Development of a qPCR Workflow for Screening Culicoides for Viral Pathogens

Research paper on experimentation performed under the umbrella of S. Zapata's research at the Center of Vector Biology at Rutgers University.

Abstract
Biting midges, or Culicoides, are a type of arboviral vector that pose substantial economic and environmental threats. This experimentation is part of a larger effort to survey Culicoides midges across New Jersey, identifying what species are present and what viral or bacterial elements they may carry. Samples of midges were collected and divided into diverse-species pools, which were RNA extracted to facilitate county-level surveillance of two Culicoides-transmitted vectors: epizootic hemorrhagic disease virus (EHD) and bluetongue virus (BTV). Two quantitative PCR assays were adopted and integrated to form a qPCR duplex, which was used to test midge samples after a substantial validation process. Preliminary results indicate the presence of EHD, which will be confirmed with a secondary qPCR followed up by sequencing analysis.

​Introduction
Culicoides biting midges are a hematophagous species with a global range of habitation. They are known for their role in transmitting numerous pathogens, many of which pose significant risks to livestock and wildlife as arbovirus vectors (Mellor et al., 2000). In New Jersey, the primary arboviruses of concern in relation to Culicoides are epizootic hemorrhagic disease virus (EHD) and bluetongue virus (BTV). Currently, there are only two confirmed Culicoides species vectors in North America: C. sonorensis (EHD and BTV) and C. insignis (BTV) (McGregor et al., 2022). Recent studies have additionally identified C. stellifer and C. venustus as possible vectors of EHD (McGregor et al., 2019). The distinction between confirmed and possible vectors follows the four criteria established by the WHO Study Group on Arthropod-Borne Viruses in 1960, which require multiple lines of evidence to classify a species as an arbovirus vector (Köhler, 1968, p. 22). Confirmed and potential Culicoides vectors for EHD have been recognized, however, none of these species have been documented in New Jersey, leading to questions about what other Culicoides species might be acting as vectors in this region.
Evidence suggests that Culicoides species capable of transmitting EHD and BTV share many similarities. This is supported by the fact that both viruses infect several overlapping ruminant hosts, such as white-tailed deer, sheep, and cattle (Jiménez-Cabello et al., 2023). The viruses are similar in structure and evolutionary origin, as both are non-enveloped, double-stranded RNA viruses belonging to the order Orbivirus (Jiménez-Cabello et al., 2023). The diseases caused by these viruses, EHD and bluetongue disease (BT), produce severe symptoms–including cyanosis, internal hemorrhaging, fever, and dehydration–although the severity of symptoms varies between ruminant species (Becker et al., 2020). Outbreaks of EHD and BT have been documented globally, causing direct economic losses by reducing livestock populations and indirect losses due to trade restrictions and other factors (Jiménez-Cabello et al., 2023). In New Jersey, a significant outbreak of EHD in 2021 received considerable public attention, following outbreaks in 1988, 1996, 2007, and 2012 (Stallknecht et al., 2015; NJDEP).

Materials and Methods
Sample Collection
Mixed species samples were collected using a CDC miniature UV/LED light trap. Midges in the samples were then isolated from other arthropods and separated into multiple diverse-species pools based on their morphological characteristics following the dichotomous keys of Jamnback (1965) and Wirth et al. (1985). Each pool contained 50-100 individuals and was labeled by county of origin and sample location.

DNA/RNA Extraction and Purification
Diverse-species samples were extracted using a Qiagen AllPrep DNA/RNA Mini kit. Initially homogenized using a VWR Mini Bead Mill, the samples were centrifuged, and the supernatant was transferred to AllPrep DNA spin columns. The spin columns were stored at 4°C for subsequent DNA purification to be used in a different line of experimentation, while the flow-through was utilized for RNA purification. RNeasy spin columns were used for RNA purification, and samples were stored at -80°C until qPCR analysis was performed.

qPCR Duplex Validation
The EHD assay described by Portanti et al. (2023) and the BTV assay described by Becker et al. (2020) were selected for validation. Initially, the assays were validated separately using only a corresponding positive DNA control. Once the reaction conditions were shown to be effective the assays were then also tested using both positive DNA controls at once (ie EHD assay tested with the EHD and BTV controls) to ensure no non-specific amplification occurred. This was then repeated with midge homogenate for the same purpose, but using the entire genome. After the assays were confirmed to not produce unwanted PCR products, they were combined into a preliminary duplex. The concentrations of the assays relative to each other were optimized to produce the version of the duplex used in sample analysis.

Sample Testing and Analysis
The ideal primer and probe concentrations for the duplex determined by the validation process were used to set up qPCR reactions on the purified RNA samples isolated from collected midge pools. Positive pools will be run in a confirmatory qPCR reaction and samples that test positive twice will be further confirmed for their presence of EHD or BTV using sequencing analysis

Results

​Figure 1. Amplification Plot of EHD/BTV qPCR Duplex
​After preliminary analysis of the BTV and EHD assays individually to confirm their efficacy, they were combined to validate the duplex. This is one such step in the validation process, where a 10-fold serial dilution of the EHD and BTV positive controls was created over 6 steps to visualize different reaction conditions in the presence of midge homogenate. The EHD Ct values were nearly unchanged by these conditions at higher concentrations of the serial dilution, while the BTV primers and probe proved to be more sensitive.

Figure 2. Standard Curve Plot of EHD/BTV Duplex
​This standard curve graph was generated by the analysis software based on the experimentation of Figure 1. The graph is based on the assumption that the samples of different reaction conditions were actually ‘replicates’ of the same concentration of positive control at the same conditions, meaning differences in absorption across a single reaction condition set alter both the efficiency percentage and the R^2 value calculated by the software. As it stands, both efficiency values are over 60% and the R^2 values are over 0.99.

Figure 3. Amplification Plot of 2023 Midge Pools
​The EHD/BTV duplex was used to test RNA samples from 2023 midge collections. This amplification shows amplification of the EHD and BTV positive controls, as well as EHD amplification in 4 samples. These samples need to be retested to confirm they are positive for EHD, followed by sequencing to confirm.

Discussion
​Development of the qPCR duplex involved a series of validation steps to ensure the primers and probes were present at concentrations that enable the detection of both EHD and BTV. Optimization was performed using a 10-fold serial dilution in 6 steps of both positive controls to confirm the amplification efficiency was linear for each target. Potential cross-reactivity was assessed by combining the two assays, and none was observed. The duplex assay was then tested using midge homogenate to confirm its function in the target species without amplification of off-target sequences. Testing a variety of duplex reaction conditions yielded an overall EHD efficiency of 60.7% and a BTV efficiency of 63.4%, with R² values of 0.94 and 0.99, respectively. The most efficient reaction conditions were chosen for sample testing.
The duplex was then used to screen for EHD and BTV in diverse-species pools collected in 2023. These samples had already been homogenized, and undergone RNA extraction and purification, meaning they were ready to undergo qPCR testing. This resulted in, as shown in Figure 3, 4 samples being identified as positive for EHD. These samples must now be retested using the qPCR assay to ensure they are true positives, in which case they will be sequenced to ultimately confirm the presence of EHD. The same testing procedures will be performed on other 2023 samples, as well as 2024 samples currently being processed, with multiple levels of confirmation for both EHD and BTV positives.

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