Identification of small-animal and primate models for evaluation of vaccine candidates for human metapneumovirus (hMPV) and implications for hMPV vaccine design

Mia MacPhail1, Jeanne H. Schickli1, Roderick S. Tang1, Jasmine Kaur1, Christopher Robinson1, Ron A. M. Fouchier2, Albert D. M. E. Osterhaus2, Richard R. Spaete1 and Aurelia A. Haller1,{dagger}

1 MedImmune Vaccines Inc., 297 North Bernardo Avenue, Mountain View, CA 94043, USA
2 Department of Virology, Erasmus Medical Center, Rotterdam, The Netherlands

Correspondence
Aurelia A. Haller
aurelia.haller{at}globeimmune.com


   ABSTRACT
Top
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Human metapneumovirus (hMPV), a recently identified paramyxovirus, is the causative agent of respiratory tract disease in young children. Epidemiological studies have established the presence of hMPV in retrospective as well as current clinical samples in Europe, USA, Canada, Hong Kong and Australia. The hMPV disease incidence rate varied from 7 to 12 %. This rate of disease attack places hMPV in severity between respiratory syncytial virus and human parainfluenza virus type 3, two common respiratory pathogens of young children, the elderly and immunosuppressed individuals. To evaluate the effectiveness and safety of future hMPV antiviral drugs, therapeutic and prophylactic monoclonal antibodies (mAbs), and vaccine candidates, it was necessary to identify small-animal and primate models that efficiently supported hMPV replication in the respiratory tract and produced neutralizing serum antibodies, commonly a clinical correlate of protection in humans. In this study, various rodents (mice, cotton rats, hamsters and ferrets) and two primate species, rhesus macaques and African green monkeys (AGMs), were evaluated for hMPV replication in the respiratory tract. The results showed that hamsters, ferrets and AGMs supported hMPV replication efficiently and produced high levels of hMPV-neutralizing antibody titres. Hamsters vaccinated with subgroup A hMPV were protected from challenge with subgroup A or subgroup B hMPV, which has implications for hMPV vaccine design. Although these animal models do not mimic human hMPV disease signs, they will nevertheless be invaluable for the future evaluation of hMPV antivirals, mAbs and vaccines.

{dagger}Present address: GlobeImmune Inc., Aurora, CO 80010, USA.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Human metapneumovirus (hMPV) is a recently identified paramyxovirus pathogen associated with upper and lower respiratory tract infections. Initially, hMPV was isolated from children with clinical symptoms indicative of human respiratory syncytial virus (RSV) infection where RSV was not detected (van den Hoogen et al., 2001). Manifestations of hMPV disease range from mild upper respiratory problems to severe lower respiratory disease such as cough, bronchiolitis and pneumonia (van den Hoogen et al., 2001, 2003; Boivin et al., 2002). Nearly all children in The Netherlands have been exposed to hMPV by the age of 5 years and the virus has been detected in clinical samples from 50 years ago (van den Hoogen et al., 2001). Preliminary epidemiological reports have estimated an hMPV disease incidence rate of 7–12 % in young children (Williams et al., 2004; Peiris et al., 2003; Freymuth et al., 2003; Maggi et al., 2003; Jartti et al., 2002; Nissen et al., 2002; Peret et al., 2002; van den Hoogen et al., 2001). The signs of hMPV infection are similar to those caused by RSV and human parainfluenza virus type 3 (hPIV3) and hospitalization of children with acute lower respiratory tract infections is necessary in some cases (Hall, 2001; Pelletier et al., 2002; van den Hoogen et al., 2003). Recently, Greensill et al. (2003) reported the detection of hMPV in bronchoalveolar lavage fluids from 21 of 30 infants (70 %) ventilated for RSV bronchiolitis. FDA-approved hMPV vaccines, monoclonal antibodies (mAbs) and antivirals to combat hMPV infection and disease are currently not available for treatment or prevention.

On the basis of electron microscopy and comparison of viral genome sequence organization, hMPV was assigned to the Metapneumovirus genus of the Paramyxoviridae family. hMPV contains a non-segmented, negative-sense RNA genome approximately 13 370 nucleotides in length (van den Hoogen et al., 2002; Biacchesi et al., 2003). The genomic organization for hMPV is similar but not identical to that of RSV. hMPV harbours open reading frames (ORFs) for at least eight viral proteins (3'-N-P-M-F-M2-SH-G-L-5'). However, hMPV lacks the non-structural proteins NS1 and NS2 of RSV, and the gene order of RSV and hMPV differs significantly. In hMPV, the M2-1 and small hydrophobic (SH) protein ORFs are located between the fusion (F) and glycoprotein (G) genes, which is unlike the RSV genomic organization. The M2-1 protein of RSV promotes processive RNA synthesis and readthrough at RSV gene junctions. Future studies will determine whether the M2-1 protein of hMPV plays a similar role in hMPV replication. Deletion of a number of RSV genes such as M2-2, SH, G, NS1 and NS2 was not deleterious to the virus and such RSV deletion mutants have been evaluated in primates as putative live attenuated vaccine candidates (Jin et al., 2003). Similar strategies will be employed to generate live attenuated hMPV vaccine candidates once a reverse genetics system is established for hMPV. Two subgroups of hMPV (subgroups A and B) were identified based on sequence comparison of the F and G genes derived from a number of different clinical isolates. Within each subgroup, two genetic sublineages were categorized, A1 and A2 for hMPV subgroup A, and B1 and B2 for hMPV subgroup B. The F genes of subgroups A and B are highly conserved and display >95 % identity at the amino acid level. In contrast, the hMPV G proteins are variable and show only 35 % amino acid identity (van den Hoogen et al., 2004; Biacchesi et al., 2003). The F protein of hMPV/NL/1/00 displays a high degree of conservation with the F protein of a related metapneumovirus, avian pneumovirus subgroup C (APV C), a fowl pathogen. Avian pneumovirus causes ‘swollen head syndrome’ in chickens and rhinotracheitis in turkeys (Buys et al., 1989). In particular, the ectodomains of the F proteins of APV C and hMPV are closely related. hMPV may have evolved from the avian metapneumovirus, although hMPV is not infectious for birds (van den Hoogen et al., 2001). Efforts to generate live attenuated hMPV vaccines, as well as to prepare neutralizing hMPV mAbs, have been initiated by a number of research institutions.

We recently reported the generation of a putative hMPV vaccine candidate, a recombinant live attenuated bovine/human PIV3 expressing the hMPV F protein (Tang et al., 2003). In the future, a plethora of putative hMPV vaccines, mAbs and antivirals will be generated, and animal models will be needed to evaluate these approaches for safety, efficacy and immunogenicity, where applicable. In this study, a number of small-animal and primate models were investigated to determine their ability to support hMPV replication in the respiratory tract and to produce an effective immune response. Mice, cotton rats, hamsters and ferrets were studied as small-animal models for hMPV replication, and rhesus monkeys and African green monkeys (AGMs) were tested as primate models. The results showed that Syrian golden hamsters, ferrets and AGMs were highly susceptible to hMPV infection and supported high levels of hMPV replication in the lower (LRT) and upper respiratory tract (URT). The data generated using these animal models will support licensing of hMPV vaccines, antivirals and prophylactic mAbs for high-risk children.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Cells and viruses.
Vero cells were maintained in minimal essential medium (MEM; JRH Biosciences) supplemented with 10 % foetal bovine serum (FBS), 2 mM L-glutamine, non-essential amino acids and antibiotics. All media components were purchased from HyClone Laboratories. hMPV/NL/1/00 (prototype A1), hMPV/NL/1/99 (prototype B1) and RSV A2 were grown in Vero cells in Opti-MEM (Gibco-BRL) in the presence of gentamicin. Vero cells were infected with hMPV at an m.o.i. of 0·1 and incubated at 35 °C for 9–11 days without media changes in the absence of trypsin. To generate RSV A2 virus stocks, Vero cells were infected with an m.o.i. of 0·1 and incubated at 35 °C for 4–5 days. The cells and supernatants were collected, stabilized by adding 10x SPG (2·18 M sucrose, 0·038 M KH2PO4, 0·072 M K2HPO4, 0·054 M L-glutamate) to a final concentration of 1x and the virus stocks were stored at –70 °C. The virus titres were determined by plaque assays on Vero cells. Plaques were quantified after immunoperoxidase staining using hMPV guinea pig, hMPV ferret or RSV goat polyclonal antisera (Biogenesis).

Small-animal studies.
Five-week-old hamsters (Mesocricetus auratus), BALB/c mice (Mus musculus), cotton rats (Sigmodon hispidus) (six animals per group) or ferrets (Mustela putorius) (four animals per group) were infected intranasally with 1x106 p.f.u. hMPV/NL/1/00. The animals were housed in individual micro-isolator cages. Four days post-infection, the nasal turbinates and lungs of the animals were harvested and homogenized. The titre of virus present in the tissues was determined by plaque assays on Vero cells, which were immunostained with hMPV polyclonal antisera. Ferrets were also monitored for changes in body temperature. For polyclonal antibody production, infected ferrets were maintained for 28 days post-infection at which time the animals were exsanguinated.

For the challenge studies, hamsters (18 animals per group) were infected intranasally with 1x105 p.f.u. hMPV/NL/1/00, hMPV/NL/1/99 or placebo medium (Opti-MEM) in a 0·1 ml volume. The different groups were maintained separately in micro-isolator cages. Four days post-infection, the nasal turbinates and lungs of six animals were harvested and homogenized. The titre of virus present in the tissues was determined by plaque assays on Vero cells by immunostaining with hMPV polyclonal antisera. Four weeks post-immunization, the remaining 12 animals were challenged intranasally with 1x106 p.f.u. hMPV/NL/1/00 (six animals) or 2x105 p.f.u. hMPV/NL/1/99 (six animals) in a 0·1 ml volume. Four days post-challenge, the nasal turbinates and lungs of the animals were collected and assayed for challenge virus replication by plaque assays on Vero cells. Plaques were visualized for quantification by immunostaining with hMPV polyclonal antisera.

Primate studies.
Three hMPV-seronegative and three RSV-seronegative AGMs (Cercopithecus aethiops) and four hMPV-seronegative rhesus monkeys (Macaca mulatta) (1–4 years old, 2–5 kg) were identified using an hMPV plaque reduction neutralization assay (PRNA) (described below) and an RSV F IgG ELISA (Immuno-Biological Laboratories), respectively, using primate pre-sera collected on day –18 prior to the study start date. The primates were housed in individual micro-isolator cages. The monkeys were anaesthetized with a ketamine/valium mixture and infected intranasally and intratracheally with hMPV/NL/1/00 or RSV A2. On day 1, each animal received a dose of 2 ml containing 1·3x105 p.f.u. hMPV ml–1 or 3·5x105 p.f.u. RSV ml–1. Nasopharyngeal (NP) swabs were collected daily for 11 days and tracheal lavage (TL) specimens were collected on days 1, 3, 5, 7 and 9 post-immunization. Blood samples for serological assays were collected on days 1, 7, 14, 21 and 28. The animals were monitored for body temperature changes indicating a fever, signs of a cold, runny nose, sneezing, loss of appetite and change in body mass. hMPV or RSV present in the primate NP and TL specimens was quantified by plaque assay using Vero cells. Mean peak virus titres represent the mean of the peak virus titre measured for each animal on any of the 11 days following immunization.

Plaque reduction neutralization assay.
PRNAs were carried out for sera obtained on days 1 and 28 post-infection from hamsters and primates infected with hMPV/NL/1/00 or hMPV/NL/1/99. The animal sera were serially twofold diluted and incubated with 100 p.f.u. hMPV in the presence of guinea pig complement for 1 h at 4 °C. The virus/serum mixtures were transferred to Vero cell monolayers and overlaid with OPTI-MEM containing 1 % methylcellulose. After 6 days of incubation at 37 °C, the monolayers were immunostained using hMPV ferret polyclonal antiserum for quantification. Neutralization titres were expressed as the reciprocal log2 of the highest serum dilution that inhibited 50 % of virus plaques.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Propagation of hMPV in tissue culture
hMPV was isolated from clinical samples derived from Dutch children who were negative for RSV yet displayed severe respiratory distress (van den Hoogen et al., 2001). Respiratory samples from these children were inoculated in tertiary monkey kidney (tMK) cells and hMPV was identified. tMK cells are not practical for routine propagation of hMPV and therefore a number of immortal cell lines were tested for support of hMPV replication. Initially only the growth of hMPV/NL/1/00, the A1 prototype, was tested in a number of different cell lines such as Vero, HEp-2 and LLC-MK2 cells. Vero and LLC-MK2 cells supported hMPV/NL/1/00 replication to titres of 106–107 p.f.u. ml–1. When replication of hMPV/NL/1/99, the B1 prototype, was evaluated in Vero cells, similar titres of 106–107 p.f.u. ml–1 were achieved. Neither virus had a trypsin requirement for propagation, which is unlike other hMPV strains. Representatives of subgroup A2, such as hMPV/NL/17/00, or subgroup B2, such as hMPV/NL/1/94, displayed a stringent requirement for trypsin in the culture medium for growth in Vero cells. The trypsin dependence of hMPV subgroups A2 and B2 may have a correlate in animal models, although this was not observed for parainfluenza viruses. hPIV1 is trypsin dependent for growth in cell culture, but this phenotype does not influence its replication properties in vivo in the respiratory tract of hamsters, where virus titres similar to those for the non-trypsin-dependent hPIV3 and hPIV2 strains were achieved (Tao et al., 1998). hMPV did not replicate efficiently in HEp-2 cells, a cell substrate commonly used to grow RSV, or fertilized hens' eggs employed to propagate influenza viruses. The cytopathic effects observed in hMPV-infected Vero cell monolayers after 10 days of incubation at 35 °C were subtle and involved only a modest degree of syncytium formation and cell rounding (data not shown). Immunostaining of infected Vero cells using hMPV polyclonal antiserum revealed small plaques with limited spread and small syncytia for both hMPV subgroups. Virus stocks for hMPV subgroups A and B were generated in Vero cells and used for the development of small-animal and primate models.

Identification and characterization of small-animal models supporting hMPV replication in the respiratory tract
To identify a small-animal model that will support hMPV replication in the respiratory tract, hamsters, mice, cotton rats and ferrets, previously used to study other paramyxoviruses, were investigated. Small animals have been employed as models to screen attenuation phenotypes of live virus vaccines as well as to evaluate the immunogenicity elicited by the virus vaccine candidates. Syrian golden hamsters have been used to characterize live attenuated PIV3 vaccine candidates (Haller et al., 2000; Skiadopoulos et al., 1999). Live attenuated candidate RSV vaccines have been studied in cotton rats and BALB/c mice (Jin et al., 2000). Ferrets have been employed to study attenuation phenotypes of live influenza virus vaccines (Maassab et al., 1982). Therefore, these animals were chosen initially to study their permissiveness to hMPV infection. The animals were dosed intranasally with 106 p.f.u. hMPV, and 4 days post-infection the nasal turbinate and lung tissues were assayed for virus replication. The results showed that Syrian golden hamsters and ferrets supported hMPV replication in the respiratory tract to high titres. As shown in Table 1, hMPV titres of 5·3 and 4·3 log10 p.f.u. (g tissue)–1 in the URT and LRT, respectively, of hamsters were observed. Ferret nasal and lung tissues yielded hMPV titres of 4·7 and 4·0 log10 p.f.u. (g tissue)–1, respectively. The body temperature of the ferrets was monitored daily but the animals did not develop a fever during the course of virus infection. Signs of illness such as a cold, runny nose, sneezing and loss of appetite were not observed for either hamsters or ferrets. Both hamsters and ferrets developed neutralizing hMPV antibodies ranging in titre from 3 to 8 reciprocal log2 in the individual animals 4 weeks post-infection (data not shown). In contrast, BALB/c mice displayed hMPV replication titres of only 3·4 and 2·4 log10 p.f.u. (g tissue)–1 in the URT and LRT, respectively (Table 1). hMPV replication was not observed in either nasal turbinate or lung tissue of infected cotton rats (Table 1), which is in contrast to RSV. To study further the kinetics of hMPV replication in vivo, a time course was carried out in hamsters for 6 days (Table 2). Days 3 and 4 post-infection displayed the highest levels of hMPV replication. hMPV titres of 4·5–5·7 log10 p.f.u. (g tissue)–1 in the URT and 4·3–4·8 log10 p.f.u. (g tissue)–1 in the LRT were observed. The hamsters appeared to clear the virus infection by day 6. Hamsters are more cost-effective than ferrets and immunological reagents for hamsters are more readily available than for ferrets. Therefore, Syrian golden hamsters were chosen as a small-animal model to analyse further the immune response as well as replication of hMPV from other subgroups.


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Table 1. Replication of hMPV/NL/1/00 in the respiratory tracts of mice, cotton rats, hamsters and ferrets

 

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Table 2. Time course of hMPV/NL/1/00 replication in the respiratory tract of Syrian golden hamsters

 
hMPV from subgroups A and B can provide cross-challenge protection in hamsters
In order to design appropriate hMPV vaccine candidates, it was necessary to determine whether hMPV subgroup A can generate an immune response that will not only protect from subgroup A but also from subgroup B infection.

To study whether immunization with hMPV from subgroup A can protect from subsequent subgroup B infection, cross-challenge studies were carried out in hamsters (Table 3). The animals received 105 p.f.u. hMPV/NL/1/00 or hMPV/NL/1/99 intranasally. Four days later, six animals were sacrificed and virus titres in the nasal turbinate and lung tissue were determined. Hamsters that received hMPV/NL/1/00 displayed titres of 5·7 and 4·8 log10 p.f.u. (g tissue)–1 in the URT and LRT, respectively. The animals that were infected with hMPV/NL/1/99 replicated to titres of 4·2 and 5·0 log10 p.f.u. (g tissue)–1 in the URT and LRT, respectively (Table 3). Separate groups of hamsters immunized with hMPV/NL/1/00 or hMPV/NL/1/99 were challenged on day 28 post-infection with both hMPV/NL/1/00 and hMPV/NL/1/99. The results showed that animals vaccinated with hMPV/NL/1/00 were completely protected from both hMPV subgroups A and B. Similarly, hamsters that received hMPV/NL/1/99 were completely protected from challenge with hMPV/NL/1/00 or hMPV/NL/1/99 (Table 3). Only the animals that were administered placebo medium showed high levels of ~5 log10 p.f.u. (g tissue–1) of hMPV subgroup A or B replication in the URT and LRT (Table 3). This result demonstrated that both hMPV subgroups A and B have the ability to replicate to high titres in the respiratory tract of hamsters if the dose is >105 p.f.u. In hamsters, infection with subgroup A hMPV produced an immune response that conferred protection from subsequent subgroup B infection. Similarly, hMPV subgroup B-immunized hamsters were protected from subgroup A challenge.


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Table 3. hMPV from subgroups A and B can provide cross-challenge protection in hamsters

 
Prior to challenge on day 28, serum was collected from the vaccinated hamsters and analysed for the presence of subgroup A and B neutralizing hMPV antibodies using a 50 % plaque reduction neutralization assay. Animals immunized with hMPV/NL/1/00 produced neutralizing antibody titres of 8·4 log2 to the homologous antigen (hMPV/NL/1/00) and 4·4 log2 to the heterologous antigen (hMPV/NL/1/99) (Table 4). Hamsters that were administered the subgroup B hMPV displayed antibody titres of 8·6 log2 for the homologous antigen (hMPV/NL/1/99) and 4·2 log2 for the heterologous antigen (hMPV/NL/1/00). Despite the 4 log2-reduced antibody titres for the hMPV/NL/1/99 antigen present in sera obtained from hMPV/NL/1/00-infected hamsters, the animals were completely protected from hMPV/NL/1/99 challenge (Table 3). Similar results were observed for the hMPV/NL/1/99-infected hamsters that displayed decreased subgroup A neutralizing antibody titres but were protected from the NL/1/00 challenge virus.


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Table 4. hMPV neutralizing antibodies specific for subgroups A and B elicited in hMPV-infected hamsters

 
Replication of hMPV in the respiratory tract of rhesus monkeys and AGMs
Evaluation of live or killed vaccines, mAbs or antivirals in primates for efficacy is often necessary to support licensing of new biological products. Two species of non-human primates were tested for their ability to support hMPV replication in the respiratory tract. Rhesus monkeys have been used previously to evaluate hPIV3 vaccine candidates such as bovine PIV3 and cp-45 hPIV3 (Pennathur et al., 2003; Karron et al., 2003). AGMs have been employed to study safety and immunogenicity of a number of live attenuated RSV vaccine candidates (Jin et al., 2003). Chimpanzees have also been shown to support replication of both PIV3 and RSV (van Wyke Coelingh et al., 1988; Teng et al., 2000); however, their scarcity prohibited their evaluation as an hMPV primate model in this study.

Briefly, hMPV-seronegative rhesus monkeys were infected intratracheally and intranasally with a 1 ml dose at each site containing 1·3x105 p.f.u. hMPV. Virus shedding was monitored daily for 11 days post-infection in the nasopharynx and on days 1, 3, 5, 7 and 9 post-infection in the trachea. From the four rhesus monkeys that received hMPV/NL/1/00, only a single animal showed a virus titre of 4·0 log10 p.f.u. ml–1 in the URT (Table 5). This animal shed virus for 5 days in the URT and for only 3 days in the LRT where hMPV titres of 1·8 log10 p.f.u. ml–1 were observed. The other three rhesus monkeys displayed very low levels of virus shedding of 1·3–1·8 log10 p.f.u. ml–1 in the URT and titres ranging from 1·3 log10 p.f.u. ml–1 to below the assay detection limit in the LRT. The mean hMPV peak replication titre in the nasopharynx of rhesus monkeys was 2·2 log10 p.f.u. ml–1 and 1·3 log10 p.f.u. ml–1 in the trachea (Table 5). Neutralizing serum antibody titres were not determined, since only low levels of replication were observed.


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Table 5. Replication of hMPV/NL/1/00 in the respiratory tract of rhesus macaques and AGMs

 
The design of the AGM study was identical to that of the rhesus macaque study, with the exception that an RSV A2 control group was included. hMPV replicated more efficiently in the respiratory tract of AGMs. Mean peak titres of 3·7 log10 p.f.u. ml–1 and 5·0 log10 p.f.u. ml–1 were observed for the URT and LRT of AGMs, respectively (Table 5). hMPV was shed for 7–11 days in the nasopharynx and for 8–9 days in the trachea of AGMs. In comparison, RSV A2 displayed mean peak titres of 3·3 log10 p.f.u. ml–1 in the nasopharynx and 5·0 log10 p.f.u. ml–1 in the trachea of AGMs (Table 5). However, in this study, one animal did not shed high titres of RSV in the URT, which lowered the mean replication in the trachea. RSV was shed for 2–9 days in the nasopharynx and 7–9 days in the trachea of AGMs (Table 5). These results showed that RSV and hMPV replicated to similar titres in the URT and LRT of AGMs, although RSV may replicate to slightly higher titres than hMPV in the URT since two of three animals displayed titres of 4 log10 p.f.u. ml–1 and only the third animal failed to shed high levels of RSV. Disease signs such as a fever, runny nose, sneezing and loss of appetite were not observed for either rhesus monkeys or AGMs.

The kinetics of hMPV replication in the URT and LRT during the course of infection was compared with that observed for RSV (Fig. 1). The progress of infection for two animals each infected with hMPV or RSV in the URT showed that virus shedding lasted for >7 days post-infection. The peak of virus replication in the URT occurred between days 4 and 5 for hMPV and days 6 and 8 for RSV A2 (Fig. 1A) Virus shedding in the LRT showed that hMPV reached peak titres between 5 and 7 days post-infection and RSV displayed peak titres on day 5 post-infection with similar time periods of virus shedding (Fig. 1B).



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Fig. 1. (A) Kinetics of URT hMPV and RSV A2 infection in AGMs. Nasopharyngeal swab (N) samples were collected daily for 11 days post-infection and analysed by plaque assay on Vero cells. hMPV achieved peak titres between days 4 and 5 in the URT, while RSV replication peaked between days 6 and 8. (B) hMPV and RSV infection in the LRT of AGMs. Bronchoalveolar lavage (B) samples were collected on days 1, 3, 5, 7 and 9 post-infection and analysed by plaque assay on Vero cells. Peak titres for hMPV and RSV replication were observed on day 5 post-infection in the LRT. The duration of virus replication and times of peak titre achievement for hMPV/NL/1/00 and RSV A2 were similar in the URT and LRT of AGMs.

 
AGM sera obtained on day 28 were analysed for the presence of hMPV subgroup A and B neutralizing antibody titres (Table 6). All three animals displayed high neutralizing antibody titres with a mean reciprocal titre of 9·1 log2 when tested using the hMPV/NL/1/00 subgroup A antigen and 8·4 log2 when using the subgroup B antigen (Table 6). Therefore, AGMs infected with hMPV subgroup A produced neutralizing antibody titres that cross-neutralized subgroup B hMPV, although the antibody titres using the homologous antigen were slightly higher.


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Table 6. hMPV/NL/1/00-infected AGMs produce neutralizing antibodies specific for subgroups A and B of hMPV

 

   DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
The goal of this study was the identification of small-animal and primate models that would support hMPV replication in the respiratory tract for evaluation of future hMPV vaccines, mAbs and antivirals, and to determine whether a vaccine based on a single hMPV subgroup will elicit an immune response sufficient to protect animals from subsequent infections with different hMPV subgroups.

Therefore, a number of small-animal and primate models were evaluated for permissiveness to hMPV infection. Animal models currently used to evaluate RSV or hPIV3 vaccines or mAbs support high levels of RSV or PIV3 replication in the respiratory tract. Usually virus titres of 104–105 p.f.u. ml–1 are observed in the URT and LRT of RSV or PIV3-immunized animals (Haller et al., 2000; Jin et al., 2000, 2003; Pennathur et al., 2003). High levels of virus replication in the animal models are desirable for the following reasons: (i) The degree of attenuation of vaccine strains can be better measured. Reduced replication in the respiratory tract of animals should correlate with reduced disease in humans. (ii) High levels of virus replication in the animals should stimulate strong cellular and humoral immune responses and elicit high neutralizing antibody titres, in general a correlate of immune protection against respiratory virus infections. (iii) Immune protection of vaccinated animals will result in a marked reduction in challenge virus titres (>2 log10) demonstrating the effectiveness of the vaccine.

Four small-animal models, mice, hamsters, cotton rats and ferrets, were tested for permissiveness to hMPV infection, replication in the respiratory tract and induction of neutralizing antibodies. The results showed that Syrian golden hamsters and ferrets supported hMPV replication to high titres of 4–5 log10 p.f.u. ml–1 in the URT and LRT. None of the animals showed signs of hMPV disease. Hamsters appeared to clear most of the hMPV infection by day 6 post-infection. Hamsters and ferrets produced a neutralizing antibody response, displaying titres ranging from 3 to 8 log2 for the individual animals. BALB/c mice were semi-permissive for hMPV infection and cotton rats did not display detectable hMPV titres in the nasal turbinates or lungs 4 days post-infection.

Two primate models, rhesus macaques and AGMs, were evaluated as potential hMPV primate models. Rhesus monkeys were not very permissive for hMPV infection and did not display high hMPV titres in the URT and LRT, even though a dose of >105 p.f.u. was administered intranasally and intratracheally. Only one of four animals displayed an hMPV titre of 4 log10 p.f.u. ml–1 in the URT; the other three animals shed only low levels of hMPV. hMPV replication in the LRT of rhesus monkeys was not detected in two animals and was very low (1·3 and 1·8 log10 p.f.u. ml–1) in the other two animals. In contrast, hMPV mean replication titres of 3·7 and 5·0 log10 p.f.u. ml–1 were observed in the nasopharynx and trachea of AGMs, respectively. The results obtained for hMPV replication in rhesus monkeys and AGMs were similar to those observed for RSV. It has previously been demonstrated that cynomolgus macaques are only semi-permissive for RSV infection, while AGMs support high levels of RSV replication (A. Haller, unpublished observation; Jin et al., 2003). In AGM sera collected 28 days post-hMPV/NL/1/00 infection, mean hMPV neutralizing antibody titres of 9 log2 were observed for the homologous antigen. Slightly lower neutralizing antibody levels were observed when the heterologous virus (hMPV/NL/1/99) was used in the neutralization assay. Similar levels of RSV neutralizing antibody titres have been observed for sera obtained from RSV-infected AGMs (Jin et al., 2003). It is expected that AGMs will also be permissive for hMPV subgroup B infection. Neither rhesus monkeys nor AGMs displayed signs of hMPV disease. Therefore, AGMs did not mimic the hMPV infection observed in humans. The observation that sera obtained from hMPV-infected AGMs contained similar levels of subgroup A and B neutralizing antibody titres suggested that the immune systems of non-human primates and hamsters are different in nature. Hamsters appeared to produce more hMPV subgroup-specific neutralizing antibody classes, while AGMs elicited antibodies that neutralized both subgroups equally well.

Syrian golden hamsters were used to study whether infection with hMPV subgroups A or B could generate an immune response that would cross-protect. These findings have implications on vaccine design and the number of vaccine strains necessary for immune protection from wild-type hMPV infection. The results showed that hamsters vaccinated with hMPV/NL/1/00 (subgroup A) were protected from challenge with hMPV/NL/1/99, a subgroup B representative. Similarly, hamsters that had received a subgroup B virus (hMPV/NL/1/99) were protected from challenge with a subgroup A virus (hMPV/NL/1/00). These results suggest that an hMPV vaccine based on either the subgroup A or subgroup B genetic backbone can provide sufficient protection from infection by other hMPV subgroups. This is an important finding that will facilitate rational vaccine design, since future hMPV vaccines may not need to contain both hMPV subgroups to be efficacious. hMPV subgroup cross-protection ability was likely due to the highly conserved hMPV F protein, although some contributions may stem from the less conserved hMPV G protein. The hMPV F protein is a viral surface glycoprotein and is thought to be responsible in part for eliciting the neutralizing antibody response. Although the hMPV neutralizing antibody titres for the heterologous hMPV antigen were lower by as much as 4 log2, the neutralizing antibodies induced effectively neutralized and protected the hamsters from the heterologous hMPV challenge virus. Based on the importance of cell-mediated immunity for humans to recover from paramyxovirus infections, it is likely that T cell responses to hMPV also contributed to the high level of heterologous protection observed after hMPV challenge.

Similar observations have been made for RSV, a related paramyxovirus. For RSV, subgroup A can provide immunological protection from subgroup B (Jin et al., 2003). Therefore, an RSV vaccine based on subgroup A should be sufficient to protect from RSV subgroup B. The RSV F proteins of subgroups A and B are highly conserved and display an 89 % amino acid identity, while the G proteins are divergent and only show 53 % identity. In contrast, hPIV3 will not induce an immune response that will protect from hPIV1 infection (Tao et al., 2000). The lack of immune cross-protection is most likely due to a greater degree of divergence of the surface glycoproteins of hPIV3 and hPIV1. The F and haemagglutinin–neuraminidase (HN) proteins of hPIV3 and hPIV1 display amino acid identities of only 41·9 and 35·5 %, respectively. These results suggest that immune cross-protection requires at least one highly conserved viral surface glycoprotein among the different virus strains to elicit cross-reactive neutralizing antibodies. The high degree of hMPV F protein conservation suggests that antibodies directed against the hMPV F protein should neutralize both subgroups A and B of hMPV, which is supported by the results obtained from hamsters presented in this study.

In summary, this study identified two small-animal models, hamsters and ferrets, that supported efficient hMPV replication in the respiratory tract and produced high hMPV neutralizing antibody titres. The hamster studies showed that hMPV subgroups A and B could elicit cross-subgroup immune protection and this finding will influence rational vaccine design. AGMs were also shown to be permissive for hMPV infection, and high levels of hMPV replication were observed in the respiratory tract. Furthermore, hMPV infection of AGMs induced a high level of hMPV neutralizing antibodies. These results will have an important impact on future hMPV vaccine design and facilitate evaluation of hMPV vaccines, mAbs and antivirals.


   ACKNOWLEDGEMENTS
 
We thank Leenas Bicha and Fiona Fernandes for technical assistance with the animal sample analyses and MedImmune Vaccines' Animal Care Facility staff for their technical support with the small-animal studies. We thank Ken Draper and Brad Saville from Sierra Biomedical for advice on the primate studies.


   REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
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Received 13 November 2003; accepted 13 February 2004.