RT-LAMP assay: African Horse Sickness virus detection in Horses (Equis) | Vetlexicon
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RT-LAMP assay: African Horse Sickness virus detection

ISSN 2398-2977

Synonym(s): Reverse transcription loop-mediated isothermal amplification assay for the rapid detection of African Horse Sickness virus, Field test for the detection of African Horse Sickness virus


  • Laboratory confirmation of clinical suspicion of African horse sickness (AHS) African horse sickness has been traditionally made by detection of AHS virus (ASHV) in blood or post-mortem tissues using classical virus isolation (VI), immunohistochemistry or antigen detection ELISA techniques. 
  • However these methods have now been superseded by the adoption of more rapid, sensitive and accurate molecular diagnostic techniques such as reverse-transcription-PCR (RT-PCR) and real-time RT-PCR (rRT-PCR).
  • These assays are widely used in most AHSV diagnostic laboratories and are the recommended tests by the OIE (World Organisation for Animal Health) (OIE, 2012) not only for confirmation of clinical diagnosis but also for animal certification purposes, import/export testing and surveillance.
  • Whilst current RT-PCR and rRT-PCR methods are accurate, rapid and sensitive, they are required to be performed in laboratory settings and samples must be transported under the appropriate conditions from the point of collection to the laboratory. This process can delay the confirmation of suspicion of AHSV infection and the implementation of adequate control measures.
  • The development of rapid field-based diagnostic assays that can be used either in the laboratory or at the point of sample collection therefore enable the fast implementation of animal movement controls, since unlike rRT-PCR a result is obtained within 30 min.
  • Loop-mediated isothermal amplification (LAMP) is an isothermal, autocyling, strand-displacement DNA amplification technique.
  • LAMP utilizes 4-6 primers which target 6-8 regions of the pathogen’s genome.
  • LAMP can combine a reverse transcription step (RT-LAMP) to detect RNA viruses such as AHSV.
  • LAMP can be performed at a single temperature using either a fluorimeter, eg Genie® II device or PCR thermocycler, or be performed using a heat block/water bath combined with simple, disposable visualization using molecular lateral-flow devices (LFD’s).
  • LAMP is a highly sensitive technique that allows for the detection of a very low number of nucleic acid copies with comparable analytical sensitivity to rRT-PCR.
  • LAMP assays can be multiplexed to detect multiple pathogens in a single reaction.



  • Within the literature there are an increasing number of LAMP assays for the detection of veterinary pathogens.
  • The most common application of LAMP is as a simple screening assay to rapidly confirm disease (<30 min) by detecting very low amounts of circulating pathogen nucleic acid (DNA or RNA). This protocol describes exactly this method.
  • However, in combination with the development of portable LAMP platforms or visualization of LAMP products by LFD, enables disease confirmation at the point of suspicion, eg on the stable yard.
  • Quick confirmation of disease enables control measures to be implemented immediately.

In combination

Other points

  • LAMP can be performed in laboratory settings using automated nucleic acid extraction platforms, or can be performed in the field using manual extraction kits.
  • LAMP assays are highly sensitive with comparable analytical sensitivity to rRT-PCR.
  • LAMP assays are less prone to sample derived inhibition and can be performed on a wide range of clinical samples. Some LAMP assays have been validated for use without combining an extraction step.


Source of test material

  • AHSV can be detected in EDTA blood collected during the early febrile stage, or in post-mortem tissues, eg spleen, lung and lymph nodes.

Quantity of test material

  • The quantity of test material will depend on the extraction kit used, however most nucleic acid extraction kits only require 50-100 µl of original sample.
  • For tissue, approximately 1 g/tissue of material is required.

Sample collection technique

  • Blood should be collected into an EDTA (purple/pink) BD Vacutainer and inverted 8-10 times following manufacturers guidelines.
  • Tissue, eg spleen, lung and lymph nodes, can be collected at post-mortem.

Quality control


  • Whilst AHSV is not considered a zoonotic risk, normal levels of care should be applied to the handing of blood and tissues.

Sample storage

  • EDTA blood and tissues should be stored at 4˚C/39.2˚F if testing on the same day, or frozen at -80˚C/-112˚F if testing is to be carried out at a later time point.

Sample transport

  • Transportation of samples should comply with the OIE (Chapter 7.2-7.4 of the Terrestrial Animal Health Code) and International Air Transport Association (IATA) requirements for air, sea and land.
  • See Office International des Epizooties (OIE).



  • Extract RNA in duplicate from the sample (EDTA blood or tissue) using an appropriate extraction kit.
  • Heat the extracted RNA at 95˚C/203˚F for 5 min.
  • RT-LAMP is performed in a total reaction mixture of 25 µl containing: 15 µl isothermal master mix ISO-001, optimized primer concentrations African Horse Sickness: RT-LAMP amplification - oligonucleotide primers, 2 U (0.2 µl) AMV reverse transcriptase, 5 µl RNA template and made up to volume with nuclease-free water.
  • RT-LAMP reactions are run at 65˚C/149˚F for 30 min on a portable LAMP machine, eg Genie® II device.
  • Samples should be tested in duplicate.
  • Post-amplification anneal analysis should be performed on LAMP products by heating the LAMP reaction to 98˚C/208.4˚F for 1 min, then cooling to 80˚C/176˚F decreasing at 0.05˚C/39.09˚F per sec (using fluorescence detection) using the portable LAMP machine, eg Genie® II.


  • The Genie® II can be purchased from OptiGene Ltd, UK.
  • ISO-001 can be purchased from OptiGene Ltd, UK in either a wet format for the user to add primers and AMV or in a lyophilized format already containing AMV and disease specific primers.



  • The AHSV RT-LAMP analytical sensitivity is comparable to rRT-PCR over a 4 log10 dilution range.
  • The AHSV RT-LAMP diagnostic sensitivity is 97.4%.


  • The specificity of the AHSV RT-LAMP assay is 100% when assessed against related viruses such as Bluetongue virus (BTV) and Equine encephalosis virus (EEV).

Technique intrinsic limitations

  • At present the test has only been validated for use on extracted RNA, therefore an extraction kit (either manual or automated) must be used in combination.
  • At present the test must be performed and analysed on a fluorimeter platform, eg field based such as the Genie® II or any laboratory based thermocycler.
  • Good quality pipettes are required due to the aliquotting of small volumes.

Technician extrinsic limitations

  • Test should only be performed by technicians whom are familiar with working with exotic pathogens for the knowhow surrounding appropriate biosecurity Biosecurity and biosafety Laboratory: safety.

Result Data

Normal (reference) values

  • ISO-001 contains an intercalating dye, enabling results to be visualized using fluorescence collected at 1 min intervals. A positive RT-LAMP reaction is indicated by an exponential increase in fluorescence (δR), with the Genie® II automatically reporting the time to positivity (Tp). The lower the Tp value the more viral RNA copies present in the sample.
  • Samples should be considered positive (AHSV-specific) if amplification occurs, eg a Tp value is obtained, and the LAMP products anneal in the AHSV amplicon-specific temperature range of 87.0-88.6˚C/188.6-191.48˚F.

Abnormal values

  • Any LAMP amplicons which have an anneal temperature outside the range of 87.0-88.6˚C/188.6-191.48˚F should be retested.

Errors and artifacts

  • Errors are minimized by the performance of an anneal analysis at the end of the LAMP reaction.

Further Reading


Refereed Papers

  • Recent references from PubMed and VetMedResource.
  • Fowler V L et al (2016) Development of a reverse transcription loop-mediated isothermal amplification assay for the detection of vesicular stomatitis New Jersey virus: use of rapid molecular assays to differentiate between vesicular disease viruses. J Virol Methods 234, 123-131 PubMed.
  • Fowler V L (2016) Development of a novel RT-LAMP assay for the rapid detection of African Horse Sickness Virus. Transbound Emerg Dis PubMed.
  • Howson E L A et al (2015) Evaluation of Two Lyophilized Molecular Assays to Rapidly Detect Foot-and-Mouth Disease Virus Directly from Clinical Samples in Field Settings. Transbound Emerg Dis PubMed.
  • Waters R A et al (2014) Preliminary validation of direct detection of foot-and-mouth disease virus within clinical samples using reverse transcription loop-mediated isothermal amplification coupled with a simple lateral flow device for detection. PLoS One 9 (8), e105630 PubMed.
  • Guthrie A J (2013) Diagnostic accuracy of a duplex real-time reverse transcription quantitative PCR assay for detection of African horse sickness virus. J Virol Methods 189 (1), 30-35 PubMed.
  • Quan M et al (2010) Development and optimisation of a duplex real-time reverse transcription quantitative PCR assay targeting the VP7 and NS2 genes of African horse sickness virus. J Virol Methods 167 (1), 45-52 PubMed.
  • Fernandez-Pinero J et al (2009) Rapid and sensitive detection of African horse sickness virus by real-time PCR. Res Vet Sci 86 (2), 353-358 PubMed.
  • Aguero M et al (2008) Real-time fluorogenic reverse transcription polymerase chain reaction assay for detection of African horse sickness virus. J Vet Diagn Invest 20 (3), 325-328 PubMed.
  • Maree S & Paweska J T (2005) Preparation of recombinant African horse sickness virus VP7 antigen via a simple method and validation of a VP7-based indirect ELISA for the detection of group-specific IgG antibodies in horse sera. J Virol Methods 125 (1), 55-65 PubMed.
  • Notomi T (2000) Loop-mediated isothermal amplification of DNA. Nucleic Acids Res 28 (12), E63 PubMed.
  • Sailleau C (1997) Detection of African horse sickness virus in the blood of experimentally infected horses: comparison of virus isolation and a PCR assay. Res Vet Sci 62 (3), 229-232 PubMed.
  • House J A (1996) A blocking ELISA for detection of antibody to a subgroup-reactive epitope of African horsesickness viral protein 7 (VP7) using a novel gamma-irradiated antigen. Ann N Y Acad Sci 791, 333-344 PubMed.
  • Zientara S et al (1995) Application of the polymerase chain reaction to the detection of African horse sickness viruses. J Virol Methods 53 (1), 47-54 PubMed.
  • Bremer C W, du Plessis D H & van Dijk A A (1994) Baculovirus expression of non-structural protein NS2 and core protein VP7 of African horsesickness virus serotype 3 and their use as antigens in an indirect ELISA. J Virol Methods 48 (2-3), 245-256 PubMed
  • Zientara S et al (1993) Diagnosis and molecular epidemiology of the African horse sickness virus by the polymerase chain reaction and restriction patterns. Vet Res 24 (5), 385-395 PubMed.
  • Chuma T et al (1992) Expression of the major core antigen VP7 of African horsesickness virus by a recombinant baculovirus and its use as a group-specific diagnostic reagent. J Gen Virol 73 (4), 925-931 PubMed.
  • Laviada M D et al (1992) Detection of African horsesickness virus in infected spleens by a sandwich ELISA using two monoclonal antibodies specific for VP7. J Virol Methods 38 (2), 229-242 PubMed.

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