Understanding foot-and-mouth disease virus transmission biology: identification of the indicators of infectiousness
© Chase-Topping et al.; licensee BioMed Central Ltd. 2013
Received: 18 February 2013
Accepted: 5 June 2013
Published: 3 July 2013
The control of foot-and-mouth disease virus (FMDV) outbreaks in non-endemic countries relies on the rapid detection and removal of infected animals. In this paper we use the observed relationship between the onset of clinical signs and direct contact transmission of FMDV to identify predictors for the onset of clinical signs and identify possible approaches to preclinical screening in the field. Threshold levels for various virological and immunological variables were determined using Receiver Operating Characteristic (ROC) curve analysis and then tested using generalized linear mixed models to determine their ability to predict the onset of clinical signs. In addition, concordance statistics between qualitative real time PCR test results and virus isolation results were evaluated. For the majority of animals (71%), the onset of clinical signs occurred 3–4 days post infection. The onset of clinical signs was associated with high levels of virus in the blood, oropharyngeal fluid and nasal fluid. Virus is first detectable in the oropharyngeal fluid, but detection of virus in the blood and nasal fluid may also be good candidates for preclinical indicators. Detection of virus in the air was also significantly associated with transmission. This study is the first to identify statistically significant indicators of infectiousness for FMDV at defined time periods during disease progression in a natural host species. Identifying factors associated with infectiousness will advance our understanding of transmission mechanisms and refine intra-herd and inter-herd disease transmission models.
Foot-and-mouth disease virus (FMDV), a member of the Aphtho virus genus within the Picornaviridae family, is the causative agent of foot-and-mouth disease (FMD), one of the world’s most important infectious animal diseases, responsible for huge global losses of livestock production and trade, as well as frequent and highly disruptive large-scale epidemics [1, 2]. The disease is characterised by a short lasting fever, epithelial lesions on the tongue, dental pad and inner mouth area leading to excessive salivation and drooling and lesions on the feet causing lameness. Secondary infection of epithelial lesions can significantly increase the severity of disease [3, 4].
There are seven immunologically distinct serotypes and more than 60 antigenic variations [5, 6] and many are endemic in large parts of Asia, Africa and South America . Here, we focus on serotype O, which is the most prevalent serotype globally and shown to be transmitted by several different routes. One of the most common routes of transmission in ruminants is by direct contact between infected and naïve animals. Indirect contact also occurs by mechanical transfer via people, wild animals and birds, vehicles, fomites and animal products e.g. milk or meat products [8–13]. The virus may also spread by inhalation of infectious droplets and droplet nuclei originating mainly from the breath of infected animals  which can be wind borne . Wind borne transmission occurs infrequently, as it requires particular climatic and epidemiological conditions [16–18].
A recent publication  reported the results of experimental studies of direct FMDV transmission in cattle. The results of that study suggested that conditions promoting transmission exist for only a brief period and showed that infectiousness is a complex phenomenon related not just to virus dynamics but also to host responses and clinical signs, which is consistent with a common but rarely tested expectation that disease signs may be functionally linked to infectiousness. Prior to this research, studies into FMDV transmission had used proxy measures for infectiousness (for example the detection of virus in the blood or other tissues) rather than directly demonstrating transmission to another animal. Recent results  highlighted that cattle infected with FMDV are substantially less likely to be infectious before showing clinical signs than was previously realized. As such there is a need for more robust empirical evidence on relationships between clinical signs and infectiousness.
The aim of the present study was to utilize the relationship between the onset of clinical signs and direct contact transmission of FMDV to identify possible predictors of the onset of clinical signs as well as identify candidates for preclinical screening in the field. Such information will advance our knowledge of the transmission mechanisms and improve the model predictions that are used in disease control. The assumption that the likelihood of transmission is decreased if control can be implemented just 24 h earlier provides strong support for investment in the development of practical tools for preclinical diagnosis. If we can identify infected cattle before they show signs of disease using tests in the laboratory then perhaps these can be used in the field during an outbreak. Measures of concordance between qualitative real time (qRT)-PCR results and virus isolation results were also determined in each experiment. These measures of concordance are useful in evaluating the performance of both methods of virus detection.
Materials and methods
Details of the methods used in this paper have been published elsewhere  but are described in brief below. All experiments were approved by the Institute’s ethical review process and were in accordance with national guidelines on animal use.
Animal experiments and samples
Four individual animal experiments using 100–150 kg Holstein Friesian calves were performed. For each experiment, two animals (referred to later as inoculates) were selected at random, and were needle challenged intradermolingually (1) with 1 × 105.7 TCID50 of cattle adapted FMDV O UKG 34/2001. Forty eight hours after challenge naïve animals (“donors”, 2 animals for each of 4 experiments) were introduced to the inoculates and were challenged by direct contact exposure for 24 h. The inoculates were removed from the study and the animals exposed to infection (n = 8) were used to attempt transmission to further naive cattle (“recipients”) at two, four, six and eight (in experiments 3 and 4 only) days post infection for a period of 8 h each time. A total of 28 recipients were used in this study design.
Individual donors were examined daily for the presence of clinical signs (lesions in the mouth, tongue, snout, feet and the presence of nasal discharge). Rectal temperature was also recorded. Blood and nasal fluid samples were taken daily for the first 8 days following challenge and then every other day for up to 14 days after challenge. The samples were transferred immediately to the laboratory for processing; nasal fluid was stored at −80°C and heparinised blood aliquoted and stored for subsequent virus detection in bovine thyroid cells (BTY) culture as described earlier . Aliquots of serum were stored at −80°C for subsequent total antibody (Ab) detection, nucleic acid extraction and analysis by qRT-PCR. Samples of oropharyngeal fluid (OPF) were collected by probang cup from all the animals before challenge and thereafter from the donors daily for the first week and fourteen days after challenge. All probang samples were stored at −80°C for subsequent virus detection using BTY cell culture and genome detection using real-time quantitative PCR.
Several air samples using multiple devices were collected simultaneously, each hour, during all but 2 of the 28 challenge periods. Air samples were collected using an all-glass Cyclone sampler (operated for 5 min at a flow rate of approximately 390l/min) and an all glass Porton impinger sampler (operated for 5 min at a flow rate of 11l/min). These sampling periods were determined as the optimal sampling configurations for the instruments [20–22]. The collecting media employed in these samplers was modified eagle’s medium (MEM) -HEPES with antibiotics and 0.1% (w/v) BSA [16, 23]. The concentration of virus per litre of air was determined by endpoint titration for each particular air-sample, which was multiplied by the volume of the collecting fluid and the flow of the sampler. The amount of infectivity recovered was expressed as the total amount (50% tissue culture infectious dose, TCID50) of airborne FMDV per animal per challenge period (8 hrs).
Live virus was detected in the biological samples collected (heparinised blood, nasal swabs, OPF) and in the collecting media from the air samples using BTY primary cell cultures [16, 23, 24]. Given the large number of samples taken, they were first screened to determine the presence of virus, and then a tenfold dilution series of those verified to be virus positive were made and each dilution inoculated onto five BTY tubes. Titres were calculated by the Karber equation according to Lennette . The specificity of any cytopathic effect was confirmed by an antigen capture ELISA [26–28].
Viral nucleic acid purification
RNA (200 μL of sample, mixed with 300 μL of MagNA Pure LC total nucleic acid Lysis/Binding Buffer) was extracted using the MagNA Pure LC total nucleic acid isolation kit (Roche, UK) and an automated nucleic acid robotic workstation according to the manufacturer’s instruction (MagNA Pure LC, Roche, UK). The samples were eluted in a volume of 50 μL and stored at −80°C for later analysis.
FMDV RNA standards and reverse transcription
As these experiments were performed with the O UKG 34/01 isolate, a homologous RNA standard was used (synthesized in vitro from a plasmid containing a 500 base pair insert of the internal ribosomal entry site (IRES) of FMDV O UKG 34/01) as described by Quan et al. . The reverse transcription was performed as previously described [29–31].
To determine the amount of FMDV RNA in extracts of the total nucleic acid from blood, OPF, and nasal fluid a qRT-PCR was performed according to the methodology previously described [30, 32]. In the PCR reaction, primers SA-UK-IRES-248F (50-AAC CAC TGG TGA CAG GCT AAG G-30)/SA-UK-IRES-308R (50-CCG AGT GTC GCG TGT AC CT-30) and a UK-IRES- 271T (6-FAM-TGC CCT TTA GGT ACC C-MGB) TaqMan® Minor Groove Binding probe (Applied Biosystems) were used, as this primer/probe set was designed for optimum detection of the FMDV O UKG 2001 virus . The PCR was performed on a Stratagene® MX3005P™ QPCR system using MXPro-MX3005 v 3.00 Build 311 software (Stratagene, UK), and fifty PCR cycles were carried out. Once obtained, results and amplification plots were analysed and standard curves constructed from cycle threshold values [32–34] of the RNA standard dilutions, to provide a measure of the number of FMDV genome copies.
Assay for FMDV specific antibodies
Assay for interferon detection
Type-I interferon (IFN) biological activity was measured in serum samples from donor animals by using an Mx/chloramphenicol acetyltransferase (Mx/CAT) reporter gene assay .
Database management and statistical analysis
List of virological, immunological and environmental variables generated in this study and included in statistical analysis
Quantity of live virus in oropharyngeal fluid (OPF) (log10 TCID50/mL)
Quantity of FMDV genome copies in OPF (log10 copies/mL)
Quantity of live virus in the blood (log10 TCID50/mL)
Quantity of FMDV genome copies in blood (log10 copies/mL)
Type-1 interferon in serum (IU/mL)
Nasal fluid VI
Quantity of live virus in the nasal fluid (log10 TCID50/mL)
FMDV-specific antibodies detected in serum (titre/mL)
Total airborne FMDV per animal per challenge period
Only data up to and including day 8 were used in the statistical analyses as after day 8 data was not collected daily. In addition, all donor animals had exhibited clinical signs by day 8. To account for the repeated measures structure in the data (i.e. daily sampling) analyses using multiple explanatory variables were performed using a GLMM with covariance pattern model (Proc Glimmix SAS version 9.3, SAS Institute Inc., Cary, NC). The GLMM was fitted with a binomial distribution and a logit link function. The response variable was the presence or absence of any clinical signs. Donor was fitted as the sole random effect, other variables were fitted as fixed effects. Given the small sample size (n=8 donors) only single factor models were generated. In total 63 variables were analysed.
Diagnostics were performed and plots of residuals were examined, confirming the goodness-of-fit of each model. Odds ratios (OR) and their associated 95% confidence intervals were estimated in the final models for factors statistically significantly associated with the onset of clinical signs. The generalised chi-square/DF (χ2/DF) was used to compare the fit of different models. Unfortunately no significance test of model fit is available, however, the χ2/DF can identify models that given their fixed and random effect specifications more (or less) closely meet the specified distribution (binomial logit). The closer the χ2/DF is to unity the better the model and data meet the assumed residual distribution.
Virus in the blood and OPF was measured using both the virus isolation method (from heparinised blood samples) and the qRT-PCR method (from serum samples). The relationship between the two methods was examined by looking at the temporal trends using PROC LOESS (SAS version 9.3). LOESS is a nonparametric method for estimating local regression surfaces. In addition, agreement between the two methods was examined using Cohen’s kappa (StatXact v.8, Cytel Software Corp, Cambridge, MA, USA).
Preclinical predictors were identified in the data by examining the virological and immunological variables in the time frame surrounding the onset of clinical signs. Data from ±4 days from the onset of clinical signs for each donor animal were included.
Prior to analysis, it was specified that results with p <0.05 would be reported as exhibiting formal statistical significance.
Associations with transmission
As previously described  there were 28 attempts to transmit the disease from donor to recipient animals over the 8h periods, 8 (29%) of which were successful in transmitting disease. Six of these transmissions occurred on either day 4 or day 6 since exposure of the donor to the virus. Only one transmission event occurred on each of day 2 and day 8. One donor transmitted the disease on two occasions, days 4 and 6 post exposure to the virus. One cow failed to transmit FMDV, even though FMDV was detected in the nasal fluid (NF) and oesophageal-pharyngeal fluid (OPF), although not the blood.
Transmission was significantly associated (Fisher’s exact = 6.16; p = 0.021) with the onset of clinical signs (Figure 1). The peak at time 0, illustrates that for most cows transmission occurs on the same day as clinical signs appear. Only one transmission event occurred prior to the onset of clinical signs. However, this animal showed overt signs the next morning, approximately 16 h after the end of the successful challenge period. The clinical signs observed were varied: nasal discharge and lesions in the mouth or tongue were the most frequently reported “first” signs (Figure 1).
GLMM air sampling
Predictors of transmission and the onset of clinical signs
GLMM Onset of clinical signs
>5.0 log10 TCID50/mL
<5.0 log10 TCID50/mL
Onset OPF VI > CT
>6.5 log10 copies/mL
<6.5 log10 copies/mL
Onset OPF qRT-PCR > CT
>2.4 log10 TCID50/mL
<2.4 log10 TCID50/mL
Appear -1D blood VI > 0
Appear -2D blood VI > 0
Onset blood VI > CT
Onset -1D blood VI > CT
>4.6 log10 copies/mL
<4.6 log10 copies/mL
Appear -1D blood qRT-PCR > 0
Onset blood qRT-PCR > CT
Onset -1D blood qRT PCR > CT
Nasal fluid VI
>4 log10 TCID50/mL
<4 log10 TCID50/mL
Appear of nasal fluid VI > 0
Onset nasal fluid VI > CT
Type 1 IFN
Onset Type 1 IFN > CT
VI vs qRT-PCR measurements
In summary, virus is first detectable in the OPF, but detection of virus in the blood and nasal fluid may also be good candidates for preclinical indicators. Interestingly, the donor that did not transmit in this study never had any measurable amount of virus in the blood. However, virus was detectable in the OPF and nasal fluid.
Previous analysis of the experimental data used in this study showed that transmission was associated with onset of clinical signs . In this study we further characterize this relationship and look for predictors of the onset of clinical signs as a proxy for transmission, thus increasing the statistical power to identify indicators of infectiousness. This is important because experiments with large animals held in high containment facilities are challenging and, inevitably, it is only feasible to use a low number of replicates. Despite this, we were able to identify factors significantly associated with different stages of FMDV infection. However, the association between transmission and the onset of clinical signs implies that relying on the detection of clinical infections will not facilitate the removal of infected animals before they become infectious, so preclinical diagnosis is required to achieve this.
All immunological and virological variables measured (with the exception of total FMDV-specific antibodies) were positively associated with onset of clinical signs. This result is not surprising given that they were chosen a priori as useful measures of the transmission of FMDV . It does appear that onset of clinical signs only occur when virus levels exceed thresholds. This will surely be a useful measure for monitoring animals in further research programs and possibly in the field.
Air sample results were not included in previous analysis of this data  because problems associated with including missing data in an ordination analysis. This study, however, has shown that there is a significant association between virus detected in the air and transmission of FMDV. This does not mean that virus in the air is a vehicle of transmission in this study. We believe, however, that it is a marker for transmission. We note that the initial designs of this transmission study included an indirect transmission element (data not shown). Although airborne virus was detected in the air during the challenge periods no transmission occurred by this indirect route so further attempts at indirect transmission were not done. With such a small sample size the confidence limits are large, however, this possibly suggests that this is not a major route for disease spread between cattle, even though it appears to be an indicator of when an animal is infectious. Though the lack of airborne transmission might be due to the properties of this particular strain of virus as there was only a limited number of documented cases of airborne spread [40, 41]. Planned future research will include more rigorous air sampling as these results suggest that air sampling shows great promise as a predictor of transmission and may also prove to be useful to detect preclinical infection. Hand held devices have been developed and their feasibility for monitoring shedding of FMDV in cattle is being tested .
Temperature was a good indicator of the onset of clinical signs and of transmission . However, elevated temperatures do not occur early in the course of infection. Temperature is, of course, a non-specific clinical sign as such would have limited utility as a pre-clinical screening tool.
For OPF there was no difference between the two methods of virus detection. The levels recorded using qRT-PCR tended to be higher at the peak but for both methods there was perfect agreement with respect to detection of virus. Measurement of virus in the blood using the VI method resulted (in some cases) in low levels of virus being detected earlier. Virus was detectable by qRT-PCR (in some cases) even when it was not using the VI method, but this always occurred later in the course of infection. It is unknown whether the virus detected by qRT-PCR at these stages is inactivated or at such a low concentration that it is unable to be detected by virus isolation.
This analysis has confirmed the close association between the onset of clinical signs and the transmission of FMDV from an infected bovine reported previously . In addition, we have identified predictors of clinical signs, namely virus present in the OPF, blood or nasal fluid but, importantly, only above a measured threshold. We also find that transmission is strongly associated with detectable levels of virus in the air although this need not imply that air-borne spread is itself a major route of transmission.
It has been argued that early detection of FMDV infection was critical to effective control of outbreaks and could help remove the need for pre-emptive culling . As clinical signs appear very close to the onset of infectiousness these are not ideal indicators. Here, we report that the detection of virus in OPF provides the earliest indication; however, this is unlikely to be practicable in the field. Alternatively, detection of virus in the blood or nasal fluid is possible days before the appearance of clinical signs. In terms of early detection of infection, the VI method performs best but, in contrast to PCR methods, is not a good basis for a rapid penside test. In future work, given that we have demonstrated that virus can be detected early in the course of infection in OPF samples and nasal swabs, we intend to explore the possibility of developing more sensitive air sampling methods as the most obviously practicable approach to mass screening.
Paul V Barnett and Bryan Charleston are Jenner Institute investigators.
We thank Luke Fitzpatrick, Colin Randal and Mark Jenkins for their assistance with the handling and management of experimental animals. We also thank John Gloster, Eoin Ryan and Caroline Wright for their assistance with air sampling, Pip Hamblin and Phil Keel for help and advice with serology assays and David Paton for valuable advice on study design. The work was funded by the Biotechnology and Biological Sciences Research Council (grant ref. BBSB00549).
- Belsham GJ: Distinctive features of foot-and-mouth disease virus, a member of the picornavirus family: aspects of virus protein synthesis, protein processing and structure. Prog Biophys Mol Biol. 1993, 60: 241-260. 10.1016/0079-6107(93)90016-D.View ArticlePubMedGoogle Scholar
- Knowles NJ, Samuel AR: Molecular epidemiology of foot-and-mouth disease virus. Virus Res. 2003, 91: 65-80. 10.1016/S0168-1702(02)00260-5.View ArticlePubMedGoogle Scholar
- Sobrino F, Domingo E: Foot and mouth disease: Current Perspectives. 2004, Norfolk, UK: Horizon Bioscience/CRC PressGoogle Scholar
- Ryan E, Gloster J, Reid SM, Li Y, Ferris NP, Waters R, Juleff N, Charleston B, Bankowski B, Gubbins S, Wilesmith JW, King DP, Paton DJ: Clinical and laboratory investigations of the outbreaks of foot-and-mouth disease in southern England in 2007. Vet Rec. 2008, 163: 139-147. 10.1136/vr.163.5.139.View ArticlePubMedGoogle Scholar
- Haydon DT, Bastos AD, Knowles NJ, Samuel AR: Evidence for positive selection in foot-and-mouth disease virus capsid genes from field isolates. Genetics. 2001, 157: 7-15.PubMed CentralPubMedGoogle Scholar
- Knowles NJ, Samuel AR, Davies PR, Kitching RP, Donaldson AI: Outbreak of foot-and-mouth disease virus serotype O in the UK caused by a pandemic strain. Vet Rec. 2001, 148: 258-259.PubMedGoogle Scholar
- Rweyemamu M, Roeder P, Mackay D, Sumption K, Brownlie J, Leforban Y, Valarcher JF, Knowles NJ, Saraiva V: Epidemiological patterns of foot-and-mouth disease worldwide. Transbound Emerg Dis. 2008, 55: 57-72. 10.1111/j.1865-1682.2007.01013.x.View ArticlePubMedGoogle Scholar
- Alexandersen S, Brotherhood I, Donaldson AI: Natural aerosol transmission of foot-and-mouth disease virus to pigs: minimal infectious dose for strain O1 Lausanne. Epidemiol Infect. 2002, 128: 301-312.PubMed CentralPubMedGoogle Scholar
- Alexandersen S, Zhang Z, Reid SM, Hutchings GH, Donaldson AI: Quantities of infectious virus and viral RNA recovered from sheep and cattle experimentally infected with foot-and-mouth disease virus O UK 2001. J Gen Virol. 2002, 83: 1915-1923.View ArticlePubMedGoogle Scholar
- Donaldson AI, Alexandersen S: Relative resistance of pigs to infection by natural aerosols of FMD virus. Vet Rec. 2001, 148: 600-602. 10.1136/vr.148.19.600.View ArticlePubMedGoogle Scholar
- Ryan E, Mackay D, Donaldson A: Foot-and-mouth disease virus concentrations in products of animal origin. Transbound Emerg Dis. 2008, 55: 89-98. 10.1111/j.1865-1682.2007.01004.x.View ArticlePubMedGoogle Scholar
- Donaldson AI: Foot-and-mouth disease: the principal features. Irish Vet J. 1987, 41: 325-327.Google Scholar
- Donaldson AI: Risk of spreading foot and mouth disease through milk and dairy products. Rev Sci Tech. 1997, 16: 117-124.PubMedGoogle Scholar
- Sellers R, Gloster J: Foot-and-mouth disease: a review of intranasal infection of cattle, sheep and pigs. Vet J. 2008, 177: 159-168. 10.1016/j.tvjl.2007.03.009.View ArticlePubMedGoogle Scholar
- Gloster J, Freshwater A, Sellers RF, Alexandersen S: Re-assessing the likelihood of airborne spread of foot-and-mouth disease at the start of the 1967–1968 UK foot-and-mouth disease epidemic. Epidemiol Infect. 2005, 133: 767-783. 10.1017/S0950268805004073.PubMed CentralView ArticlePubMedGoogle Scholar
- Donaldson AI, Gibson CF, Oliver R, Hamblin C, Kitching RP: Infection of cattle by airborne foot-and-mouth disease virus: minimal doses with O1 and SAT 2 strains. Res Vet Sci. 1987, 43: 339-346.PubMedGoogle Scholar
- Donaldson AI, Gloster J, Harvey LD, Deans DH: Use of prediction models to forecast and analyse airborne spread during the foot-and-mouth disease outbreaks in Brittany, Jersey and the Isle of Wight in 1981. Vet Rec. 1982, 110: 53-57. 10.1136/vr.110.3.53.View ArticlePubMedGoogle Scholar
- Alexandersen S, Kitching RP, Mansley LM, Donaldson AI: Clinical and laboratory investigations of five outbreaks of foot-and-mouth disease during the 2001 epidemic in the United Kingdom. Vet Rec. 2003, 152: 489-496. 10.1136/vr.152.16.489.View ArticlePubMedGoogle Scholar
- Charleston B, Bankowski BM, Gubbins S, Chase-Topping ME, Schley D, Howey R, Barnett PV, Gibson D, Juleff ND, Woolhouse MEJ: Relationship between clinical signs and transmission of an infectious disease and the implications for control. Science. 2011, 332: 726-729. 10.1126/science.1199884.View ArticlePubMedGoogle Scholar
- Errington FP, Powell EO: A cyclone separator for aerosol sampling in the field. J Hyg (Lond). 1969, 67: 387-399. 10.1017/S0022172400041802.View ArticleGoogle Scholar
- May KR, Harper GJ: The efficiency of various liquid impinger samplers in bacterial aerosols. Br J Ind Med. 1957, 14: 287-297.PubMed CentralPubMedGoogle Scholar
- Amaral Doel CM, Gloster J, Valarcher J-F: Airborne transmission of foot-and-mouth disease in pigs: evaluation and optimisation of instrumentation and techniques. Vet J. 2009, 179: 219-224. 10.1016/j.tvjl.2007.09.010.View ArticlePubMedGoogle Scholar
- Gibson CF, Donaldson AI: Exposure of sheep to natural aerosols of foot-and-mouth disease virus. Res Vet Sci. 1986, 41: 45-49.PubMedGoogle Scholar
- Snowdon WA: Growth of foot-and mouth disease virus in monolayer cultures of calf thyroid cells. Nature. 1966, 210: 1079-1080. 10.1038/2101079a0.View ArticlePubMedGoogle Scholar
- Lennette EH: Diagnostic procedures for viral and rickettsial diseases. 1964, New York: American Public health associationGoogle Scholar
- Ferris NP: Selection of foot and mouth disease antisera by ELISA. Rev Sci Tech. 1988, 7: 331-346.Google Scholar
- Ferris NP, Dawson M: Routine application of enzyme-linked immunosorbent assay in comparison with complement fixation for the diagnosis of foot-and-mouth and swine vesicular diseases. Vet Microbiol. 1988, 16: 201-209. 10.1016/0378-1135(88)90024-7.View ArticlePubMedGoogle Scholar
- Hamblin C, Armstrong RM, Hedger RS: A rapid enzyme-linked immunosorbent assay for the detection of foot-and-mouth disease virus in epithelial tissues. Vet Microbiol. 1984, 9: 435-443. 10.1016/0378-1135(84)90064-6.View ArticlePubMedGoogle Scholar
- Quan M, Murphy CM, Zhang Z, Alexandersen S: Determinants of early foot-and-mouth disease virus dynamics in pigs. J Comp Pathol. 2004, 131: 294-307. 10.1016/j.jcpa.2004.05.002.View ArticlePubMedGoogle Scholar
- Zhang Z, Alexandersen S: Detection of carrier cattle and sheep persistently infected with foot-and-mouth disease virus by a rapid real-time RT-PCR assay. J Virol Methods. 2003, 111: 95-100. 10.1016/S0166-0934(03)00165-4.View ArticlePubMedGoogle Scholar
- Reid SM, Ferris NP, Hutchings GH, De Clercq K, Newman BJ, Knowles NJ, Samuel AR: Diagnosis of foot-and-mouth disease by RT-PCR: use of phylogenetic data to evaluate primers for the typing of viral RNA in clinical samples. Arch Virol. 2001, 146: 2421-2434. 10.1007/s007050170012.View ArticlePubMedGoogle Scholar
- Reid SM, Ferris NP, Hutchings GH, Zhang Z, Belsham GJ, Alexandersen S: Detection of all seven serotypes of foot-and-mouth disease virus by real-time, fluorogenic reverse transcription polymerase chain reaction assay. J Virol Methods. 2002, 105: 67-80. 10.1016/S0166-0934(02)00081-2.View ArticlePubMedGoogle Scholar
- Zhang Z, Murphy C, Quan M, Knight J, Alexandersen S: Extent of reduction of foot-and-mouth disease virus RNA load in oesophageal-pharyngeal fluid after peak levels may be a critical determinant of virus persistence in infected cattle. J Gen Virol. 2004, 85: 415-421. 10.1099/vir.0.19538-0.View ArticlePubMedGoogle Scholar
- Reid SM, Grierson SS, Ferris NP, Hutchings GH, Alexandersen S: Evaluation of automated RT-PCR to accelerate the laboratory diagnosis of foot-and-mouth disease virus. J Virol Methods. 2003, 107: 129-139. 10.1016/S0166-0934(02)00210-0.View ArticlePubMedGoogle Scholar
- Ferris NP: Selection of foot and mouth disease antisera for diagnosis. Rev Sci Tech. 1987, 6: 127-140.Google Scholar
- Ferris NP, Kitching RP, Oxtoby JM, Philpot RM, Rendle R: Use of inactivated foot-and-mouth disease virus antigen in liquid-phase blocking ELISA. J Virol Methods. 1990, 29: 33-41. 10.1016/0166-0934(90)90005-Z.View ArticlePubMedGoogle Scholar
- Fray MD, Mann GE, Charleston B: Validation of an Mx/CAT reporter gene assay for the quantification of bovine type-I interferon. J Immunol Methods. 2001, 249: 235-244. 10.1016/S0022-1759(00)00359-8.View ArticlePubMedGoogle Scholar
- Sing T, Sander O, Beerenwinkel N, Lengauer T: ROCR: visualizing classifier performance in R. Bioinformatics. 2005, 21: 3940-3941. 10.1093/bioinformatics/bti623.View ArticlePubMedGoogle Scholar
- Howey R, Bankowski B, Juleff J, Savill NJ, Gibson D, Frazakerley J, Charleston B, Woolhouse ME: Modelling the within-host dynamics of the foot-and mouth disease virus in cattle. Epidemics. 2012, 4: 93-103. 10.1016/j.epidem.2012.04.001.View ArticlePubMedGoogle Scholar
- Gloster J, Champion HJ, Mansley LM, Romero P, Brough T, Ramirez A: The 2001 epidemic of foot-and-mouth disease in the United Kingdom: epidemiological and meteorological case studies. Vet Rec. 2005, 156: 793-803.View ArticlePubMedGoogle Scholar
- Valarcher JF, Gloster J, Doel CA, Bankowski B, Gibson D: Foot-and-mouth disease virus (O/UKG/2001) is poorly transmitted between sheep by the airborne route. Vet J. 2008, 177: 425-428. 10.1016/j.tvjl.2007.05.023.View ArticlePubMedGoogle Scholar
- Christensen LS, Brehm KE, Skov J, Harlow KW, Christensen J, Haas B: Detection of foot-and-mouth disease virus in the breath of infected cattle using a hand-held device to collect aerosols. J Virol Methods. 2011, 177: 44-48. 10.1016/j.jviromet.2011.06.011.View ArticlePubMedGoogle Scholar
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