Abstract
The feasibility of using Flinders Technology Associates (FTA) cards for the molecular detection of avian metapneumovirus (aMPV) by reverse transcriptase-polymerase chain reaction (RT-PCR) was investigated. Findings showed that no virus isolation was possible from aMPV-inoculated FTA cards, confirming viral inactivation upon contact with the cards. The detection limits of aMPV from the FTA card and tracheal organ culture medium were 101.5 median ciliostatic doses/ml and 100.75 median ciliostatic doses/ml respectively. It was possible to perform molecular characterization of both subtypes A and B aMPV using inoculated FTA cards stored for up to 60 days at 4 to 6°C. Tissues of the turbinate, trachea and lung of aMPV-infected chicks sampled either by direct impression smears or by inoculation of the tissue homogenate supernatants onto the FTA cards were positive by RT-PCR. However, the latter yielded more detections. FTA cards are suitable for collecting and transporting aMPV-positive samples, providing a reliable and hazard-free source of RNA for molecular characterization.
Introduction
Avian metapneumovirus (aMPV) is a member of the subfamily Pneumovirinae in the family Paramyxoviridae (Pringle, Citation1998). It is recognized as the cause of turkey rhinotracheitis virus, a widespread disease of turkeys and chickens, and has been reported in some other avian species such as ducks, geese, guinea fowl, pheasants and pigeons (Gough et al., Citation1988; Catelli et al., Citation2001; Bennett et al., Citation2005; Toquin et al., Citation2006). The virus is considered to be associated with swollen head syndrome in broiler and broiler breeders (Cook et al., Citation1988; Jones et al., Citation1991; Gough et al., Citation1994; Cook, Citation2000), and egg production losses in laying turkeys and chickens (Hafez et al., Citation1990; Cook et al., Citation1996; Hess et al., Citation2004; Sugiyama et al., Citation2006). The acute upper respiratory tract infection in turkeys and chickens associated with aMPV is characterized by oculonasal discharge, coughing, sneezing, tracheal râles, foamy conjunctivitis and swelling of infraorbital sinuses as a result of the secondary infection (Gough & Jones, Citation2008). Reverse transcriptase-polymerase chain reaction (RT-PCR) has been established for detection of aMPV (Cavanagh et al., Citation1999). However, in some countries, due to lack of RT-PCR technology, expertise and economic factors, it is expedient to send the specimens for diagnosis to specialized laboratories in countries where expertise exists. Shipments of infectious material must comply with strict regulation in most of these countries and organisms must be inactivated by chemicals, such as phenol or formalin, before being transported (Snyder, Citation2002). However, chemically inactivated samples might not always prove efficient in terms of subsequent virus detection due to problems in nucleic acid extraction (Coombs et al., Citation1999). An alternative and safe form of transportation of infectious material is the use of Flinders Technology Associates (FTA) cards (Moscoso et al., Citation2004). FTA cards comprise cotton-based cellulose paper that, when treated with anionic detergents and buffer, provides a stable matrix for the immobilization of genomes for molecular characterization but free from the living host cells or organisms (Natarajan et al., Citation2000; Whatman, Citation2009). This technology simplifies the method of genome collection, storage, transportation and extraction, and consequently reduces the cost and time required (Mbogori et al., Citation2006). The ability to use the FTA card for detection of several avian pathogens has been shown previously, including infectious bronchitis virus (Moscoso et al., Citation2005), Newcastle disease virus (Perozo et al., Citation2006), infectious bursal disease virus (Moscoso et al., Citation2006; Purvis et al., Citation2006), Marek's disease virus (Cortes et al., Citation2009), avian influenza virus (AIV) (Abdelwhab et al., Citation2011), fowl adenovirus (Moscoso et al., Citation2007) and mycoplasmas (Moscoso et al., Citation2004).
To date, no information has been published on the use of FTA cards for detection of aMPV. In this study, using aMPV subtypes A and B, we investigated: the use of the FTA cards in inactivating aMPV (Experiment 1); validating the detection limits for aMPV-inoculated FTA cards (Experiment 2); and the effects of different storage temperatures on the detection of aMPV (Experiment 3). We also compared the aMPV detection using two sampling techniques: tissue impression smears directly onto the FTA cards, and the same tissues macerated, and subjected to a freeze–thaw process and the supernatant inoculated onto FTA cards (Experiment 4).
Materials and Methods
Viruses
aMPV subtype A and subtype B strains available at the Institute of Infection and Global Health, University of Liverpool, UK were propagated and titrated in chicken tracheal organ cultures (TOC) as described previously (Cook et al., Citation1976). The median ciliostatic doses (CD50/ml) were calculated by the method of Reed & Muench (Citation1938). The titres obtained for subtypes A and B were 104.56 CD50/ml and 104.51 CD50/ml respectively.
Flinders Technology Associates cards
FTA cards are a commercial product developed by Whatman Corporation (Fisher Scientific Limited, Loughborough, UK) designed to transport and store a variety of biological materials including viral and bacteriological samples and blood. Infectious material applied to the cards is inactivated on contact as the cells are lysed but the genome is preserved for molecular recognition (Whatman, Citation2009).
Experiment 1: confirmation of aMPV inactivation by FTA cards
One hundred microlitres of stock culture of aMPV subtype A or subtype B were spotted onto the matrix area of an FTA card and allowed to air-dry at room temperature for 1 h. The spotted areas of the FTA card were excised using sterile scissors and forceps. Each sample was placed in a bijou bottle containing 2 ml TOC medium (Eagle's serum-free minimum essential medium with glutamine and streptomycin [50 mg/ml] and penicillin [50 iu/ml]) and processed for virus isolation as described previously (Cook et al., Citation1976). Briefly, 100 µl from each sample of the suspension were aseptically inoculated into replicates of three tracheal rings. The TOCs were checked daily for up to 10 days post inoculation and each sample was passaged three times. An absence of ciliostasis indicates an absence of viable aMPV.
Experiment 2: detection limits of aMPV on FTA cards
To determine the sensitivity of the RT-PCR in detecting subtype A or subtype B aMPVs, serial 10-fold dilutions up to 10−7 were made from each subtype from the initial stock. For each dilution, 100 µl was applied to the matrix areas of the FTA cards, which were air-dried for an hour at room temperature away from direct light. For comparison, 300 µl of the same viral dilutions were subjected directly to RT-PCR (see below). The RT-PCR reactions were conducted for each sample to determine the lowest dilution at which viral RNA can be detected.
Experiment 3: effects of storage temperature on detection of aMPV on FTA cards
A volume of 100 µl stock culture of aMPV subtype A or subtype B was spotted onto the matrix circles of FTA cards. After air-drying as above, the cards were placed in sealed polythene bags and kept at three different temperatures: in a refrigerator, with temperature range 4 to 6°C; in an incubator at 25°C; or in an incubator at 41°C. On each of days 1, 5, 10, 15, 20, 25, 30, 40, 50 and 60, one of the FTA cards was removed for the detection of aMPV by RT-PCR.
Experiment 4: detection of aMPV subtypes from FTA cards versus tissue supernatants
To assess the different methods of sample submissions and to demonstrate any variations in aMPV detection, 90-day-old specific pathogen free chicks were hatched and reared at the Institute of Infection and Global Health, University of Liverpool, UK. Food and drinking water were provided ad libitum. At 7 days of age, chicks were randomly divided into three groups of 30 birds each. Group 1 was inoculated oculonasally with 100 µl (103.56 CD50/chick) aMPV subtype A virus, Group 2 was inoculated with 100 µl (103.51 CD50/chick) of aMPV subtype B virus, while Group 3 was inoculated with virus-free TOC supernatant. At 1, 3, 5, 7, 9 and 14 days post infection (d.p.i.), five birds from each group were humanely killed and tissues of the turbinate, trachea and lung were collected.
The five tissues were each applied directly onto FTA cards. Tissues of the same type were pooled. An impression smear was made by gently pressing the tissues onto the matrix area of the cards. The cards were labelled and air-dried as described above. The remaining tissues were pooled according to the type, group and day of sampling and were stored at −70°C until further use.
Tissues of the same type were pooled and manually macerated with sterile sand in a pestle and mortar, and 3 ml TOC medium. After three cycles of freeze–thawing and centrifugation, 100 µl each supernatant was spotted directly onto an FTA card and allowed to dry for 1 h at room temperature. The FTA cards prepared with tissue impression smears and those that were inoculated with processed tissue supernatants were all examined for presence of aMPV RNA by RT-PCR.
Sample preparation and aMPV RT-PCR
Samples impregnated onto the matrix of the FTA cards (described above) and those with impression smears of fresh tissues were excised using sterile scissors and forceps, were placed in bijou bottles containing 2 ml guanidinium thiocyanate (solution D) and were then stored at −20°C. For extraction of the RNA from TOC medium, 300 µl inoculated TOC medium was added to 300 µl solution D and this was processed in the same way as for FTA papers. The RNA was extracted using the guanidinium thiocyanate–phenol chloroform method as described previously (Chomczynski & Sacchi, Citation2006). The aMPV genome was detected by RT-nested PCR based on the attachment protein gene (G) as described before (Cavanagh et al., Citation1999; Ganapathy et al., Citation2005), which allowed the differentiation of aMPV subtypes, with amplicon sizes of 268 base pairs and 361 base pairs for subtypes A and B respectively.
Results
Experiment 1: confirmation of aMPV inactivation by FTA cards
No ciliostasis was detected in the TOCs inoculated with elutions from either aMPV subtype-inoculated matrix of FTA cards. In contrast, ciliostasis was detected within 5 to 7 days post inoculation in TOC rings that received the stock culture of subtype A or subtype B.
Experiment 2: detection limits of aMPV on FTA cards
The detection limits of RT-PCR for the subtype A and subtype B aMPVs, based on either serially diluted stock cultures in the TOC medium or equivalent viral numbers, onto the matrix of the FTA cards are presented in . Detection limits for subtypes A and B were equivalent to ? 101.56 CD50/ml and 101.51 CD50/ml respectively for the samples processed on the FTA cards. The detection limits of the subtypes processed from TOC medium were 1 log higher than those for the FTA cards.
Table 1. Detection limits of aMPV subtype A or subtype B on FTA cards.
Experiment 3: effects of storage temperature on detection of aMPV on FTA cards
The stability of viral genome on FTA cards was measured by performing RT-PCR on the stock virus spotted onto FTA cards and stored at different temperatures for up to 2 months. The viral genome of both subtypes A and B was detected when stored at 4°C for up to 60 days post storage. Subtype A RNA was detected only up to 1 day post storage at 25°C and 41°C. In contrast, subtype B RNA was detected up to 30 days post storage at 25°C and for up to 5 days post storage when stored at 41°C ().
Table 2. Stability of aMPV RNA on the FTA cards under differing storage temperatures.
Experiment 4: detection of aMPV subtypes from FTA cards versus tissue supernatants
No virus was detected in the uninfected birds. presents the results for the detection of aMPV by RT-PCR for the infected groups. In general, subtype A was detected less frequently than subtype B. Subtype A was detected up to 7 d.p.i. only, compared with subtype B that was detectable up to 14 d.p.i. (end of the experiment). For subtype A, there were marginally more frequent detections in the tissue impression smears. For subtype B there were no obvious differences with detection in the turbinates with either method of sampling; virus was detected at almost all sampling intervals, except at 5 d.p.i. where the virus was detected only in the tissue impression sample. For tracheal samples, there was an obvious difference between the methods, but for lungs there was a slightly longer detection period by the tissue impression method (up to 9 d.p.i.). Overall, for subtype A and subtype B infections, by either sampling method (impression smear or tissue supernatant), the highest level of detection was seen at 5 d.p.i.
Table 3. Comparison of detection of aMPV by RT-PCR on samples collected on FTA cards either as tissue impression smear or inoculated with tissue supernatant.
Discussion
Definitive diagnosis of aMPV infection requires identification of the virus in clinical material, which is most often achieved nowadays by RT-PCR. However, PCR technology is unavailable in some countries and requires transportation of samples in a safe way to specialist laboratories, with international shipment of samples needing high standards of biosafety procedures (Snyder, Citation2002), including for aMPV. In assessing the suitability of detecting aMPV subtype A or subtype B, or both, we are reporting the results of experiments on inactivation of aMPV on FTA cards, aMPV detection limits when the viruses are sampled on FTA cards, effects of different temperatures on storage of FTA cards containing aMPV, and a cross-comparison between two sampling methods (direct impression smear versus inoculation of tissue supernatant onto FTA cards).
The absence of ciliostasis of the TOCs inoculated with elution from aMPV subtype A or subtype B indicated no viable virus presence, confirming that the FTA card inactivates the aMPV viruses. The ability of FTA cards to inactivate other avian pathogens such as infectious bronchitis virus, Newcastle disease virus, infectious bursal disease virus, AIV and fowl adenovirus have been reported before (Moscoso et al., Citation2005, Citation2006, Citation2007; Perozo et al., Citation2006; Narayanan et al., Citation2010; Abdelwhab et al., Citation2011).
In our study, the detection limits for subtype A and subtype B on FTA cards were 101.56 CD50/ml and 101.51 CD50/ml respectively. These values were approximately 1 log lower than the detection limits of viruses in the TOC medium. These data are in agreement with a similar study on detection of multiple poultry respiratory pathogens on FTA cards (Awad et al., Citation2012). It appears that there could be a small loss of RNA in the process of inoculation onto the FTA card matrix, storage or extraction processes. However, such a loss would outweigh the benefit of aMPV detection, particularly for samples transported thousands of miles.
Ideally, for successful virus detection, the FTA card should preserve and protect against deteriorations of viral genome. The FTA card manufacturer states that the card could be used for sampling and transportation of RNA viruses and the FTA card can be stored for a short time at room temperature or for longer at −20°C or −70°C (Whatman, Citation2009). However, to date there is no published information of the effects of various environmental temperatures on the detection of aMPV by RT-PCR. Our study demonstrated that, following inoculation of FTA cards and storage in air-tight plastic bags at 4 to 6°C, subtype A and subtype B were both detected for up to 60 days. At 25°C aMPV subtype B was detected up to 30 days but subtype A for only 1 day. The reason for this discrepancy is unclear even though virus titres of the viruses were almost the same. When the FTA cards were stored at 41°C, mimicking summer temperature in some countries, the subtype A was again only detected for 1 day but the subtype B was detected up to 5 days post storage. This reflects poor preservation of aMPV RNA at this temperature. The decrease in the sensitivity could be due to the fact that RNA denatures over time and is faster at higher temperatures (Rogers & Burgoyne, Citation2000; Moscoso et al., Citation2005, Citation2006). As such, FTA cards intended for aMPV RT-PCR ideally need to be stored at as low a temperature as possible, preferably 4 to 6°C. RT-PCR detection of infectious bronchitis virus or infectious bursal disease virus in samples stored on FTA cards at −20°C, 4°C or 41°C indicates that the RNA was stable for at least 15 days (Moscoso et al., Citation2005, Citation2006). Newcastle disease virus was stable on the FTA card for at least 20 days at both room temperature and 4°C, and for 18 months at −20°C (Narayanan et al., Citation2010). AIV RNA was also stable on FTA cards for at least 5 months at room temperature (Abdelwhab et al., Citation2011). It appears that, other than the intrinsic nature of the viruses, the variations in the virus titres and differences in the PCR protocols may have influenced the outcome.
Experiment 4 was organized to answer field questions on the appropriate methods of sampling onto FTA cards for optimal detection. The question was whether it was better to do a tissue impression smear directly onto the FTA card or whether the tissues should be processed in the normal manner for virus isolation and freeze–thawed three times, before applying a drop of 100 µl onto the FTA cards. Our study showed that there were slightly better chances of detecting aMPV RNA when the sample supernatants were used compared with tissue impression smears. These findings suggest that with tissue impression smears there could be a reduced amount of RNA left on the FTA card. In contrast, for the tissue supernatant, it is likely that maceration and later freeze–thaw processes have allowed cells to rupture and release more viral RNA into the supernatant. In a study with AIV, RNA from swab samples from the field adsorbed to and extracted from FTA cards was detected with reduced sensitivity when compared with RNA directly extracted from swab fluids (Abdelwhab et al., Citation2011). Thus, it appears that to increase the likelihood of aMPV detection, it is beneficial to grind the tissues (a pool of five or more tissues), subject them to a freeze–thaw process, centrifuge (or allow to stand still) and apply the supernatant onto the FTA cards.
These findings reveal that aMPV could be detected on FTA card samples taken from the turbinate, trachea and lung. However, the length of time for which detection was possible varied according to the organ type, sampling time and variation between aMPV subtypes A and B. For example, with samples taken from the turbinate, aMPV subtype A and subtype B were detectable up to 7 d.p.i. and 14 d.p.i. respectively, and with those from the lung were positive up to 5 d.p.i. and 9 d.p.i. for aMPV subtype A and subtype B respectively. These indicate that subtype B virus can be detected for a longer period than subtype A (Aung et al., Citation2008), even though similar titres of the viruses were used. The highest detections of aMPV was recorded at 5 d.p.i and this is in agreement with previous work in broilers (Gharaibeh & Shamoun, Citation2012) and turkeys (Liman & Rautenschlein, Citation2007).
We conclude that FTA cards are suitable for collecting and transporting aMPV samples worldwide, but the cards must be stored at lower temperatures for optimal chances of detection.
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