journa l homepage: www.e lsev ier .com/ locate / jv i romet
evelopment and validation of a novel SYBR Green real-time RT-PCR assay forhe detection of classical swine fever virus evaluated on different real-time PCRlatforms
ester Josué Péreza,1, Heidy Díaz de Arcea,∗,1, Joan Tarradasb, Rosa Rosellb,c, Carmen Laura Pereraa,arta Munozb, Maria T. Fríasa, José Ignacio Nunezb, Llilianne Gangesb
Centro Nacional de Sanidad Agropecuaria (CENSA), San Jose de las Lajas, La Habana, Apdo. 10, CubaCentre de Reserca in Sanitat Aimal (CReSA), UAB Bellaterra, 08193 Barcelona, SpainDepartament d’Agricultura, Alimentación i Acció Rural de la Generalitat de Catalunya (DAR), Spain
rticle history:eceived 14 August 2010eceived in revised form 16 March 2011ccepted 23 March 2011vailable online 31 March 2011
eywords:
a b s t r a c t
Classical swine fever is a highly contagious viral disease that causes significant economic losses in pigproduction on a global scale. The rapid dissemination of the virus and the variability of the clinical signsmerit the development of swift and accurate classical swine fever virus (CSFV) detection methods, whichcan assist in disease control. The development and evaluation of a novel quantitative real-time RT-PCRassay for CSFV detection, based on SYBR Green coupled to melting curve analysis, is described. The ana-lytical and diagnostic performances of the method using two real-time PCR instruments were compared.
lassical swine fever virusYBR Green real-time RT-PCReal-time PCR instrument
The assay was specific and detected the major genotypes of CSFV. The limit of detection in cell culturemedium and serum was 0.1 TCID50/reaction, while in tissue homogenate for both platforms, it was 1TCID50/reaction. The limit of detection was 1, 10 and 102 gene copies/�L when nuclease-free water,serum and tissue homogenate, respectively, were used as sample matrices for both instruments. Theanalysis of 108 tissue homogenate and serum samples from animals infected with CSFV naturally andexperimentally and non-infected animals showed that the assay provided a highly sensitive and specific
e fev
method for classical swin
. Introduction
Classical swine fever (CSF) is a highly contagious viral diseasef domestic pigs and wild boar (Moennig et al., 2003) that causesajor losses in stock farming due to high mortality rates, compul-
ory pig slaughter policies and bans on trade of live pigs and pigroducts (Terpstra and de Smit, 2000). CSF is a notifiable disease tohe World Organization for Animal Health (OIE, 2008a).
Classical swine fever virus (CSFV) is a member of the genus Pes-ivirus, which also includes Bovine viral diarrhoea virus 1 (BVDV-1),ovine viral diarrhoea virus 2 (BVDV-2), Border disease virus (BDV)nd a fifth tentative species, pestivirus of the giraffe (Thiel et al.,
005). The pestivirus genome consists of a single plus-strand RNAontaining a single large open reading frame (ORF) flanked by twontranslated regions (UTRs). The ORF encodes a polyprotein ofpproximately 3900 amino acids, which is processed subsequently
by cellular and viral proteases into 12 mature proteins: four struc-tural (C, Erns, E1 and E2) and eight non-structural proteins (Npro,P7, NS2, NS3, NS4A, NS4B, NS5A and NS5B) (Meyers and Thiel,1996). The rapid dissemination of the CSFV, as well as the variabilityof the clinical signs described among animals, compel the adoptionof rapid and accurate methods as a basis for the implementation ofeffective control measures to prevent further spread of the disease(Moennig, 2000). The real-time RT-PCR method described belowprovides novel rapid means of virus detection in diagnostic labora-tories. The advantages of this approach over conventional RT-PCRmethods include an enhanced sensitivity, a large dynamic range, areduced risk of cross contamination, the possibility of scale up andthe potential for an accurate target quantitation (Hoffmann et al.,2009).
Recently, real-time RT-PCR assays based on the hydrolysisprobe format (Hoffmann et al., 2005; Liu et al., 2007; JamnikarCiglenecki et al., 2008; Cheng et al., 2008) have been successfully
used for an improved detection of CSFV. However, the possibilityof false-negative test results poses a substantial problem in diag-nostic TaqMan assays because a single point mutation within theprobe-binding site could prevent annealing of the probe and thesubsequent detection (Hughes et al., 2004; Pham et al., 2005; King
PAV-250 Centro de Investigación en Sanidad Animal, SpainMargarita CENSA, Cuba15 Cuban field isolatesa CENSA, CubaClinical samples from experimentally infected pigswith Kozlov reference strain
Subtype 1aNADL EU Reference Laboratory for CSF, GermanyOregon Centro de Investigación en Sanidad Animal, SpainSinger Centro de Investigación en Sanidad Animal, Spain
Subtype 1b Osloss EU Reference Laboratory for CSF, Germany
Non-determined7 field isolates from cattle, Cuba/2002–2003 CENSA, Cuba8 field isolates from cattle, Spain/1990–1994b CReSA/DAR, Spain
BVDV-2 New York Hipra laboratory, Spain
BDV
Genotype 1Frijters ID-DLO Laboratory, Netherland.Moredun EU Reference Laboratory for CSF, Germany137/4 Central Veterinary Laboratory of Weybridge,
United KingdomGenotype 3 Clinical samples from experimentally infected pigs
with Gifhorn reference strainEU Reference Laboratory for CSF, Germany
Genotype 45 BDV isolates from sheepc CReSA/DAR, Spain2 BDV isolates from sheep Neiker, SpainBDV isolated from pig Spain/2006d CReSA, Spain
TGEV Purdue 115 Pathobiology Laboratory, Gelph University, CanadaEMCV Field isolate CENSA, Cuba
eso2baofa
paHaa
2
2
ia
a Díaz de Arce et al. (2005).b Vega et al. (2000).c Vega et al. (2002).d Rosell et al. (2008).
t al., 2006; Belák, 2007). Therefore, alternative detection formats,uch as primer probe energy transfer (PriProET) (Liu et al., 2009)r SYBR Green coupled to melting curve analysis (Martínez et al.,008; Kong et al., 2009; Tam et al., 2009; Tan et al., 2009) haveeen suggested to detect RNA viruses showing a high genomic vari-bility. The main purpose of this study was therefore to develop,ptimise and evaluate a novel, quantitative real-time RT-PCR assayor CSFV detection based on SYBR Green coupled to melting curvenalysis.
Taking into account the fact that the type of thermocyclinglatform used to perform the real-time RT-PCR assay could biasnalytical and diagnostic test performances (Bentley et al., 2005;ayden et al., 2008), two different apparatus were used to comparessay results: an ABI PRISM 7500 (AP7500) (Applied Biosystems)nd a LightCycler 1.5 (LC1.5) (Roche).
. Materials and methods
.1. Viruses
The reference strains and field isolates used in this study, whichncluded CSFV, non-CSFV pestiviruses and other porcine viruses,re listed in Table 1.
2.2. Cell cultures and virus propagation
Viruses were cultured and titrated in PK15 (CSFV strains) andMDBK cell lines (BVDV and BDV strains) following the standard pro-cedures described by the OIE Manual (OIE, 2008a,b,c). The CubanEMCV 744/03 strain was propagated on BHK-21 cells in mini-mum essential medium (MEM) supplemented with 5% FBS. Afteran approximately 16 h incubation at 37 ◦C, when more than 80%of the cells showed cytopathology, the monolayer was exposed tothree freeze–thaw cycles. PK15, MDBK and BHK-21 non-infectedcell cultures were also used in the specificity assays. The cell cul-tures and media were previously tested to be free of contaminatingpestiviruses (Edwards, 1993; Bolin et al., 1994).
2.3. Clinical standard samples
The availability of standard samples is crucial for a reliable assayvalidation. Therefore, a panel of 40 pig sera from two Interna-tional Interlaboratory Comparison Tests conducted in 2007 and
2008 by the Community Reference Laboratory (CRL) for ClassicalSwine Fever, Hanover, Germany, was used. Briefly, the serum sam-ples with different titres of the virus were obtained as a result of theexperimental infection of pigs with different subtypes and geno-types of pestivirus strains. A BVDV-positive fetal calf serum and
Table 2Sensitivity of the real-time RT-PCR assay to different CSFV genotypes evaluated forboth platforms.
CSFV strain Titre of viral stock(TCID50/mL)
Sensitivity(TCID50/mL)
Alfort 187 (Genotype 1.1) 106.50 101.50
5.00 1.00
L.J. Pérez et al. / Journal of Viro
negative commercial pig serum were also included. Each panelf samples was tested at least three times at the CRL with all theoutine procedures, which included the real-time RT-PCR assayescribed by Hoffmann et al. (2005) and one SYBR Green basedeal-time RT-PCR using the primer pair described by Vilcek et al.1994), for CSFV and non-CSFV pestivirus detection, respectively.
.4. Samples from animals infected experimentally
A total of 38 serum samples from animals infected experimen-ally were evaluated, and the results were compared with those ofhe CRL reference real-time RT-PCR protocol reported by Hoffmannt al. (2005). Briefly, 15 pigs were infected with 105 TCID50 (tissueulture infectious dose 50%) of CSFV (strain Margarita) by intra-uscular injection in the neck region. The serum samples from pigsere collected prior to inoculation (n = 15) at 7 days post-infection
p.i.) (n = 15) and at 13 days p.i. (n = 8).A total of 38 serum samples from animals infected experimen-
ally were evaluated, and the results were compared with thosef the CRL reference real-time RT-PCR reported by Hoffmann et al.2005). Positive results were considered as threshold cycle valuesCt) less than or equal to 42. For samples where fluorescence wasot detectable, the results were considered negative. In short, 15igs were infected intramuscularly in the neck region with 105
CID50 of CSFV (strain Margarita).International standards for animal welfare were used for all ani-
al experiments.
.5. Field samples
A collection of tissue homogenate samples (lymph nodes,pleens or tonsils) from CSFV naturally infected animals (n = 18)nd healthy field animals (n = 12) tested in a previous study (Díaze Arce et al., 2009) were used to assess the performance of thessay in tissues.
.6. RNA isolation and cDNA synthesis
The RNA was extracted from cell culture supernatants, sera andissue homogenates using the RNA viral isolation kit nucleospin IIccording to the manufacturer’s instructions (Macherey-Nagel). Inll cases, an initial volume of 150 �L was used to obtain a final vol-me of 50 �L of RNA, which was stored at −80 ◦C. The synthesis ofDNA was performed by random priming and using M-MLV reverseranscriptase, as described previously by Díaz de Arce et al. (2009).
.7. SYBR Green real-time RT-PCR experimental andhermocycling conditions for LightCycler1.5 and ABIPrism7500latforms
In both cases, the real-time RT-PCR was performed usinghe Quantitect SYBR Green PCR kit (Qiagen GmbH, Hilden,ermany). For CSFV specific detection, the primer pair (CSFV1: CCT-AGGACCAAACACATGTTG/CSFV2: TGGTGGAAGTTGGTTGTGTCTG)
eported previously by Díaz de Arce et al. (1998), which targets aegion corresponding to the NS5B protein, was used. To minimiserimer dimer formation, primer set concentration and thermocy-ling conditions were both optimised (data not shown).
The final experimental and thermocycling conditions of thessay for each platform were the following:
(i) For the LightCycler1.5 (Roche Applied Science, Mannheim,Germany), the reagent/primer final concentration was1xQuantitect-SYBR-Green PCR kit/0.4 �M, with each sam-ple in a 20 �L total reaction volume that included 5 �L of cDNAtemplate. The thermal profile used was the following: 10 min
at 95 ◦C, followed by 40 cycles at 95 ◦C for 10 s, 58 ◦C for 5 sand 72 ◦C for 8 s. After the PCR cycles, a melting curve wasgenerated (0 s at 95 ◦C, 15 s at 65 ◦C, 0 s at 95 ◦C) to discriminatebetween specific amplicons and non-specific amplificationproducts (in all cases the ramp time was 20 ◦C per second).
(ii) For the ABIPrism 7500 (Applied Biosystem), the reagent/primerfinal concentration was 1xQuantitect-SYBR-Green PCRkit/0.4 �M, and each sample was in a 25 �L total reactionvolume that included 5 �L of cDNA template. The thermalprofile used was the following: 10 min at 95 ◦C, followed by 40cycles at 95 ◦C for 30 s, 58 ◦C for 30 s and 72 ◦C for 35 s. Afterthe PCR cycles, a melting curve was generated (15 s at 95 ◦C,1 min at 65 ◦C, 15 s at 95 ◦C) to discriminate between specificamplicons and non-specific amplification products (in all casesthe ramp time was 1 ◦C per second).
2.8. Analytical sensitivity
2.8.1. Analytical sensitivity evaluation by ten-fold dilutions ofCSFV Margarita strain
In this study, the assay detection limit was determined bythe efficacy of the entire method, including the sensitivity of theprimers and the optimisation of the procedure. The real-time RT-PCR assay was evaluated by testing the viral RNAs eluted fromsequential ten-fold dilutions of titrated (105.67 TCID50/mL) CSFVMargarita strain in cell culture medium, negative-CSFV serum andnegative-CSFV 10% tissue homogenate to establish whether thedifferent sample matrices could bias analytical sensitivity. Eachdilution was tested in three replicates.
2.8.1.1. Analytical sensitivity evaluated by infectious doses from dif-ferent CSFV genotypes. Viral RNAs were extracted from sequentialten-fold dilutions of different CSFV strains in cell culture mediumand then evaluated to determine the possible influence of the CSFVgenotype on the test analytical sensitivity (Table 2).
2.8.2. Analytical sensitivity evaluated by target copy numbers2.8.2.1. In vitro transcription. To determine the assay limit of detec-tion in terms of RNA copy numbers, in vitro-transcribed RNAs fromthe target gene were analysed. The target gene of CSFV Alfort strainwas amplified by RT-PCR using the reverse primer coupled to theT7 promoter. The amplification product was transcribed with theRiboMAXTM Large Scale RNA Production System (Promega, Madi-son, WI, USA) in accordance with the manufacturer’s instructions.Finally, the RNA transcript was quantified by spectrophotometry,and the exact number of RNA molecules was calculated follow-ing the formula reported by Fronhoffs et al. (2002). Thus, a total of1.8 × 1013 RNA copies/�L were obtained.
2.8.2.2. Detection limit by RNA copies. The detection limit of the
assay in terms of RNA copy number was determined by testingsequential ten-fold dilutions of the in vitro-transcribed RNA, whichwere obtained as described in Section 2.8.2.1, in nuclease free water(Promega, Madison, USA), negative-CSFV serum and negative-CSFVtissue homogenate at 10%.
The specificity of the assay was assessed by analysis of viral RNAsxtracted from non-CSFV pestivirus (representatives of BVDV-1,VDV-2 and BDV), CSFV strains from the major genotypes (1.1,.2, 2.1, 2.2, 2.3 and 3.4) and other non-pestivirus RNA virusesommonly infecting pigs (Table 1). Each strain used was tested inriplicate.
.10. Intra- and inter-assay variability
The repeatability of the assay was determined by the repeatedesting of a strong positive (105.0 TCID50/mL), medium positive104.0 TCID50/mL) and weak positive (102 TCID50/mL) sample. Thehree samples were obtained by diluting the CSFV Margarita strainn cell culture medium. To evaluate intra-assay variability, eachilution was analysed in triplicate. To measure inter-assay variabil-
ty, each dilution was analysed in three different runs performedy two different operators on different days. The coefficients ofariation for each platform were determined in accordance withreviously published guidelines (U.S. Environmental Protectiongency, 2004).
.11. Interpretation of results
Specimens were considered CSFV positive when (i) the Ct valueas below 40, (ii) the curve in the amplification plot showed an
xponential increase and (iii) a CSFV-specific melting curve (match-ng with that of the positive CSFV control in the same run) wasbtained.
.12. Statistical analysis
The means and standard deviations of the cycle thresholdf SYBR Green real-time RT-PCR for each platform were gener-ted using each instrument’s incorporated software. Significantifferences among mean cycle threshold values of the three con-entration values (high, medium and low TCID50/mL and RNAopies/�L) were analysed by a one-way analysis of varianceANOVA) using the SAS v.7.0 program with a Kruskal–Wallis test.tatistical significance was defined by values of P < 0.05.
. Results
.1. Analytical sensitivity of the SYBR Green real-time RT-PCR
.1.1. Analytical sensitivity evaluation by ten-fold dilutions ofSFV Margarita
The assay’s limit of detection in cell culture medium and serumas 0.1 TCID50/reaction, while in tissue homogenate it was 1
CID50/reaction for both platforms. To determine the linearity ofhe reaction and the PCR efficiency, the threshold cycle values ofndividual dilutions were plotted against the initial gene copy num-er. Thus, the quantitative linear ranges (QLRs) of the assay spanned05–102 TCID50/mL and 105–103 TCID50/mL when evaluated in cellulture medium/serum and tissue homogenate, respectively.
The correlation coefficients (R2) on the AP7500 platformere 0.99 for CSFV-detection in cell culture medium, 0.92 forSFV-detection in serum and 0.86 for CSFV-detection in tissue
omogenate; for the LC1.5 platform they were 0.99 for CSFV-etection in cell culture medium, 0.98 for CSFV-detection in serumnd 0.87 for CSFV-detection in tissue homogenate (data not shown).owever, when the potential influence on analytical sensitivity bySFV genotypes was evaluated, the assay’s limit of detection fluc-
l Methods 174 (2011) 53–59
tuated from 101 to 101.5 TCID50/mL, which was equivalent to 0.03and 0.12 TCID50/reaction, respectively (Table 2).
3.1.2. Analytical sensitivity of the SYBR Green real-time RT-PCR interms of RNA copy number
The limit of detection was 1, 10 and 102 gene copies/�L whennuclease free water, serum and tissue homogenate, respectively,were used as sample matrices for both instruments. The lin-ear ranges of the SYBR-Green real-time RT-PCR assay spanned1010–104 gene copies/�L, 1010–105 gene copies/�L and 1010–105
gene copies/�L for RNA diluted in nuclease free water, serum andtissue homogenate, respectively. Likewise, for the AP7500 plat-form, the correlation coefficients (R2) were 0.99, 0.98 and 0.92 forthe RNA gene copies detected in nuclease free water, serum and tis-sue homogenate, respectively. However, for the LC1.5 platform, thecorrelation coefficients for the RNA gene copies detected in nucle-ase free water, serum and tissue homogenate were 0.99, 0.98 and0.97, respectively (data not shown).
3.2. Analytical specificity
The SYBR Green real-time RT-PCR assay was able to detect allthe CSFV strains and isolates belonging to genotypes and subtypeslisted in Table 1. The melting temperature values of the specificamplified products ranged between 80.0 and 81.6 ◦C (80.7 ± 0.4 ◦C).No specific amplification curves were obtained with any of theother pestivirus (non-CSFV) strains and isolates listed in Table 1.Specific amplification curves were not obtained from heterologousRNA porcine viruses and non-infected cell cultures. Identical resultswere obtained on both platforms.
3.3. Intra-/inter-assay variability
The amplifications were highly reproducible with coefficientsof variation within runs (intra-assay variability) ranging from 0% to2%. The inter-assay variability results ranged from 1.3% to 2.5%.
3.4. Diagnostic performance assessment of the SYBR Greenreal-time RT-PCR
The results of the SYBR Green real-time RT-PCR assay appliedto the standard samples (n = 40) agreed with the expected results(100%) from the CRL (Table 3). Likewise, the serum sample evalu-ation results (n = 38) obtained from pigs infected experimentallywere all 100% in accordance with the outcomes obtained fromthe real-time RT-PCR reported by Hoffmann et al. (2005). Whenthe tissue field sample collection originated from pigs infectednaturally was evaluated using SYBR Green real-time RT-PCR, spe-cific amplification curves were obtained from all positive samples(n = 18). Negative results were obtained from non-infected animals(n = 12). Therefore, taking into consideration all the samples anal-ysed (n = 108), the diagnostic sensitivity and specificity values wereboth 100% in comparison with the reference procedures, becauseno false-negative or false-positive results were observed. No differ-ences in the results were observed when the assay was performedin a different platform.
4. Discussion
The use of real-time PCR in clinical diagnosis is replacing con-
ventional PCR methods and other established diagnostic methods,such as antigen-ELISA and cell culture isolation (Hoffmann et al.,2009). Specifically, real-time RT-PCR has become an essentialmethod used by laboratories for routine diagnosis of CSFV (Depneret al., 2007; Greiser-Wilke et al., 2007).
L.J. Pérez et al. / Journal of Virological Methods 174 (2011) 53–59 57
Table 3Evaluation and comparison of the diagnostic performance of the SYBR-Green real-time RT-PCR using ABI Prism 7500 and LightCycler 1.5 platforms from international standardsamples.
In the present study, a SYBR Green real-time RT-PCR coupled toelting curve analysis for specific CSFV detection was developed.
n addition, the analytical and diagnostic performances of the assayere evaluated on two different commercially available real-timelatforms.
TaqMan-based real-time RT-PCR for the detection of CSFV haseen described previously (Hoffmann et al., 2005; Liu et al., 2007;amnikar Ciglenecki et al., 2008; Cheng et al., 2008). In particu-ar, the real-time RT-PCR described by Hoffmann et al. (2005) haseen reported as one used most frequently (Koenig et al., 2007;epner et al., 2007) tests and has also been proposed as a tool toe used for routine diagnosis of CSFV (Depner et al., 2007). Never-heless, single point mutations within the probe-binding site couldias the outcomes of these types of assays (Belák, 2007). In fact,igh false-negative rates of TaqMan assays for RNA virus detectionPapin et al., 2004; Hughes et al., 2004; Pham et al., 2005; King et al.,006) including CSFV (Leifer et al., 2010) have been reported.
The SYBR Green real-time RT-PCR assay is less influenced byequence variations. In addition, the inclusion of post-amplificationelting curve analysis ensures highly specific detection (Martínez
t al., 2008; Tam et al., 2009; Tan et al., 2009; Kong et al., 2009).oreover, SYBR Green-based real-time RT-PCR is considered a flex-
ble and cost-effective approach (Martínez et al., 2008). Therefore,he SYBR Green real-time RT-PCR assay developed in this study isroposed as an alternative method for CSFV diagnosis.
Within the area of assay design, relevant issues includechemistry, target selection, cycling conditions and thermocyclingplatform selection (Hayden et al., 2008). On the one hand, theselected target comprised a region from the NS5B gene that is highlyconserved among CSFV strains but is also significantly divergentwhen compared with other pestiviruses (Díaz de Arce et al., 2009).On the other hand, even though the chemistries of real-time RT-PCR assays are the same, the chosen platform could vary betweenlaboratories (Bentley et al., 2005). Each instrument has inherentcharacteristics that must be addressed as an essential step in theprocess of assay validation (Bentley et al., 2005). For this reason, theanalytical and diagnostic performances of this assay implementedon two different real-time platforms were compared.
Considering that the temperature transition rate recommendedfor the LightCycler 1.5 instrument is 20 ◦C per second, while therate suggested for the ABI prism 7500 platform is 1 ◦C per second,the time it takes to get from one temperature to the next one inthe heating/cooling cycle is longer in the ABI prism 7500 than inthe LightCycler 1.5 instrument. Consequently, the assay was fasterwhen performed in the LightCycler 1.5 (Fig. 1).
Interestingly, in spite of the Ct value shift observed among theinstruments (Fig. 1), no differences were found between these plat-forms in the analytical and diagnostic performances of the currenttest. However, the analytical sensitivity was higher when the viraldilutions were assessed in cell culture medium or serum than when
Fig. 1. The means of the cycle thresholds of SYBR Green real-time RT-PCR testedfor each platform. High, medium and low values of TCID50/mL and RNA copies/�Lof the CSFV were assessed on AP7500 and LC1.5 platforms. HIGH: 105 TCID50/mL,1010 RNA copies/�L; MEDIUM: 103 TCID50/mL, 106 RNA copies/�L; and LOW: 102
TPco
aesc(
thtans
tbfgtgSd
sslufa
CID50/mL, 104 RNA copies/�L (a, b: statistical significance defined by values of< 0.05 for AP7500 and LC1.5 platforms for TCID50/mL of CSFV; c, d: statistical signifi-ance defined by values of P < 0.05 for AP7500 and LC1.5 platforms for RNA copies/�Lf target gene).
ssessed in homogenate tissue samples. These results could bexplained by the fact that it is harder to extract RNA from tissueamples because of the higher level of organic material, the complexhemical composition, and the potential presence of PCR inhibitorsCone et al., 1992; Petrich et al., 2006).
Similarly, the level of organic material inherent to the serum andissue sample could bias the linearity of the test (data not shown);ence, the importance of considering the sample to be evaluatedo select the same biological matrix for standard dilution. Thus, anccurate quantitation based on infectious doses or viral RNA copyumbers should be accomplished in the assessed sample. In thisense, the assays proposed showed a wide range of quantitation.
The SYBR Green real-time RT-PCR test was proven to be ableo detect the major genotypes of CSFV and was demonstrated toe very sensitive and specific. However, slight differences wereound when analytical sensitivity was assessed from different CSFVenotypes (Table 2). This result may be more related to the initialitre of each CSFV-strain, along with the dilution factor, than to theenomic variability among CSFV genotypes. Therefore, the currentYBR Green real-time RT-PCR seems not to be influenced by theifferent genetic backgrounds of the viruses.
According to Weesendorp et al. (2009,2010), the titre of the CSFVtrains in whole blood may vary depending on the virulence of the
train. The lower titres found in whole blood of pigs infected withow-virulence CSFV strains were in the range of 104–102 TCID50/mLntil day 40 post-infection (Weesendorp et al., 2009,2010). There-ore, taking into account that the viral titre in persistently infectednimals can be higher than 102 TCID50/mL (Weesendorp et al.,
l Methods 174 (2011) 53–59
2009,2010) and that the analytical sensitivity of the test in serumsamples was 0.1 TCID50/reaction, then the current SYBR-Greenreal-time RT-PCR can be used for detecting CSFV in pooled serumsamples. Hence, the proposed assay can be a useful tool in large-scale PCR screening.
With the aim of preventing false negative RT-PCR results dueto inhibitory effects or degradation of the target RNA, an internalpositive control (IPC) should ideally be included in each sample(Hoffmann et al., 2006). Even though a recent report (Weesendorpet al., 2010) suggested that RNA degradation from field samples isnot a common issue for CSFV, the lack of an IPC in the present assayis still considered a limitation regarding the potential presence ofinhibitory substances. Therefore, the development of an IPC is animportant task for a future improvement of the described assay.
5. Conclusion
In conclusion, a validated SYBR Green real-time RT-PCR assayfor the detection and quantitation of CSFV is proposed. A compari-son of two real-time platforms with different ramp times regardingthe analytical and diagnostic performances of the assay was con-ducted. The successful test validation on these two real-time PCRinstruments with the most divergent ramping conditions demon-strated that the assay was not affected by the type of platform andthat it can be used as a reliable tool for large-scale CSFV detection.
Acknowledgements
Work at CReSA was supported by the project BIO2008-04487-C03-03, Consolider Ingenio 2011 “Nconsolider” and a scholarship(2009/10) from the MAEC-AECID program, all from the Spanishgovernment.
References
Belák, S., 2007. Molecular diagnosis of viral diseases, present trends and futureaspects. A view from the OIE Collaborating Centre for the Application of Poly-merase Chain Reaction Methods for Diagnosis of Viral Diseases in VeterinaryMedicine. Vaccine 25, 5444–5452.
Bentley, H.A., Belloni, D.R., Tsongalis, G.J., 2005. Parameters involved in the conver-sion of real-time PCR assays from the ABI prism 7700 to the Cepheid SmartCyclerII. Clin. Biochem. 38, 183–186.
Bolin, S.R., Ridpath, J.F., Black, J., Macy, M., Roblin, R., 1994. Survey of cell lines in theAmerican type culture collection for bovine viral diarrhea virus. J. Virol. Methods48, 211–221.
Cheng, D., Zhao, J.J., Li, N., Sun, Y., Zhou, Y.J., Zhu, Y., Tian, Z.J., Tu, C., Tong, G.Z.,Qiu, H.J., 2008. Simultaneous detection of classical swine fever virus and NorthAmerican genotype porcine reproductive and respiratory syndrome virus usinga duplex real-time RT-PCR. J. Virol. Methods 151, 194–199.
Cone, R.W., Hobson, A.C., Huang, M.L., 1992. Coamplified positive control detectsinhibition of polymerase chain reactions. J. Clin. Microbiol. 30, 3185–3189.
Depner, K., Hoffmann, B., Beer, M., 2007. Evaluation of real-time RT-PCR assay forthe routine intra vitam diagnosis of classical swine fever. Vet. Microbiol. 121,338–343.
Díaz de Arce, H., Núnez, J.I., Ganges, L., Barreras, M., Frias, M.T., Sobrino, F., 1998. AnRT-PCR assay for the specific detection of classical swine fever virus in clinicalsamples. Vet. Res. 29, 431–440.
Díaz de Arce, H., Ganges, L., Barrera, M., Naranjo, D., Sobrino, F., Frías, M.T., Núnez,J.I., 2005. Origin and evolution of viruses causing classical swine fever in Cuba.Virus Res. 112, 123–131.
Díaz de Arce, H., Pérez, L.J., Frías, M.T., Rosell, R., Tarradas, J., Núnez, J.I., Ganges, L.,2009. A multiplex RT-PCR assay for the rapid and differential diagnosis of clas-sical swine fever and other pestivirus infections. Vet. Microbiol. 139, 245–252.
Edwards, S., 1993. Bovine viral diarrhea virus. In: Doyle, A., Griffiths, J.B., Newell,D.G. (Eds.), Cell & Tissue Culture: Laboratory Procedures. John Wiley & Sons,Chichester, UK, pp. 1–8 (Module 7B:5).
Fronhoffs, S., Totzke, G., Stier, S., Wernert, N., Rothe, M., Bruning, T., Koch, B., Sachini-dis, A., Vetter, H., Ko, Y., 2002. A method for the rapid construction of cRNAstandard curves in quantitative real-time reverse transcription polymerase
chain reaction. Mol. Cell. Probes 16, 99–110.
Greiser-Wilke, I., Blome, S., Moennig, V., 2007. Diagnostic methods for detectionof classical swine fever virus—status quo and new developments. Vaccine 25,5524–5530.
via different routes in pigs experimentally infected with classical swinefever virus strains of high, moderate or low virulence. Vet. Microbiol. 133,
L.J. Pérez et al. / Journal of Viro
Romain, C.A., Stevenson, J., Valsamakis, A., Balfour, H.H., 2008. Multicentercomparison of different real-time PCR assays for quantitative detection ofEpstein-Barr virus. J. Clin. Microbiol. 46 (1), 157–163.
offmann, B., Beer, M., Schelp, C., Schirrmeier, H., Depner, K., 2005. Validation ofa real-time RT-PCR assay for sensitive and specific detection of classical swinefever. J. Virol. Methods 130, 36–44.
offmann, B., Depner, K., Schirrmeier, H., Beer, M., 2006. A universal heterologousinternal control system for duplex real-time RT-PCR assays used in a detectionsystem for pestiviruses. J. Virol. Methods 136, 200–209.
offmann, B., Beer, M., Reid, S.M., Mertens, P., Oura, C.A.L., van Rijn, P.A., Slomka,M.J., Banks, J., Brown, I.H., Alexander, D.J., King, D.P., 2009. A review of RT-PCRtechnologies used in veterinary virology and disease control: sensitive and spe-cific diagnosis of five livestock diseases notifiable to the World Organization forAnimal Health. Vet. Microbiol. 139, 1–23.
ughes, G.J., Smith, J.S., Hanlon, C.A., Rupprecht, C.E., 2004. Evaluation of a Taq-Man PCR assay to detect rabies virus RNA: influence of sequence variation andapplication to quantitation of viral loads. J. Clin. Microbiol. 42, 299–306.
amnikar Ciglenecki, U., Grom, J., Toplak, I., Jemersic, L., Barlic-Maganja, D., 2008.Real-time RT-PCR Assay for rapid and specific detection of classical swine fevervirus: comparison of SYBR green and TaqMan MGB detection methods usingnovel MGB probes. J. Virol. Methods 147, 257–264.
oenig, P., Hoffman, B., Depner, K.R., Reimann, I., Teifke, J.P., Beer, M., 2007. Detectionof classical swine fever vaccine virus in blood and tissue samples of pigs vac-cinated either with a conventional C-strain vaccine or a modified live markervaccine. Vet. Microbiol. 120, 343–351.
ong, L.L., Omar, A.R., Hair-Bejo, M., Ideris, A., Tan, S.W., 2009. Development of SYBRgreen I based one-step real-time RT-PCR assay for the detection and differen-tiation of very virulent and classical strains of infectious bursal disease virus. J.Virol. Methods 161, 271–279.
eifer, I., Evertt, H., Hoffman, B., Sosan, O., Crooke, H., Beer, M., Blome, S., 2010. Escapeof classical swine fever C-strain vaccine virus from detection by C-strain specificreal-time RT-PCR caused by a point mutation in the primer-binding site. J. Virol.Methods doi:10.1016/j.jviromet.2010.03.004.
iu, L., Widén, F., Baule, C., Belák, S., 2007. A one-step, gel-based RT-PCR assay withcomparable performance to real-time RT-PCR for detection of classical swinefever virus. J. Virol. Methods 139, 203–207.
iu, L., Xia, H., Belák, S., Widén, F., 2009. Development of a primer-probe energy trans-fer real-time PCR assay for improved detection of classical swine fever virus. J.Virol. Methods 160, 69–73.
artínez, E., Riera, P., Sitjà, M., Fang, Y., Oliveira, S., Maldonado, J., 2008. Simultane-ous detection and genotyping of porcine reproductive and respiratory syndromevirus (PRRSV) by real-time RT-PCR and amplicon melting curve analysis usingSYBR Green. Res. Vet. Sci. 85, 184–193.
oennig, V., 2000. Introduction to classical swine fever: virus, disease and controlpolicy. Vet. Microbiol. 73, 93–102.
oennig, V., Floegel-Niesmann, G., Greiser-Wilke, I., 2003. Clinical signs and epi-demiology of classical swine fever: a review of new knowledge. Vet. J. 165,11–20.
IE, 2008a. Classical swine fever (Hog Cholera). In: Manual of Diagnostic Tests andVaccines for Terrestrial Animals , 6th ed. Office International des Epizooties,Paris, France, pp. 1092–1106 (Chapter 2.8.3).
l Methods 174 (2011) 53–59 59
OIE, 2008b. Bovine viral diarrhea. In: Manual of Diagnostic Tests and Vaccines forTerrestrial Animals , 6th ed. Office International des Epizooties, Paris, France, pp.698–711 (Chapter 2.4.8).
OIE, 2008c. Border disease. In: Manual of Diagnostic Tests and Vaccines for Ter-restrial Animals , 6th ed. Office International des Epizooties, Paris, France, pp.963–973 (Chapter 2.7.1).
Papin, J.F., Vahrson, W., Dittmer, D.P., 2004. SYBR Green-based real-time quantitativePCR assay for detection of West Nile Virus circumvents false-negative results dueto strain variability. J. Clin. Microbiol. 42, 1511–1518.
Petrich, A., Mahony, J., Chong, S., Broukhanski, G., Gharabaghi, F., Johnson, G., Louie,L., Luinstra, K., Willey, B., Akhaven, P., Chui, L., Jamieson, F., Louie, M., Mazzulli, T.,Tellier, R., Smieja, M., Cai, W., Chernesky, M., Richardson, S.E., 2006. Multicentercomparison of nucleic acid extraction methods for detection of severe acuterespiratory syndrome coronavirus RNA in stool specimens. J. Clin. Microbiol. 44,2681–2688.
Pham, H.M., Konnai, S., Usui, T., Chang, K.S., Murata, S., Mase, M., Ohashi, K., Onuma,M., 2005. Rapid detection and differentiation of Newcastle disease virus by real-time PCR with melting-curve analysis. Arch. Virol. 150, 2429–2438.
Rosell, R., Pujols, J., Munoz, I., Núnez, J.I., Ganges, L., Domingo, M., 2008. Antigenicand molecular characterization of border disease virus isolated from pigs. In:7th ESVV Pestivirus Symposium , Uppsala, Sweden.
Tam, S., Clavijo, A., Engelhard, E.K., Thurmond, M.C., 2009. Fluorescence-based mul-tiplex real-time RT-PCR arrays for the detection and serotype determination offoot-and-mouth disease virus. J. Virol. Methods 161, 183–191.
Tan, S.W., Ideris, A., Omar, A.R., Yusoff, K., Hair-Bejo, M., 2009. Detection and dif-ferentiation of velogenic and lentogenic Newcastle disease viruses using SYBRGreen I real-time PCR with nucleocapsid gene-specific primers. J. Virol. Methods160, 149–156.
Terpstra, C., de Smit, A.J., 2000. The 1997/1998 epizootic of swine fever in TheNetherlands: control strategies under a non-vaccination regimen. Vet. Micro-biol. 77 (1–2), 3–15.
Thiel, H.J., Collett, M.S., Gould, E.A., Heinz, F.X., Houghton, M., Meyers, G., Purcell, R.H.,Rice, C.M., 2005. Family flaviviridae. In: Fauquet, C.M., Mayo, M.A., Maniloff, J.,Desselberger, U., Ball, L.A. (Eds.), Virus Taxonomy: VIIIth Report of the Inter-national Committee on Taxonomy of Viruses. Academic Press, San Diego, pp.979–996.
U.S. Environmental Protection Agency. 2004. Quality assurance/quality control guid-ance for laboratories performing PCR analyses on environmental samples. U.S.Environmental Protection Agency, Washington, DC.
Vega, S., Rosell, R., Paton, D.J., Orden, J.A., De la Fuente, R., 2000. Antigenic char-acterization of bovine viral diarrhoea virus isolates from Spain with a panel ofmonoclonal antibodies. J. Vet. Med. 47, 701–706.
Vega, S., Rosell, R., Orden, J.A., De la Fuente, R., García, A., Pérez, T., Davis, L., Wakeley,P., Sandvik, T., 2002. Antigenic and molecular characterization of border diseasevirus isolates from Spain. In: 5th Pestivirus Symposium , Cambridge, UK.
Vilcek, S., Herring, A.J., Herring, J.A., Nettleton, P.F., Lowings, J.P., Paton, D.J., 1994.Pestiviruses isolated from pigs, cattle and sheep can be allocated into at leastthree genogroups using polymerase chain reaction and restriction endonucleaseanalysis. Arch. Virol. 136, 309–323.
Weesendorp, E., Stegeman, A., Loeffen, W., 2009. Dynamics of virus excretion
9–22.Weesendorp, E., Willems, E.M., Loeffen, W.L.A., 2010. The effect of tissue degradation
on detection of infectious virus and viral RNA to diagnose classical swine fevervirus. Vet. Microbiol. 141, 275–281.
