Development of a new quantitative RT-PCR to detect lymphocytic choriomeningitis virus

Lymphocytic choriomeningitis virus (LCMV) is a neglected rodent-borne virus, with a worldwide distribution. The common mouse Mus musculus acts as reservoir and vector in the biological cycle of the virus. Surveillance of LCMV infection in mice is of importance as they are a widely used animal model in research and, through contact with them or their fluids, humans can be infected. Although most human cases are asymptomatic, LCMV infection can cause mild to severe, even fatal, and new diagnostic tools need to be developed to improve its detection. In the present work we report the development of a new method for the detection of LCMV RNA by quantitative reverse transcription polymerase chain reaction (RT-qPCR), able to detect all LCMV strains described to date. RT-qPCR targeting the S segment was developed and evaluated. Specificity and sensitivity were determined, and its limit of detection (LOD) was defined. The method designed is able to detect all 5 LCMV lineages described to date, with a LOD of 5.6 genome copies/μL. Its design with a built-in internal amplification control allows the detection of false negative results. Other arenaviruses were found not to cross-react with the method designed. In conclusion, a new diagnostic RT-qPCR for the detection of LCMV have successfully designed and validated. Improved detection techniques allow to reduce the turnaround time in the diagnosis of infections and to improve epidemiological surveillance in humans and animals.

1 Introduction

Lymphocytic choriomeningitis virus (LCMV) is a zoonotic agent, which belongs to the Arenaviridae family. This family includes 5 genera, but only the genus Mammarenavirus has been described as competent to infect mammals, including humans (1). Rodents are recognized as the main natural reservoirs of the genus, although some mamarenaviruses have been isolated from fruit-eating bats (2, 3). To date, two major groups have been described within Mammarenavirus genus based on the geographical distribution of the viruses and their rodent reservoirs, the Old World Group (OWG) and the New World Group (NWG) (4). LCMV has the widest known distribution of all Mammarenaviruses, being predominant in North America and Europe. This wide distribution is explained by the geographical distribution of its main host and reservoir, the mouse Mus musculus, which is present in all continents except Antarctica and the North Pole (5, 6). Other reservoirs of the virus, and therefore potential transmitters, are the field mouse (Apodemus sylvaticus), the fawn mouse (Apodemus flavicolis), hamsters and guinea pigs (Cavia porcellus) (7–9).

LCMV is an enveloped, segmented single-stranded, ambisense RNA genome, whose segments are called L (large) and S (small) (10, 11). Considering the genetic diversity of LCMV S-segment, five lineages (Lineage I, II, III, IV and V) have been described (7, 8, 12, 13). Lineage I is the most widespread and includes most of the described strains, such as the classical laboratory strains Armstrong and WE.

In their natural host, rodents, LCMV transmission usually occurs by inhalation of aerosols generated from contaminated droppings (saliva, semen, milk, urine and faeces) and typical social behaviors of rodents, such as grooming, favor transmission. In addition, the virus can be transmitted vertically to subsequent generations by intrauterine infection, resulting in a generation of persistently infected mice that will excrete the virus throughout their lives (11, 14, 15). Because they usually exhibit no overt clinical signs, the infection can spread silently through high-density colonies typical of laboratory animal facilities. As a result, personnel who handle or work near these animals—including animal care technicians, veterinarians, and laboratory staff—are at increased risk of occupational exposure to this zoonotic pathogen (16, 17). Humans can be infected by inhalation of infected excreta that may be aerosolised during sweeping or cleaning mice (18) or by direct contact with contaminated material. Animal bites, ingestion of contaminated food and solid organ transplants are another potential route of LCMV infection (11, 19–22). Most infections are asymptomatic, or only a mild, self-limiting febrile illness develops. However, in severe cases, after a temporary improvement of these flu-like symptoms, a second phase of central nervous system involvement usually follows. The symptoms of this second phase are usually those of classical aseptic meningitis or meningoencephalitis (23, 24). Other LCMV-related complications include hydrocephalus, pancreatitis, orchitis, arthritis, parotitis, and pericarditis (11, 25, 26). LCMV can also be transmitted vertically from mother to foetus via the transplacental route (27, 28). In congenital LCMV infection the mortality rate ranges from 14 to 35% depending on the time of infection (23, 29). When infants survive, congenital malformations such as chorioretinitis, hydrocephaly, psychomotor retardation, microcephaly or ventriculomegaly are common (23, 30–32).

At present, no specific vaccine or treatment against LCMV is available either in humans or in rodents. Therefore, the cornerstone of disease control strategies should be based on regular health surveillance and good diagnosis of infection. Laboratory diagnosis of LCMV infections can be approached in two ways: using direct or indirect methods. Acute LCMV infections can be diagnosed by viral isolation or by conventional PCR assays from samples such as serum, blood, urine, cerebrospinal fluid, organs and throat swabs (33, 34). Serological methods, ELISA and immunofluorescence assay (IFA) are used to detect IgM and IgG antibodies in serum samples, although very few commercial tests are currently available (14, 35).

Based on these concerns, the present study describes the development and validation of a novel RT-qPCR assay for the detection of LCMV, designed to enhance diagnostic capacity and animal surveillance and occupational health protection settings.

2 Materials and methods

2.1 Primer and probes design

LCMV specific primers and probe were designed in the nucleoprotein (NP) region of the S segment. For this purpose, an extensive alignment of 34 LCMV sequences, representative of all LCMV lineages described to date, was performed. All sequences were available in the NCBI GenBank database (National Center for Biotechnology Information) (Table 1). Primers and probe were designed using Primer Express® v2.0 software (Applied Biosystems, Foster City, CA, USA). Primers were also analysed with the Oligo Primer Analysis v6.0 program (Molecular Biology Insights).

Table 1

Table 1. Sequences used in the design of RT-qPCR oligonucleotides for the detection of lymphocytic choriomeningitis virus.

Once designed, primers and probe were introduced into a second alignment, containing 24 different sequences of several Mammarenaviruses (Table 2), in order to perform an in silico specificity study, and to check that the sequences designed for the primers and probe in the chosen region did not have sufficient homology to be able to amplify other viruses of the same genus (36). Two different TaqMan® probes were designed, one specific LCMV probe (LCMV_probe) and an internal control specific probe (IC_Probe), labelled at the 5′-end with FAM and NED, respectively. MGB (Minor Groove Binder) and NFQ (Non-Fluorescent Quencher) were used at the 3′-ends (Table 3).

Table 2

Table 2. Sequences of viruses belonging to the Mammarenavirus genus used to study in silico the specificity of the developed LCMV RT-qPCR.

Table 3

Table 3. Primers and probes designed for LCMV and IC detection.

2.2 Design of internal control plasmid and DNA standard

To detect possible inhibitions in the PCR reaction, an internal control (IC) was designed. A method like that described by other authors was used (37–39), consisting of the synthesis of a plasmid DNA fragment containing a known sequence of the BK virus, flanked by the sequence of the primers designed in this work for LCMV amplification in the RT-qPCR (Table 4). To generate a plasmid DNA-based standard curve, primers designed for RT-qPCR were used to amplify part of the S fragment of the LCMV Amstrong strain (GenBank: AY847350). PCR products were purified and cloned using TOPO TA Cloning® kit (Invitrogen, MA, USA) according to the manufacturer’s instructions. Plasmids obtained were sequenced to confirm the absence of mutations, quantified by measuring the optical density at 260 nm using a NanoDrop spectrophotometer (Thermo Fisher Scientific, USA), and the copy number was calculated using the formula of Reed-MuenchQuantified DNA was linearized using the Not I restriction enzyme (New England Biolabs, MA, USA) and, for LCMV standard DNA, a 10-fold dilution curve was constructed from 10⁶ to 10¹ DNA copies/μL using water as diluent.

Table 4

Table 4. Primers used for internal control construction.

2.3 RNA extraction

The nucleic acids used in this work were extracted using the commercial kit QIAamp® Viral RNA (Qiagen, Hilden, Germany) following the manufacturer’s recommendations. For RT-qPCR optimisation, a RNA extract (Armstrong strain 53b; GenBank: AY847350) from a LCMV isolated in Vero E6 cells was used as template.

2.4 Real time RT-PCR assay

The RT-qPCR reaction was carried out in a 7,500 Fast Real-Time PCR system (Applied Biosystems), using the commercial kit QuantiTect® Multiplex RT-PCR Kit (Qiagen, Hilden, Germany). For each assay, 5 μL of RNA sample was mixed with 20 μL of a reaction mix containing: 3.45 μL of sterile water, 12.5 μL of Quantitec multiplex RT-PCR master, 0.25 μL of each primer (100 μM), 0.75 μL of LCMV probe (10 μM), 1.5 μL of IC probe (10 μM), 1 μL of IC (10² copies/μL) and 0.3 μL of RT mix. Amplification parameters were established as follows: an initial retrotranscription step at 50 °C for 30 min, a DNA polymerase activation cycle at 95 °C for 15 min, followed by 45 amplification cycles at 94 °C for 45 s and 60 °C for 1 min. Fluorescence measurement for FAM and NED was performed simultaneously at the end of each cycle. Cycle threshold values (Ct) were measured as the point at which the fluorescence signal of the sample fluorescence crossed a predetermined threshold value.

2.5 Evaluation of sensitivity and specificity of the qRT-PCR

To assess the sensitivity of RT-qPCR, standard plasmids were used as template. Each reaction was replicated four times in the same assay, and the tests were performed on three different days. The LOD was calculated by adaptation of the Reed–Muench formula (40). Specificity was assessed by testing a panel of 28 arenaviruses genomes and other encephalitis-producing viruses belonging to other viral families (Supplementary Table S1).

2.6 Evaluation of repeatability (intra-assay precision) and reproducibility (inter-assay precision)

The repeatability and reproducibility of the RT-qPCR assay were determined using six different concentrations (10⁶, 10⁵,10⁴, 10³, 10² and 10 copies/μL) of the standard plasmids. Quintuplicate analysis of each dilution were performed on the same day to determine intra-assay variability. Additionally, three different experiments were performed in different days to assess inter-assay variability. The coefficient of variation of the Ct values was determined based on the intra-assay and inter-assay results.

2.7 Evaluation of the possible application of the real-time PCR method using preparation of synthetic clinical samples

Due to the absence of LCMV PCR-positive human clinical samples in our laboratory, a panel of 15 simulated human clinical samples was constructed to evaluate this RT-qPCR. Six known negative samples were also included in the panel. The simulated clinical samples, urine, blood or serum, were generated by inoculating 20 μL of a titled LCMV viral culture (Armstrong strain 53b GenBank: AY847350) into 180 μL of the clinical sample to finally obtain samples containing different concentrations of viral genome (10⁴–1 copies of viral genome per microlitre). These clinical samples had previously been tested for LCMV RNA to ensure negativity using a conventional nested RT-PCR adapted from Ledesma et al. (8) by using LCMV sequences instead of degenerations (Supplementary Table S2). Viral inactivation and RNA extraction were performed using standard procedures as described above.

3 Results

3.1 Design of primers, probes and standard curve for the RT-qPCR assay

Primers and probes targeting a highly conserved fragment of the LCMV S-segment amplifying a 96 bp sequence were designed.

The standard curves for RT-qPCR were constructed using a 10-fold dilution series ranging from 10⁶ to 10 copies/μL of the corresponding DNA standard in each reaction. The standard curve for RT-qPCR showed that the slope is −3.448, the amplification efficiency (Eff%) is 95.006 and that the correlation coefficient R² is 0.999 (Figure 1). Both R² and efficiency values indicate a strong linear relationship between the template and the Ct value, validating the reliability of the newly developed RT-qPCR assay.

Figure 1

Figure 1. Amplification for the LCMV DNA standard. The copy number of the standard plasmid ranged 10⁶ (F), 10⁵ (E), 10⁴ (D), 10³ (C), 10² (B), 10¹ (A) and 10¹ (G) copies/μL. (A) Amplification curve for LCMV target representing fluorescence increase vs. cycle. (B) Standard curve representing Ct vs. DNA quantity.

3.2 Sensitivity and specificity of real-time RT-PCR

The LOD was assessed using serial dilutions of the quantified LCMV DNA standard the sensitivity limit of this RT-qPCR using the Reed–Muench formula is 5.6 copies/μL. To assess specificity, different viral genomes were used as templates of the LCMV RT-qPCR. No amplification curves were detected for any other viruses, demonstrating that the developed RT-qPCR method does not cross-react neither with other encephalitis-producing viruses nor with other arenaviruses.

3.3 Repeatability and reproducibility of the real-time PCR assay

The repeatability and reproducibility of the RT-qPCR assay were evaluated by testing different concentrations of the LCMV standard plasmids. The relationship between the standard deviation (SD) and the average (X̅) obtained for the Ct values in five replicates of each DNA standard concentration, was calculated using the coefficient of variation (CV). Intra-assay CV values ranged from 0.39 to 1.1% and between assays from 0.90 to 4.59% for the limit dilution (Table 5). These results indicated a satisfactory repeatability and reproducibility for the developed RT-qPCR assay.

Table 5

Table 5. Repeatability and reproducibility RT-qPCR assay for detection of LCMV.

3.4 Detection in synthetic clinical samples

The results of the simulated clinical samples showed that this RT-qPCR (Figure 2) was able to detect LCMV RNA with a concentration of 10 copies of viral genome/μL, in 15 of the 16 samples tested, with Ct values between 34 and 44, depending on the sample type. Samples contaminated with 1 copy of viral genome/μL were negative in all assays (Table 6).

Figure 2

Figure 2. Detection of LCMV RNA in synthetic clinical samples by RT-qPCR. Cycle threshold (Ct) values obtained for serum, blood, and urine samples spiked with LCMV at different concentrations are shown. Each sample type was tested across three independent experimental assays (Day 1–3), with duplicate reactions (D1, D2) performed on each day. Symbols correspond to the viral genome concentration used for spiking: 10¹ copies/μL (■); 10² copies/μL (●); 10³ copies/μL (▼) and 10⁴ copies/μL (♦). The assay consistently detected viral RNA down to 10 genome copies/μL in all sample types, with Ct values ranging from ~30 to 44, whereas samples spiked with 1 copy/μL were not detected (see Table 6).

Table 6

Table 6. LCMV detection in synthetic clinical samples.

4 Discussion

Lymphocytic choriomeningitis virus (LCMV) is a pathogen whose life cycle is associated with the distribution and activity of rodent populations. In wild mouse populations, studies show prevalences ranging from 0 to 25%, depending on the region and sampling conditions (41–44). In other mammals, LCMV infections are not uncommon (13, 45) and can even infect humans (24, 46, 47). Human infections are underestimated and little studied, but in Europe there are studies showing human seroprevalences of 6.8% in Croatia (48), 7% in Italy (49) and 1.7–2% in Spain (50, 51). All these data indicate that LCMV is currently circulating in our environment. In recent decades, climate change has caused global temperatures to rise, making rodents able to reproduce faster and more efficiently (52, 53). It has also led to changes in vegetation or precipitation that have caused rodent populations to seek refuge indoors. This has changed the behavior of rodent populations, making them more likely to come into contact with humans and increasing the transmission of zoonotic diseases (54, 55).

On the other hand, a significant relationship has been found to exist between rodent population density and the prevalence of LCMV infection (48). And that within infested households, LCMV infection rates in mice range from 50% to almost 100% (34, 48), increasing the risk of LCMV infection in humans living there. This increased transmission of LCMV when there is a high density of rodents may be of particular importance in animal breeding and experimental animal facilities, where, in addition to the health of the animals, the health of the workers may be affected. In 1972–1973 at the University of Rochester Medical Centre, an outbreak of LCMV was reported among 48 infected workers, caused by Syrian hamsters used in tumour research (56). In 2012 there was an LCMV outbreak in several EEUU rodent facilities, and a third of the employees were infected, developing, some of them, aseptic meningitis (57). Animal health surveillance in these facilities is essential to ensure both the health of animals and workers, as the absence of microorganisms that could interfere with research. Federation of European Laboratory Animal Science Associations, US Department of Health and Human Services recommend the implementation of surveillance programmes to detect and control potential pathogens, including LCMV (58–60). Although traditional pathogen detection methods such as necropsies remain powerful tools for the diagnosis of infectious diseases in animals, molecular methods significantly improve the sensitivity of the tests. For that purpose, we describe a new RT-qPCR assay for LCMV detection that would be suitable for rapid and specific diagnostic objectives. Currently, several conventional nested RT-PCRs have been published for the molecular diagnosis of LCMV infection, which involve multiple time-consuming steps and are associated with an increased risk of contamination. Few quantitative PCRs have been described to date. Of the published designs, some require a previous cDNA synthesis step (61, 62), and others are SYBR Green quantitative PCRs (34). Extensive efforts during the SARS-CoV-2 pandemic have focused on the validation and standardization of RT-PCR assays. These studies highlighted the need for assay optimization, sensitivity and specificity assessment, inter-laboratory comparability, and harmonized validation protocols. Lessons learned from SARS-CoV-2 diagnostics provide valuable benchmarks to strengthen molecular diagnostic strategies for other viral pathogens (63, 64). The RT-qPCR developed in this study is a one-step PCR theoretically capable of detecting all five LCMV lineages described to date. Its design is based on hydrolysis probes (Taqman probes), which increases specificity and reproducibility in the detection of PCR amplification products, compared to other qPCR chemistries. In addition, an internal control was also designed to be co-amplified in each reaction mixture to aid detection of false negative results due to lack of amplification for example by polymerase inhibition. Other features of this new RT-qPCR are a low detection limit, a high ratio of genomic load to Ct and consistently repeatability and reproducibility. Using synthetic clinical samples, the efficacy of this new PCR could be evaluated, since it was able to detect the presence of LCMV in all three types of synthetical clinical samples tested: serum, blood and urine. Using the same synthetic clinical samples, the accuracy and stability of the new PCR was also validated by comparison with a nested PCR commonly used in the laboratory for LCMV detection. The results showed that the detection limit of the new RT-qPCR in clinical samples was between 10 and 1 copies/μL, which was in line with the detection limit obtained when using standard DNA (5.6 copies/μL). Moreover, this technique was validated using RNA obtained from mouse samples. The evaluation of the specificity of RT-qPCR was performed using genomes of viruses belonging to Arenaviridae or different families, all of them producing the same or similar symptoms. The absence of amplification signal in these samples demonstrated the high accuracy of the method developed for LCMV detection, as it only detects the genome of the virus for which it has been designed. The diagnosis of infectious diseases in clinical samples requires the use of specific, highly sensitive and reproducible techniques. The RT-qPCR designed in this work provides these characteristics, guarantees the detection of all LCMV lineages circulating to date without cross-reaction with other arenaviruses or other encephalitis-producing viruses and, in addition, avoids false-negative results.

Data availability statement

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found in the article/Supplementary material.

Ethics statement

Ethical review and approval was not required for the study on human participants in accordance with the local legislations and institutional requirements. The participants provided their written informed consent to participate in this study.

Author contributions

LH: Validation, Methodology, Data curation, Formal analysis, Writing – review & editing, Conceptualization, Writing – original draft. NL: Conceptualization, Supervision, Methodology, Writing – review & editing, Data curation, Investigation, Writing – original draft, Visualization, Formal analysis. FM: Methodology, Writing – review & editing. AP: Methodology, Writing – review & editing. MS-S: Writing – review & editing, Formal analysis, Supervision. AV: Data curation, Conceptualization, Writing – review & editing, Formal analysis, Supervision, Funding acquisition.

Funding

The author(s) declare that financial support was received for the research and/or publication of this article. This work was supported by Instituto de Salud Carlos III (“PI19CIII/00014 and PI24CIII/00038) and the project PLEC2021–007968 (NEXTHREAT) funded by MCIN/AEI/10.13039/501100011033 and European Union NextGenerationEU/PRTR.

Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

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Supplementary material

The Supplementary material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fvets.2025.1651039/full#supplementary-material

Abbreviations

LCMV, lymphocytic choriomeningitis virus; LOD, limit of detection; qPCR, quantitative polymerase chain reaction; RT-qPCR, quantitative reverse transcription polymerase chain reaction; Ct, cycle threshold; CV, coefficient of variation; IC, internal control; Bp, base pairs.

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