journal of Clinical Investigation 94, 1953-1958.[Medline]
Vai, J. A., Leary, P. L. & Levin, M. J. (1985). Specificity of the blastogenic response of human mononuclear cells to herpes virus antigens. Infection and Immunity 20, 646.
Van de Winkel, J. G. & Capel, P. J. A. (1993). Human IgG Fc receptor heterogeneity: molecular aspects and clinical implications. Immunology Today 14, 215-221.[Medline]
Waris, M. E., Tsou, C., Erdman, D. D., Zaki, S. R. & Anderson, L. J. (1996). Respiratory syncytial virus infection in BALB/c mice previously immunized with formalin-inactivated virus induces enhanced pulmonary inflammatory response with a predominant Th2 like cytokine pattern. Journal of Virology 70, 2852-2860.[Abstract]
Received 30 May 2001;
accepted 31 July 2001.