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Article

Development of Quantitative Real-Time PCR Tests for the Identification of Biting Midge Species and Clades (Diptera: Ceratopogonidae) of the Obsoletus Group (Subgenus Avaritia), Including Important Viral Vectors in Europe

by
Oliver Dähn
1,*,
Bernd Hoffmann
1,
Doreen Werner
2,
Bruno Mathieu
3 and
Helge Kampen
1
1
Friedrich-Loeffler-Institut (FLI), Federal Research Institute for Animal Health, Südufer 10, 17493 Greifswald, Germany
2
Leibniz Centre for Agricultural Landscape Research (ZALF), Eberswalder Str. 84, 15374 Müncheberg, Germany
3
Institutes of Bacteriology and Parasitology, Medical Faculty, University of Strasbourg, UR 3073 PHAVI, 67000 Strasbourg, France
*
Author to whom correspondence should be addressed.
Insects 2025, 16(5), 500; https://doi.org/10.3390/insects16050500
Submission received: 22 March 2025 / Revised: 2 May 2025 / Accepted: 5 May 2025 / Published: 7 May 2025
(This article belongs to the Special Issue Approaches to Diseases Control Centered on Arthropod Vectors of Animal and Human Pathogens)
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Simple Summary

Biting midges of the genus Culicoides are small hematophagous flies, responsible for the spread of economically important arboviruses such as bluetongue virus. In Europe, the species of the Obsoletus Group have been identified as the main vectors, but the knowledge of the geographical distribution and ecology of many taxa of this group is scarce, not least of all because of missing identification tools. To address this issue, a high-throughput PCR approach was developed that allows the simultaneous examination of large quantities of individuals for the presence of isomorphic Obsoletus Group taxa, even in a pool of unsorted insects. In this work, the difficulties in the development of the PCR are addressed. The unresolved phylogeny of certain taxa and the difficulty in obtaining suitable DNA material for PCR validation have posed significant obstacles. Despite these challenges, the new PCR approach proved its potential by accurately identifying their target taxa. This establishes a crucial foundation for the routine implementation of surveillance studies, enabling detailed insights into the spatiotemporal distribution of vector species. As a result, current and future risk scenarios can be assessed, and countermeasures can be initiated to prevent viral spread. Especially in times of increasing globalization and a rapidly changing climate, this helps protect livestock and mitigate economic losses.

Abstract

Species of the widespread Obsoletus Complex (Culicoides subgenus Avarita Fox, 1955) have been implicated as potential key vectors during the bluetongue and Schmallenberg epidemics in Central Europe in 2006 and 2012. Although extensive efforts have been made to clarify vector–pathogen relationships, one of the most important steps in this process—correct species identification—remains difficult, due to the presence of isomorphic species within the Obsoletus Group. To overcome the difficulties in morphological species identification, several PCR tests were developed. With the aim of developing a high-throughput PCR, capable of differentiating all putative vector species and newly described haplotypes of the subgenus Avaritia present in Europe, a dataset of 4407 published sequences of the mitochondrial (mt) cytochrome c oxidase subunit I (COI) was used to develop specific primers and probes, which can either be applied in a singleplex PCR or in different multiplex PCR approaches. The real-time PCR achieved very high diagnostic sensitivity (100%) and specificity (91.7%) and reliably detected the three clades of C. obsoletus sensu stricto (s.s.) in a pool of specimens. Thus, the new real-time PCR approach will provide an excellent tool for large-scale monitoring, which could improve the understanding of the biology, geographical distribution, and habitat preference of European biting midge species involved in the transmission of bluetongue, Schmallenberg, and epizootic hemorrhagic disease viruses.

1. Introduction

Hematophagous insects are important vectors of disease agents worldwide, particularly species belonging to the dipteran families Culicidae, Ceratopogonidae, Psychodidae, and Simuliidae [1,2]. Contrary to mosquitoes (Diptera: Culicidae), which are certainly the most prominent representatives of this group of arthropods, biting midges of the genus Culicoides Latreille, 1809 (Diptera: Ceratopogonidae) have received much less public attention, even though they have been identified as vectors of filarial nematodes, protozoan parasites, and more than 50 viruses of veterinary and medical importance [3,4].
In Europe, indigenous biting midge species had been neglected as potent vectors until the unexpected emergence and fulminant spread of bluetongue virus (BTV) in several countries outside of the known distribution area of its main vector C. imicola. BTV can cause severe infection in ruminants, especially sheep [5,6]. The bluetongue disease (BTD) has a long history in Africa [7] and was initially reported in South Africa in the late eighteenth century [8]. Meanwhile, BTD has become endemic in the Mediterranean basin [9,10], and occasional spreading events take place in Central Europe, as recently observed during the incursion of BTV serotype 3 (BTV-3) and BTV-12 [11,12,13,14,15]. With the emergence of these virus strains, around one-third of all yet-known serotypes (13 of 36) [16] have already been confirmed for Europe [17] but the accessibility of effective vaccines [18] has so far prevented the disease from becoming endemic. The closely related epizootic hemorrhagic disease virus (EHDV) of the genus Orbivirus, on the other hand, was first described in North America where it was isolated from a wild-tailed deer [19]. It had never been reported in Europe until the recent isolation of EHDV-8 from infected farm animals in Italy (Sicily, Sardinia) and southern Spain (Andalusia) at the end of 2022 [20]. A high genetic similarity between the virus strain from Italy to the EHDV strains circulating in Tunisia in 2021 and 2022 suggests its introduction via the North African route [20], as also suspected for BTV in 2006 [21]. In the meantime, EHDV-8 has also been detected in Portugal and southern France [22]. In contrast to BTV and EHDV, nothing is known about the origin of the ruminant-pathogenic Orthobunyavirus Schmallenberg virus (SBV). Since its first appearance near the German–Dutch border in 2011, it has never completely disappeared [23] and rapidly established an enzootic status in many European countries [24,25].
Several native biting midge species have been implicated as potential vectors of these viruses, although true evidence of vector competence is scarce due to difficulties in rearing them under laboratory conditions. However, initial infection studies investigating the susceptibility of field-caught Culicoides to BTV [26,27] and EHDV [28,29] and virus isolations from field-collected specimens identified members of the Obsoletus Group (subgenus Avaritia Fox, 1955) as the main vectors of BTV [30,31,32,33,34,35], SBV [36,37,38,39,40,41,42,43,44,45,46,47], and EHDV [48].
In Europe, the Obsoletus Group was believed to consist of five species, including the eponymous C. obsoletus Meigen, C. scoticus Downes & Kettle, C. chiopterus (Meigen, 1830), C. dewulfi Goetghebuer, 1936, and C. montanus Shakirzjanova [49,50]. However, there are still taxonomic obscurities that have not yet been resolved completely, highlighting the ongoing challenge of identifying endemic vector species. Due to the analysis of various genetic markers, C. dewulfi has been excluded from the Obsoletus Group [51,52,53,54,55], while the phylogenetic status of C. chiopterus is still ambiguous. Depending on the gene markers analyzed, the latter is assigned to the Obsoletus Group in some cases (COI, ITS-1) [56,57] and excluded in others (28S rDNA) [56]. The remaining three taxa, C. obsoletus, C. scoticus, and C. montanus, are closely related species whose isomorphic females cannot be distinguished based on their morphological features. To facilitate morphological classification, these species are collectively referred to as ‘Obsoletus Complex’ or ‘Obsoletus/Scoticus Complex’ [58]. Culicoides montanus was thought to only be present further south in Europe [49]. A recent study reviewed all available COI sequences of subgenus Avaritia species published in GenBank, and the re-classification of some sequences revealed a much more northern distribution (UK, Germany, Poland) of the species [59]. Its taxonomic status within the Obsoletus Complex is still controversially discussed. Some authors propose that C. montanus might be a genetic variant of C. obsoletus rather than a separate species due to very low differences in the DNA sequence at certain gene loci [58,60]. Additionally, over the past two decades, further genetic variants of C. obsoletus [51,61,62,63] and C. scoticus [54,58] have been identified, with increasing evidence for C. obsoletus clades O1, O2, and O3 representing independent species [59].
All these native vector species feed on livestock and find excellent conditions for the development of their larvae in the surroundings of livestock farms [64], which significantly increases the risk of spreading BTV, SBV, and EHDV. Culicoides obsoletus and C. scoticus are generalists with opportunistic feeding behavior whose larvae can be found in a wide range of habitats such as forest litter, tree holes, silage residue, or manure [65]. Culicoides chiopterus and C. dewulfi almost exclusively take their blood meal from cattle and are strongly adapted to cattle dung as a breeding substrate [64,66,67]. Given this, along with the high density of animal farms and the extensive livestock trade within the European Union, it is not surprising that culicoid-borne pathogens spread and establish quickly, with considerable consequences for livestock industries. Although the dimensions of direct and indirect economic losses are difficult to assess [64], the financial impact e.g., of BTV-8 in Germany alone is estimated to be between 157 and 203 million Euro from the first cases in 2006 to the re-emergence in 2018 [68]. These immense costs emphasize the importance of effective vector management, but the required knowledge of the biology, ecology, and spatiotemporal distribution of native biting midge species is still limited.
One of the main reasons for these knowledge gaps is the costly and time-intensive nature of conducting surveillance studies, which rely on the consistent cooperation of selected livestock farms, skilled scientific personnel, and entomologists with years of experience to accurately identify captured biting midges. The identification of adult Culicoides is usually performed based on their morphology [54,69] but is rarely achievable for the isomorphic specimens of the Obsoletus Group [58,70].
Accordingly, for the reliable identification of these taxa, the development of different molecular methods has been the subject of numerous studies (e.g., [71,72,73]). By far the most widely used molecular techniques are those based on DNA amplification through polymerase chain reaction (PCR) [69]. Meanwhile, direct sequence analysis (DNA barcoding) of several mitochondrial (e.g., COI, 18S rRNA, cytb), ribosomal (ITS1, ITS2), and nuclear (e.g., CAD) markers has become a standard procedure for species identification and phylogenetic studies within the genus Culicoides [69]. However, a major disadvantage of DNA barcoding is the need to compare generated sequences with public genome databases such as GenBank, which have been shown to contain a substantial number of incorrect entries [59,74]. Additionally, the method allows only a single specimen to be processed per reaction tube [74], posing a major limitation for surveillance studies that often collect thousands of individuals per night [69]. This may be particularly problematic if species of the Obsoletus Group constitute a significant proportion of the captured specimens [73,75]. Hence, several medium-throughput multiplex PCR assays have been developed for the identification of the different Obsoletus Group members (e.g., [57,70,76,77]). However, most of these PCR tests are relatively complex to perform and analyze, and they are generally limited to specialized research groups, as they require the pre-sorting of captured insects and depend on time-consuming gel electrophoresis. To address at least some of these challenges, conventional multiplex PCRs for identifying almost all west Palearctic taxa of the subgenera Culicoides and Avaritia were recently developed [59,74]. The PCR tests are easy to analyze (producing one specific band per taxon) and can be run simultaneously using a standardized PCR protocol. While these PCR tests enable the analysis of multiple individuals in a pooled sample, they still require the morphological pre-sorting of catches at least to the group or complex level, which is why these tests are rather unsuitable for high-throughput screening.
A solution to this bottleneck may be provided using real-time PCR tests that are capable of identifying pooled Culicoides specimens, but only three of these assays have been developed so far for the differentiation of specimens belonging to the Obsoletus Group/Complex [62,73,78]. Of these assays, one was developed for the determination of the relative proportion of species in a pool of insects [73], which would make the analysis of large Culicoides collections to the species level viable [69]. However, this PCR is not up to date as it does not consider the different clades of C. obsoletus, whose geographical distribution and role in the spread of BTV, SBV, and EHDV is unknown. To address this issue, a real-time PCR approach was developed in this study that considers the latest taxonomic findings and is intended to quantitatively detect all potential vector species in unsorted pool samples.

2. Materials and Methods

2.1. Insect Collection and Morphologic Pre-Identification

Biting midges and non-culicoid Diptera (bycatch) were trapped year-round, both inside and outside livestock holdings, using BG-sentinel UV-light suction traps (Biogents, Regensburg, Germany), primarily in the framework of a Germany-wide monitoring project conducted from 2019 to 2022. Individuals of a few subgenus Avarita taxa have not been found in the collections and were obtained from other samplings in different European countries, Russia, and the USA. All captured midges were morphologically pre-identified at least to the group or complex level using common identification keys [79,80,81,82] and kept in 75% EtOH for further molecular analysis.

2.2. Extraction of Genomic DNA

DNA extraction was performed using either whole insects or parts of them (thorax, abdomen) (Table S1). For DNA extraction from whole specimens, the insects were removed from the storage vessel and placed on a clean paper tissue. After evaporation of the remaining preservative for 1 min at room temperature, the specimens were individually transferred to a clean 2 mL reaction tube containing either 180 µL buffer ATL and 20 µL proteinase K (QIAamp DNA Mini Kit) or 350 µL of house-intern ZB5d medium supplemented with antibiotics as described in Ries et al. [83]. Three steel beads with a diameter of 3 mm (TIS GmbH, Gauting, Germany) were added, and samples were completely homogenized for 3 min at 30 Hz using a TissueLyser II device (Qiagen, Hilden, Germany). Total genomic DNA was isolated with the QIAamp DNA Mini Kit (Qiagen) from ATL/Proteinase K containing homogenate in a 50 µL elution volume or with the NucleoMag VET Kit (Macherey-Nagel, Düren, Germany) from samples agitated in ZB5d/antibiotics mixture and finally eluted in 100 µL VEL-buffer. For DNA extraction from insect abdomens, total abdomens were removed from the insects and directly transferred to 200 µL buffer ATL/proteinase K mixture for further processing, as mentioned above. In some cases, DNA available from previous extractions (thorax and partial abdomen) [53,55] was directly used.

2.3. DNA Amplification and Sequencing

DNA extracts of single specimens or their body parts were used for the amplification of taxon-specific COI gene fragments, as described [59]. In total, 125 extracts were used for PCR amplification. Generated PCR products were subjected to agarose gel electrophoresis, and ethidium bromide-stained DNA bands of expected length (517 bp or 685 bp) were excised from the gels. The amplicons were recovered by the QIAquick Gel Extraction Kit (Qiagen) and further used for cycle sequencing with the PCR primers, applying the protocol of the BigDye Terminator v1.1 Cycle Sequencing Kit (Thermo Fisher Scientific, Dreieich, Germany). The DNA fragments were purified (Bioanalysis NucleoSEQ Kit, Macherey-Nagel), and 15 µL of the eluate was mixed with an equal volume of Hi-Di formamide (Thermo Fisher Scientific). The samples were sequenced either in one or both directions on a 3500 Genetic Analyzer device (Applied Biosystems/Hitachi, Darmstadt, Germany). Obtained sequences were edited with Geneious Prime software version 2021.0.1 (Biomatters, Auckland, New Zealand) and checked against NCBI GenBank (www.ncbi.nlm.nih.gov, last accessed on 11 December 2022). Edited sequences were deposited in GenBank and corresponding DNA samples were used for PCR validation.

2.4. Data Analysis and PCR Design

The real-time PCR was developed based on a multiplex in-house assay targeting the four taxa C. obsoletus, C. scoticus, C. chiopterus, and C. dewulfi. According to the reviewed consensus sequences from a recently published study [59], the PCR assay was updated and expanded to include the different clades of C. obsoletus. To do so, the specific consensus sequences of all subgenus Avaritia taxa were compared in a Geneious multiple alignment (Biomatters). Taking the resulting interspecific sequence differences into account, taxon-specific primers and probes were revised or newly designed according to common guidelines for PCR design (e.g., [84,85]). Since all genetic variants of the individual taxa had to be considered, degenerated bases were inserted at corresponding positions. Appropriate primer and probe sequences were checked in terms of melting temperature and hairpin structure or primer–dimer formation using the Oligo Analysis Tool (https://eurofinsgenomics.eu/en/ecom/tools/oligo-analysis/, last accessed on 11 December 2022) and analyzed with the NCBI BLAST Tool (https://blast.ncbi.nlm.nih.gov/Blast.cgi, 2.13.0 release, last accessed on 15 November 2022) for repetitive sequences. Selected primers (RP-Cartridge-Gold purified) and Taqman probes (HPLC purified) were ordered from Eurogentec (Kaneka Eurogentec S.A., Seraing, Belgium). The used CFX96 Touch Real-Time PCR Detection System (Bio-Rad, Feldkirchen, Germany) enables the simultaneous discrimination of a maximum of five targets in different fluorescence channels (FAM: 510–530 nm, HEX: 560–580 nm, TEX: 610–650 nm, Cy5: 675–690 nm, Quasar705: 705–730 nm), making it impossible to distinguish all six taxa in a single reaction well. Additionally, the PCR needed to be cost-effective and provide greater flexibility for the end-user. To address these requirements, the real-time PCR was designed in a modular manner, allowing the performance of each PCR in a singleplex format or their combination in various multiplex configurations, depending on the objectives of the scientific question.

2.5. Modular Real-Time PCR

Each master mix (singleplex or multiplex) was composed of 10 µL of 2× QuantiTect Multiplex PCR NoROX reagent (Qiagen), 0.375 µM of each primer, 0.125 µM of each probe, and 2 µL of DNA template, resulting in a total volume of 20 µL. The real-time PCR was performed in a 96-well format on CFX96 Touch Real-Time PCR Detection System (Bio-Rad) using the following thermoprofile: 15 min at 95 °C (activation of Taq polymerase), followed by 42 cycles of 30 s at 95 °C (denaturation), 30 s at 55 °C (primer annealing), and 30 s at 72 °C (primer elongation). The excited fluorescence of the probes was measured after each cycle at the end of the primer elongation step. The developed PCRs were tested in terms of their functionality, specificity, and capability for multiplexing and finally validated with genetically pre-identified (sequenced) material from field-collected culicoids or artificial COI genes [59,74].

3. Results

In this study, quantitative real-time PCRs targeting the mitochondrial (mt) COI gene were developed for the taxon-specific identification of six members belonging to the Culicoides subgenus Avaritia.
Taking the interspecific polymorphisms and intraspecific conserved regions of taxon-specific consensus sequences into account, 97 primers and 29 TaqMan probes were designed (Table S2). All primers and probes were pre-tested in terms of their functionality, PCR performance, and specificity to find the best working combination for the accurate detection of the different Avaritia taxa. Due to weak fluorescence signals, all Cy5 probes have been replaced by Atto647N labeled probes, which can be measured in the same detection channel (675–690 nm). The best-working primers and probes are summarized in Table 1. For four taxa, C. obsoletus clade O1, C. obsoletus clade O2, C. obsoletus clade O3, and C. scoticus clade 1, plasmids containing cloned COI target sequences [59] were used to generate standard curves and calculate the number of gene copies per single biting midge (Table S1). Of 142 individual DNA samples measured, only 57 were extracted following the same protocol (whole-body extraction with QIAamp DNA Mini Kit and an elution volume of 50 µL). Hence, these samples were used for the determination of the mt genome copy number per biting midge. The analysis revealed Ct values between 17.48 and 22.57 (mean Ct: 19.32), and a mean copy number of 1.2 × 106 was calculated. Based on these findings, serial dilutions of cloned targets were prepared for each taxon (around 100- to 1000-fold lower and higher than the mean copy number per specimen) to evaluate the PCR performance of the different taxon-specific singleplex PCRs and compare the results to those obtained with the one-tube multiplex PCR approach (Figure 1). In the cases of C. chiopterus and C. dewulfi, no plasmids with cloned COI sequences were available at the time of assay design. Hence, serial dilutions of taxon-specific COI amplicons were prepared, which achieved similar Ct values to the plasmid standards of the other taxa.
Both the singleplex PCR and the multiplex PCR approach showed an almost similar performance. The calculated standard curves of all singleplex PCRs were linear over the selected measuring range with linear regression values (R2) of 1.000 and an amplification efficiency (E) between 83.9% and 92.4%. The R2s of the different taxa in the multiplex PCR (0.995–1.000) were comparable but the amplification efficiencies were, in most cases, lower (78.8–91.5%) than in the singleplex PCRs, which might be caused by undesired interactions of primers and probes or steric hindrance. The best results were achieved with the taxon-specific PCR of C. scoticus clade 1, which shows no difference in the overlay of calculated standard curves for both singleplex and multiplex PCR (Figure 1D). In general, no significant reduction in PCR sensitivity (∆Ct < 1) was determined between all singleplex PCRs and the multiplex approach (sixplex PCR), indicating no detrimental interactions of the taxon-specific primers in the PCR mixture. However, differences in the signal intensities (∆RFU) between +11.9% (C. obsoletus clade O2 PCR) and −22.8% (C. dewulfi PCR) were noticed for the multiplex PCR, and in some cases unspecific amplification signals were observed (Figure 1D,F), suggesting interactions (cross-reactivities) of respective TaqMan probes with other molecules of the PCR mixture or even the unspecific detection of non-target taxa.
To confirm the observed non-specific amplification signals and determine the problem-causing molecules, each singleplex PCR was tested with 1 × 108 mt genome copies (equivalent to the DNA amount of 100 specimens). Ct values between 10.46 and 13.85 were achieved for the specific DNAs and negative results were observed for the non-target taxa in the case of the PCR for C. obsoletus clade O1, C. obsoletus clade O2, and C. chiopterus (Figure 2A,B,E). However, unspecific DNA amplification (Ct values ≥ 34.65) was detected for the taxon-specific primers and probes of (i) C. obsoletus clade O3 with the DNAs of C. obsoletus clade O1, C. obsoletus clade O2, and C. scoticus clade 1 (Figure 2C); (ii) C. scoticus clade 1 with the DNAs of all three clades of C. obsoletus (Figure 2D); and (iii) C. dewulfi with the DNA of C. chiopterus (Figure 2F). Based on the Ct values obtained when testing a single biting midge (Ct < 20) and according to the observed cross-reactivities, a cut-off value was implemented, which considers samples with Ct > 30 as negative.
Taking this cut-off value into account, the diagnostic sensitivity (dSe) and specificity (dSp) of the multiplex PCR concept were determined for the various biting midge taxa through the analysis of genetically pre-identified specimens from field collections (Table S1). In total, 91 DNA samples of individual biting midges (whole specimens or body parts) were analyzed in a two-step PCR approach: First, the sixplex PCR was performed to identify specimens of C. obsoletus (no clade-separation), C. scoticus clade 1, C. chiopterus, and C. dewulfi. In a second step, the samples positive for C. obsoletus were further determined to the respective clades of C. obsoletus (clade O1, clade O2, or clade O3). Table 2 shows a summary of the obtained PCR results. Except for one DNA sample, which was positive for both C. obsoletus clade O2 (Ct 25.60) and C. obsoletus clade O1 (Ct 28.56), all target taxa of subgenus Avaritia biting midges were correctly identified, giving 100% dSe and 98.9% dSp.
The capability of the developed real-time PCR to analyze pools of unsorted biting midges was further assessed by examining the DNA of 36 Culicoides species and haplotypes representing seven additional subgenera endemic in the Palaearctic region (Table 3). The sixplex PCR was negative (Ct > 30) for 33 of the 36 taxa, giving a total dSp of 91.7%. However, two taxa of the subgenus Avarita (C. montanus and C. scoticus clade 2) were incorrectly detected as C. obsoletus (Ct 19.54) and C. scoticus clade 1 (Ct 19.97), respectively. In the case of unspecific detection of C. montanus, the sample was re-tested with the triplex PCR (C. obsoletus clade identification) for further verification of the cross-reactivity. The results were negative with the C. obsoletus clade O2 and C. obsoletus clade O3 PCRs, whereas a ‘false-positive’ result was observed with the C. obsoletus clade O1 PCR (Ct 22.91). A subsequent comparison of the taxon-specific consensus sequences revealed almost 100% identity in the primer and probe binding regions for C. obsoletus clade O1 and C. montanus (Table S3). Hence, samples identified as C. obsoletus clade O1 should be considered C. obsoletus clade O1/C. montanus and might be further analyzed by DNA barcoding. The same applies to C. scoticus clades 1 and 2. The analysis of the DNAs of both taxa displayed identical primer binding site sequences and only two base differences between the C. scoticus clade 1 probe (Csco-P-FAM) and the COI sequence of C. scoticus clade 2. Additionally, the synthetic DNA of C. newsteadi haplotype N2 (subgenus Culicoides Latreille) was incorrectly identified as C. scoticus clade 1 (Ct 27.12) if the sample was tested with the sixplex PCR or fourplex approach but was not detected with C. scoticus PCR alone. Further analysis of the false-positive PCR product was performed using gel electrophoresis followed by sequence comparison of all molecules applied in the fourplex PCR with the consensus sequence of C. newsteadi haplotype N2. It turned out that an 89 bp long PCR fragment was amplified with the forward primer of C. obsoletus clade O1 (CobsO1_F) and the reverse primer of C. scoticus (Csco_R) with the DNA of C. newsteadi haplotype N2, which was detected with the specific Taq-Man probe of C. scoticus clade 1 (Csco_P-FAM) (Table S3). Hence, if there is a high chance of finding specimens of C. newsteadi haplotype N2 in field collections, a morphological pre-identification of the samples to the group level (separation of specimens according to Obsoletus Group and Pulicaris Complex via wing pigmentation) or the usage of the C. scoticus clade 1 PCR in a multiplex approach without the C. obsoletus clade O1 PCR is recommended.
To confirm the applicability of the real-time PCR for the direct measurement of unsorted insect samples (Culicoides plus bycatch), genetically pre-identified DNA samples of 15 non-culicoid Diptera (Table 4) were analyzed with the developed multiplex PCR (sixplex). For three taxa, Ct values > 35.8 were obtained with the PCRs for C. obsoletus and C. dewulfi, despite a considerable number of mismatches between the primer and probe sequences and the non-culicoid DNA (Table S4). However, all tested samples were negative if the mentioned cut-off value was applied. Hence, if the observed cross-reactivities are considered, the developed PCR approach has great potential to be used for the specific detection of Avaritia biting midge taxa in unsorted UV-light catches.

4. Discussion

In the past two decades, PCR-based methodologies have increasingly been implemented in the discipline of entomology as standard diagnostic tools, especially when dealing with sibling species of, e.g., culicoid vectors [73,87]. Several PCR tests have been developed to distinguish isomorphic specimens of the widespread Obsoletus Group, the main vectors of BTV, SBV, and EHDV in central Europe. Most of these assays include time-consuming gel electrophoresis steps [57,69,76,77,88,89] and are inefficient at analyzing large quantities of samples that usually accumulate during entomological surveillance and monitoring activities. Given these deficiencies, the knowledge about the geographic distribution and ecology of vector species is still fragmentary.
In this study, a high-throughput multiplex real-time PCR was successfully developed that enables the quantitative detection of vector species within the Culicoides subgenus Avaritia, including the clades of C. obsoletus. A major challenge was the acquisition of sufficient DNA material of the various Culicoides taxa. Therefore taxon-specific PCR development and its verification were restricted, and synthetic DNA had to be used in some cases. Additionally, the PCR was meant to consider the sequences of all genetic variants (haplotypes) of the target taxa across a wide geographical area to ensure universal applicability. To obtain the necessary genetic data, publicly accessible databases such as GenBank are available. However, sequences deposited in databases should be used with caution due to a considerable number of incorrectly annotated entries. Recently, Dähn et al. [59] successfully developed a conventional PCR for the different taxa of the subgenus Avaritia based on reviewed and scrutinized COI sequences from GenBank. This sequence dataset was successfully used in this study to design taxon-specific primers and probes.
The developed real-time PCR achieved a diagnostic sensitivity (dSE) of 100% due to the high-copy properties of the COI gene. Notwithstanding, the exact quantification of the target DNA turned out to be difficult due to inhomogeneous DNA extraction. The biting midge size proved to be particularly disadvantageous as during extraction of an individual midge, some parts were not completely lysed. This likely explains the obtained variations in the Ct-values when analyzing single midges (19.32 ± 4 SD).
High sensitivity also indicates highly efficient PCR amplification, which can even amplify the smallest traces of DNA (a few nanograms) [90]. Millions to billions of copies are rapidly produced [91], particularly when generating short amplicons, as is common in real-time PCRs (50–150 bp) [92]. This can readily lead to carryover contamination from previous PCRs [93]. When assessing the specificity of the developed PCR, this contamination was particularly problematic. Despite preventive measures, such as spatial separation of the master mix preparation and DNA sample addition, amplification signals were observed for several non-target taxa. For example, the DNA of C. newsteadi haplotype N1 (Ct 36.09) was detected with the taxon-specific PCR of C. dewulfi, even though the probe sequence (26 bp in length) exhibited 11 mismatches compared to the sequence of this DNA sample (c.f. Table S3). Since numerous PCR developments and sequencing efforts of various taxa of the subgenus Culicoides [74] and Avaritia [59] were conducted simultaneously in the same laboratory facilities, contamination appears to be the most likely explanation. This assumption could be confirmed by repetitive measurement of the sample, which only exceptionally resulted in positive detection of the non-target taxon. A cut-off value was successfully established to differentiate between non-specific and specific amplification signals in the developed PCR assay.
Other false-positive results that occurred are associated with issues arising from unclear phylogeny of certain Culicoides taxa. For instance, the taxon-specific PCR of C. obsoletus clade O1 developed in this study also detected C. montanus. Several authors suggested that C. montanus represents a genetic variant of C. obsoletus clade O1 [58,59,60], so its status as a distinct species remains controversial. Due to the extremely high degree of sequence homology between C. obsoletus and C. montanus, it has not yet been possible to distinguish both species when using the COI gene as a molecular marker for PCR development [59,62,77,88,94]. Interestingly, despite the even higher sequence similarity in ribosomal ITS1 (98.3%) and ITS2 (98.6%) spacers, as compared to the mt COI gene (97.2%), some authors have been able to discriminate C. montanus from C. obsoletus by those genetic markers [57,78]. Additionally, C. obsoletus and C. montanus specimens can easily be separated by experienced entomologists based on the morphology of both the palp and the long trichodea [82], without any overlapping between both species. The same genetic and morphological discrepancy is observed for C. scoticus clade 1 and C. scoticus clade 2 [59,82]. Assuming that C. montanus and C. scoticus clade 2 are not distinct species, the calculated diagnostic specificity of the multiplex PCR (sixplex) would need to be adjusted to more than 97.2%. However, it should be noted that, due to the limited availability of sample material, only the DNA of a single specimen or synthetic DNA representing the respective taxa has been tested so far. Further comparative morphological and genetic analyses of individual specimens of respective taxa are needed to clarify the status of C. montanus and C. scoticus clade 2 and correctly interpret the obtained false-positive results of the developed real-time PCR.
The only verified cross-reactivity below the defined cut-off value was observed with the taxon-specific PCR for C. scoticus clade 1 when tested against the DNA of C. newsteadi haplotype N2. Although the individual PCRs were specifically designed, limitations in the specificity of individual molecules can accumulate in the multiplex PCR, resulting in the generation and detection of non-specific amplicons. In this study, the forward primer of C. obsoletus clade O1 and the reverse primer of C. scoticus clade 1 amplified a PCR fragment with the DNA of C. newsteadi haplotype N2, which was detected with the probe of C. scoticus clade 1. Hence, the C. scoticus clade 1 PCR should not be used in combination with the taxon-specific PCR for C. obsoletus clade O1 if unsorted Culicoides are analyzed. This issue highlights the challenges of multiplex development, especially when the targeted DNA region offers limited space and possibilities for positioning several PCRs. The design of sequencing primers capable of amplifying a longer genomic marker and providing greater flexibility for the positioning of primers and probes would be beneficial for future PCR developments. Alternatively, minor groove binding probes could be used in A/T-rich gene regions to enhance PCR specificity [62]. The side-directed insertion of mismatch bases into primers or probes, designed to not interfere with the detection of the target taxon [59,74], was also tested in this study. However, its application in quantitative real-time PCR proved to be challenging due to observed shifts in Ct values and reduced probe signal intensities, which affected the accuracy of genome copy number calculations.
Despite the challenges mentioned, the developed real-time PCR has demonstrated high reliability when testing DNA extracts of single specimens or body parts (thorax, abdomen). The latter is a major advantage over less-sensitive methods such as DNA barcoding, which often require entire midges for the analysis, making morphological re-analysis impossible. The developed multiplex PCR demonstrated successful quantification of its target taxa with a PCR performance comparable to the singleplex PCR approach. The definition of an appropriate cut-off value even allowed the correct determination of the selected target taxa in mixed pools of up to 100 culicoid specimens or bycatch, which is a prerequisite for a high-throughput screening approach needed for surveillance studies. Thus, the developed PCR provides significant potential for monitoring the European vector species and facilitates the implementation of preventive measures to control culicoid-borne diseases such as BTD, Schmallenberg disease, and epidemic hemorrhagic disease of cervids.

5. Conclusions

Culicoid biting midges commonly achieve high population densities, resulting in tremendous numbers of specimens being captured in UV-light traps. This presents a significant challenge in entomological surveillance studies. Isomorphic species of the Obsoletus Group often comprise a substantial proportion of the collected specimens. The real-time PCR developed in this study is the first of its kind capable of efficiently processing such sample sizes while quantifying the relative proportions of all known taxa within the Obsoletus Group in Europe. Although further validation is needed to evaluate the method’s robustness, especially for analyzing pools of unsorted insect catches, the PCR has demonstrated superior efficiency and reliability compared to conventional gel-based PCRs. This advancement marks a significant step toward the routine collection of data on the distribution of ceratopogonid vector species, enabling improved assessment of current and future risk scenarios. Such capabilities are increasingly critical considering the growing globalization and climate change, allowing the timely implementation of countermeasures.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/insects16050500/s1, Table S1: List of DNA extracts used to determine the diagnostic sensitivity (dSe) and specificity (dSp) of the developed real-time PCRs for their different target taxa; Table S2: List of all primers and probes tested for the development of the taxon-specific PCRs; Table S3: Cross-reactivity of the newly developed real-time PCRs against other Culicoides taxa; Table S4: Cross-reactivity of the newly developed real-time PCRs against selected non-culicoid insect taxa.

Author Contributions

Conceptualization, O.D. and H.K.; methodology, O.D., B.H. and H.K.; validation, O.D.; formal analysis, O.D.; investigation, O.D. and H.K.; resources, H.K., D.W. and B.M.; data curation, O.D. and H.K.; writing—original draft preparation, O.D.; writing—review and editing, H.K., D.W., B.H. and B.M.; visualization, O.D.; supervision, H.K.; project administration, H.K. and D.W.; funding acquisition, H.K. and D.W. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the German Federal Ministry of Food and Agriculture (BMEL) through the Federal Office for Agriculture and Food (BLE), grant number 281B101816.

Data Availability Statement

Publicly available datasets used in this study are listed in the Supplementary Material of reference [57]. The respective data can be found in GenBank at: https://www.ncbi.nlm.nih.gov/genbank/ (accessed on 15 November 2022). Data additionally created in this study are deposited in the same data repository with the following accession numbers: Desmometopa sordida (PP110227).

Acknowledgments

We thank our colleagues Sarah Groschupp, Nathalie Richter, and Sebastian Schuran from the Leibniz Centre for Agricultural Landscape Research (ZALF) for the efforts in collecting and morphologically pre-identifying the biting midges used in this study. We also thank Maxi Uecker and Freja Pfirschke for excellent technical assistance regarding the molecular biological work at FLI.

Conflicts of Interest

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

Abbreviations

The following abbreviations are used in this manuscript:
28S rDNAstructural ribosomal RNA (large subunit)
bpbase pairs
BTbaseline threshold
BTDbluetongue disease
BTVbluetongue virus
CADcarbamoyl-phosphate synthetase 2
COImitochondrial cytochrome c oxidase subunit I
Ctcycle threshold
Cy5cyanine5
cytbmitochondrial cytochrome b
DNAdeoxyribonucleic acid
dSediagnostic sensitivity
dSpdiagnostic specificity
EHDepizootic hemorrhagic disease
EHDVepizootic hemorrhagic disease virus
FAM6-carboxyfluorescein
HEXhexachlorofluorescein
HPLCHigh-performance liquid chromatography
ITS-1internal transcribed spacer 1
ITS-2internal transcribed spacer 2
mtmitochondrial
NCBINational Center for Biotechnology Information
PCRpolymerase chain reaction
RFUrelative fluorescence unit
s.s.sensu stricto
SBDSchmallenberg disease
SBVSchmallenberg virus
TEXTexas Red (sulforhodamine 101 acid chloride)

References

  1. Vasconcelos, P.F.; Calisher, C.H. Emergence of human arboviral diseases in the Americas, 2000–2016. Vector Borne Zoonotic Dis. 2016, 16, 295–301. [Google Scholar] [CrossRef] [PubMed]
  2. Catenacci, L.S.; Nunes-Neto, J.; Deem, S.L.; Palmer, J.L.; Travassos-da Rosa, E.S.; Tello, J.S. Diversity patterns of hematophagous insects in Atlantic Forest fragments and human-modified areas of southern Bahia, Brazil. J. Vector Ecol. 2018, 43, 293–304. [Google Scholar] [CrossRef] [PubMed]
  3. Mellor, P.S.; Boorman, J.; Baylis, M. Culicoides biting midges: Their role as arbovirus vectors. Annu. Rev. Entomol. 2000, 45, 307–340. [Google Scholar] [CrossRef] [PubMed]
  4. Carpenter, S.; Groschup, M.H.; Garros, C.; Felippe-Bauer, M.L.; Purse, B.V. Culicoides biting midges, arboviruses and public health in Europe. Antivir. Res. 2013, 100, 102–113. [Google Scholar] [CrossRef]
  5. MacLachlan, N.J. The pathogenesis and immunology of bluetongue virus infection of ruminants. Comp. Immunol. Microbiol. Infect. Dis. 1994, 17, 197–206. [Google Scholar] [CrossRef]
  6. Maclachlan, N.J.; Drew, C.P.; Darpel, K.E.; Worwa, G. The pathology and pathogenesis of bluetongue. J. Comp. Pathol. 2009, 141, 1–16. [Google Scholar] [CrossRef]
  7. Kampen, H.; Werner, D. Biting midges (Diptera: Ceratopogonidae) as vectors of viruses. Microorganisms 2023, 11, 2706. [Google Scholar] [CrossRef]
  8. Coetzee, P.; Stokstad, M.; Venter, E.H.; Myrmel, M.; Van Vuuren, M. Bluetongue: A historical and epidemiological perspective with the emphasis on South Africa. Virol. J. 2012, 9, 198. [Google Scholar] [CrossRef]
  9. Zientara, S.; Sánchez-Vizcaíno, J.M. Control of bluetongue in Europe. Vet. Microbiol. 2013, 165, 33–37. [Google Scholar] [CrossRef]
  10. Alkhamis, M.A.; Aguilar-Vega, C.; Fountain-Jones, N.M.; Lin, K.; Perez, A.M.; Sánchez-Vizcaíno, J.M. Global emergence and evolutionary dynamics of bluetongue virus. Sci. Rep. 2020, 10, 21677. [Google Scholar] [CrossRef]
  11. van den Brom, R.; Santman-Berends, I.; van der Heijden, M.G.; Harders, F.; Engelsma, M.; van Gennip, R.G.P.; Maris-Veldhuis, M.A.; Feddema, A.-J.; Peterson, K.; Golender, N.; et al. Bluetongue virus serotype 12 in sheep and cattle in the Netherlands in 2024–a BTV serotype reported in Europe for the first time. Vet. Microbiol. 2024, 110365. [Google Scholar] [CrossRef] [PubMed]
  12. Holwerda, M.; Santman-Berends, I.; Harders, F.; Engelsma, M.; Vloet, R.; Dijkstra, E.; van Gennip, R.G.P.; Mars, M.H.; Roos, L.; van den Brom, R.; et al. Emergence of bluetongue virus serotype 3, the Netherlands, September 2023. Emerg. Infect. Dis. 2024, 30, 1552–1561. [Google Scholar] [CrossRef] [PubMed]
  13. ProMED-Mail. Bluetongue–Europe (13): Belgium (Antwerp) Serotype 3, Suspected, Request for Information. 9 October 2023: 20231009.8712543. Available online: https://www.promedmail.org/ (accessed on 9 March 2024).
  14. Newbrook, K.; Obishakin, E.; Jones, L.A.; Waters, R.; Ashby, M.; Batten, C.; Sanders, C. Clinical disease in British sheep infected with an emerging strain of bluetongue virus serotype 3. Vet. Rec. 2024, e4910. [Google Scholar] [CrossRef] [PubMed]
  15. Voigt, A.; Kampen, H.; Heuser, E.; Zeiske, S.; Hoffmann, B.; Höper, D.; Holsteg, M.; Sick, F.; Ziegler, S.; Wernike, K.; et al. Bluetongue virus serotype 3 and Schmallenberg virus in Culicoides biting midges, western Germany, 2023. Emerg. Infect. Dis. 2024, 30, 1438–1441. [Google Scholar] [CrossRef]
  16. Ries, C.; Beer, M.; Hoffmann, B. Bluetongue virus infection of goats: Re-emerged European serotype 8 vs. two atypical serotypes. Viruses 2022, 14, 1034. [Google Scholar] [CrossRef]
  17. Subhadra, S.; Sreenivasulu, D.; Pattnaik, R.; Panda, B.K.; Kumar, S. Bluetongue virus: Past, present, and future scope. J. Infect. Dev. Ctries. 2023, 17, 147–156. [Google Scholar] [CrossRef]
  18. van Rijn, P.A. Prospects of next-generation vaccines for bluetongue. Front. Vet. Sci. 2019, 6, 407. [Google Scholar] [CrossRef]
  19. Shope, R.E.; Lester, G.M.; Robert, M. Deer mortality–epizootic hemorrhagic disease of deer. N. J. Outdoors 1955, 6, 17–21. [Google Scholar]
  20. Lorusso, A.; Cappai, S.; Loi, F.; Pinna, L.; Ruiu, A.; Puggioni, G.; Guercio, A.; Purpari, G.; Vicari, D.; Sghaier, S.; et al. Epizootic hemorrhagic disease virus serotype 8, Italy, 2022. Emerg. Infect. Dis. 2023, 29, 1063–1065. [Google Scholar] [CrossRef]
  21. Maan, S.; Maan, N.S.; Ross-smith, N.; Batten, C.A.; Shaw, A.E.; Anthony, S.J.; Samuel, A.R.; Darpel, K.E.; Veronesi, E.; Oura, C.A.; et al. Sequence analysis of bluetongue virus serotype 8 from the Netherlands 2006 and comparison to other European strains. J. Virol. 2008, 377, 308–318. [Google Scholar] [CrossRef] [PubMed]
  22. Bulletins Hebdomadaires de Veille Sanitaire Internationale–Sainté Animale (plateforme-esa.fr). Available online: https://www.plateforme-esa.fr/fr/bulletins-hebdomadaires-de-veille-sanitaire-internationale-du-28-11-2023 (accessed on 21 March 2025).
  23. Hoffmann, B.; Scheuch, M.; Höper, D.; Jungblut, R.; Holsteg, M.; Schirrmeier, H.; Eschbaumer, M.; Goller, K.V.; Wernike, K.; Fischer, M.; et al. Novel orthobunyavirus in cattle, Europe, 2011. Emerg. Infect. Dis. 2012, 18, 469–472. [Google Scholar] [CrossRef] [PubMed]
  24. Wernike, K.; Conraths, F.; Zanella, G.; Granzow, H.; Gache, K.; Schirrmeier, H.; Valas, S.; Staubach, C.; Marianneau, P.; Kraatz, F.; et al. Schmallenberg virus–two years of experiences. Prev. Vet. Med. 2014, 116, 423–434. [Google Scholar] [CrossRef] [PubMed]
  25. Wernike, K.; Beer, M. Schmallenberg virus: A novel virus of veterinary importance. Adv. Virus Res. 2017, 99, 39–60. [Google Scholar] [CrossRef] [PubMed]
  26. Carpenter, S.; Lunt, H.L.; Arav, D.; Venter, G.J.; Mellor, P.S. Oral susceptibility to bluetongue virus of Culicoides (Diptera: Ceratopogonidae) from the United Kingdom. J. Med. Entomol. 2006, 43, 73–78. [Google Scholar] [CrossRef]
  27. Carpenter, S.; McArthur, C.; Selby, R.; Ward, R.; Nolan, D.V.; Luntz, A.J.; Dallas, J.F.; Tripet, F.; Mellor, P.S. Experimental infection studies of UK Culicoides species midges with bluetongue virus serotypes 8 and 9. Vet. Rec. 2008, 163, 589–592. [Google Scholar] [CrossRef]
  28. Federici, V.; Ippoliti, C.; Goffredo, M.; Catalani, M.; Di Provvido, A.; Santilli, A.; Quaglia, M.; Mancini, G.; Di Nicola, F.; Di Gennaro, A.; et al. Epizootic haemorrhagic disease in Italy: Vector competence of indigenous Culicoides species and spatial multicriteria evaluation of vulnerability. Vet. Ital. 2016, 52, 271–279. [Google Scholar] [CrossRef]
  29. Maurer, L.M.; Paslaru, A.; Torgerson, P.R.; Veronesi, E.; Mathis, A. Vector competence of Culicoides biting midges from Switzerland for African horse sickness virus and epizootic haemorrhagic disease virus. Schweiz. Arch. Tierheilkd. 2022, 164, 66–70. [Google Scholar] [CrossRef]
  30. Meiswinkel, R.; van Rijn, P.; Leijs, P.; Goffredo, M. Potential new Culicoides vector of bluetongue virus in northern Europe. Vet. Rec. 2007, 161, 564–565. [Google Scholar] [CrossRef]
  31. Goffredo, M.; Catalani, M.; Federici, V.; Portanti, O.; Marini, V.; Mancini, G.; Quaglia, M.; Santilli, A.; Teodori, L.; Savini, G. Vector species of Culicoides midges implicated in the 2012–2014 bluetongue epidemics in Italy. Vet. Ital. 2015, 51, 131–138. [Google Scholar] [CrossRef]
  32. Dijkstra, E.; van der Ven, I.J.K.; Meiswinkel, R.; Hölzel, D.R.; van Rijn, P.A. Culicoides chiopterus as a potential vector of bluetongue virus in Europe. Vet. Rec. 2008, 162, 422. [Google Scholar] [CrossRef]
  33. Mehlhorn, H.; Walldorf, V.; Klimpel, S.; Jahn, B.; Jaeger, F.; Eschweiler, J.; Hoffmann, B.; Beer, M. First occurrence of Culicoides obsoletus-transmitted bluetongue virus epidemic in Central Europe. Parasitol. Res. 2007, 101, 219–228. [Google Scholar] [CrossRef] [PubMed]
  34. Hoffmann, B.; Bauer, B.; Bauer, C.; Bätza, H.J.; Beer, M.; Clausen, P.H.; Geier, M.; Gethmann, J.M.; Kiel, E.; Liebisch, G.; et al. Monitoring of putative vectors of bluetongue virus serotype 8, Germany. Emerg. Infect. Dis. 2009, 15, 1481–1484. [Google Scholar] [CrossRef] [PubMed]
  35. Foxi, C.; Delrio, G.; Falchi, G.; Marche, M.G.; Satta, G.; Ruiu, L. Role of different Culicoides vectors (Diptera: Ceratopogonidae) in bluetongue virus transmission and overwintering in Sardinia (Italy). Parasit. Vectors 2016, 9, 440. [Google Scholar] [CrossRef] [PubMed]
  36. Rasmussen, L.D.; Kristensen, B.; Kirkeby, C.; Rasmussen, T.B.; Belsham, G.J.; Bødker, R.; Bøtner, A. Culicoids as vectors of Schmallenberg virus. Emerg. Infect. Dis. 2012, 18, 1204–1206. [Google Scholar] [CrossRef]
  37. de Regge, N.; Deblauwe, I.; de Deken, R.; Vantieghem, P.; Madder, M.; Geysen, D.; Smeets, F.; Losson, B.; van den Berg, T.; Cay, A.B. Detection of Schmallenberg virus in different Culicoides spp. by real-time RT-PCR. Transbound. Emerg. Dis. 2012, 59, 471–475. [Google Scholar] [CrossRef]
  38. Elbers, A.R.W.; Meiswinkel, R.; van Weezep, E.; van Sloet Oldruitenborgh-Oosterbaan, M.M.; Kooi, E.A. Schmallenberg virus in Culicoides spp. biting midges, The Netherlands, 2011. Emerg. Infect. Dis. 2013, 19, 106–109. [Google Scholar] [CrossRef]
  39. Goffredo, M.; Monaco, F.; Capelli, G.; Quaglia, M.; Federici, V.; Catalani, M.; Montarsi, F.; Polci, A.; Pinoni, C.; Calistri, P.; et al. Schmallenberg virus in Italy: A retrospective survey in Culicoides stored during the bluetongue Italian surveillance program. Prev. Vet. Med. 2013, 111, 230–236. [Google Scholar] [CrossRef]
  40. Larska, M.; Polak, M.P.; Grochowska, M.; Lechowski, L.; Związek, J.S.; Zmudziński, J.F. First report of Schmallenberg virus infection in cattle and midges in Poland. Transbound. Emerg. Dis. 2013, 60, 97–101. [Google Scholar] [CrossRef]
  41. Larska, M.; Lechowski, L.; Grochowska, M.; Żmudziński, J.F. Detection of the Schmallenberg virus in nulliparous Culicoides obsoletus/scoticus complex and C. punctatus—The possibility of transovarial virus transmission in the midge population and of a new vector. Vet. Microbiol. 2013, 166, 467–473. [Google Scholar] [CrossRef]
  42. Balenghien, T.; Pagès, N.; Goffredo, M.; Carpenter, S.; Augot, D.; Jacquier, E.; Talavera, S.; Monaco, F.; Depaquit, J.; Grillet, C.; et al. The emergence of Schmallenberg virus across Culicoides communities and ecosystems in Europe. Prev. Vet. Med. 2014, 116, 360–369. [Google Scholar] [CrossRef]
  43. Rasmussen, L.D.; Kirkeby, C.; Bødker, R.; Kristensen, B.; Rasmussen, T.B.; Belsham, G.J.; Bøtner, A. Rapid spread of Schmallenberg virus-infected biting midges (Culicoides spp.) across Denmark in 2012. Transbound. Emerg. Dis. 2014, 61, 12–16. [Google Scholar] [CrossRef] [PubMed]
  44. Elbers, A.R.W.; Meiswinkel, R.; van Weezep, E.; Kooi, E.A.; van der Poel, W.H.M. Schmallenberg virus in Culicoides biting midges in the Netherlands in 2012. Transbound. Emerg. Dis. 2015, 62, 339–342. [Google Scholar] [CrossRef] [PubMed]
  45. de Regge, N.; de Deken, R.; Fassotte, C.; Losson, B.; Deblauwe, I.; Madder, M.; Vantieghem, P.; Tomme, M.; Smeets, F.; Cay, A.B. Culicoides monitoring in Belgium in 2011: Analysis of spatiotemporal abundance, species diversity and Schmallenberg virus detection. Med. Vet. Entomol. 2015, 29, 263–275. [Google Scholar] [CrossRef] [PubMed]
  46. Pagès, N.; Talavera, S.; Verdún, M.; Pujol, N.; Valle, M.; Bensaid, A.; Pujols, J. Schmallenberg virus detection in Culicoides biting midges in Spain: First laboratory evidence for highly efficient infection of Culicoides of the Obsoletus Complex and Culicoides imicola. Transbound. Emerg. Dis. 2018, 65, e1–e6. [Google Scholar] [CrossRef]
  47. Ségard, A.; Gardès, L.; Jacquier, E.; Grillet, C.; Mathieu, B.; Rakotoarivony, I.; Setier-Rio, M.-L.; Chavernac, D.; Cêtre-Sossah, C.; Balenghien, T.; et al. Schmallenberg virus in Culicoides Latreille (Diptera: Ceratopogonidae) populations in France during 2011–2012 outbreak. Transbound. Emerg. Dis. 2018, 65, e94–e103. [Google Scholar] [CrossRef]
  48. Quaglia, M.; Foxi, C.; Satta, G.; Puggioni, G.; Bechere, R.; De Ascentis, M.; D’Alessio, S.G.; Spedicato, M.; Leone, A.; Pisciella, M.; et al. Culicoides species responsible for the transmission of epizootic haemorrhagic disease virus (EHDV) serotype 8 in Italy. Vet. Ital. 2023, 59, 83–89. [Google Scholar] [CrossRef]
  49. Nielsen, S.A.; Kristensen, M. Morphological and molecular identification of species of the Obsoletus Group (Diptera: Ceratopogonidae) in Scandinavia. Parasitol. Res. 2011, 109, 1133–1141. [Google Scholar] [CrossRef]
  50. Goffredo, M.; Meiswinkel, R.; Federici, V.; Di Nicola, F.; Mancini, G.; Ippoliti, C.; Di Lorenzo, A.; Quaglia, M.; Santilli, A.; Conte, A.; et al. The ‘Culicoides obsoletus group’ in Italy: Relative abundance, geographic range, and role as vector for bluetongue virus. Vet. Ital. 2016, 52, 235–241. [Google Scholar] [CrossRef]
  51. Schwenkenbecher, J.M.; Mordue, A.J.; Piertney, S.B. Phylogenetic analysis indicates that Culicoides dewulfi should not be considered part of the Culicoides obsoletus complex. Bull. Entomol. Res. 2009, 99, 371–375. [Google Scholar] [CrossRef]
  52. Ander, M.; Troell, K.; Chirico, J. Barcoding of biting midges in the genus Culicoides: A tool for species determination. Med. Vet. Entomol. 2013, 27, 323–331. [Google Scholar] [CrossRef]
  53. Sarvašová, A.; Kočišová, A.; Halán, M.; Delécolle, J.-C.; Mathieu, B. Morphological and molecular analysis of the genus Culicoides (Diptera: Ceratopogonidae) in Slovakia with five new records. Zootaxa 2014, 3872, 541–560. [Google Scholar] [CrossRef] [PubMed]
  54. Augot, D.; Mathieu, B.; Hadj-Henni, L.; Barriel, V.; Zapata, M.S.; Smolis, S.; Augot, D.; Mathieu, B.; Hadj-Henni, L.; Barriel, V.; et al. Molecular phylogeny of 42 species of Culicoides (Diptera, Ceratopogonidae) from three continents. Parasite 2017, 24, 23. [Google Scholar] [CrossRef] [PubMed]
  55. Mathieu, B.; Garros, C.; Balenghien, T.; Candolfi, E.; Delécolle, J.-C.; Cêtre-Sossah, C. A phylogenetic analysis of the biting midges belonging to Culicoides Latreille (Diptera: Ceratopogonidae) subgenus Avaritia using molecular data. Parasit. Vectors 2020, 13, 243. [Google Scholar] [CrossRef] [PubMed]
  56. Hajd-Henni, L.; Sauvage, F.; Ninio, C.; Depaquit, J.; Augot, D. Wing geometry as a tool for discrimination of Obsoletus Group (Diptera: Ceratopogonidae: Culicoides) in France. Infect. Genet. Evol. 2014, 21, 110–117. [Google Scholar] [CrossRef]
  57. Mathieu, B.; Perrin, A.; Baldet, T.; Delécolle, J.-C.; Albina, E.; Cêtre-Sossah, C. Molecular identification of western European species of Obsoletus Complex (Diptera: Ceratopogonidae) by an internal transcribed spacer-1 rDNA multiplex polymerase chain reaction assay. J. Med. Entomol. 2007, 44, 1019–1025. [Google Scholar] [CrossRef]
  58. Mignotte, A.; Garros, C.; Gardès, L.; Balenghien, T.; Duhayon, M.; Rakotoarivony, I.; Tabourin, L.; Poujol, L.; Mathieu, B.; Ibañez-Justicia, A.; et al. The tree that hides the forest: Cryptic diversity and phylogenetic relationships in the Palaearctic vector Obsoletus/Scoticus Complex (Diptera: Ceratopogonidae) at the European level. Parasit. Vectors 2020, 13, 265. [Google Scholar] [CrossRef]
  59. Dähn, O.; Werner, D.; Mathieu, B.; Kampen, H. Large-scale cytochrome c oxidase subunit I gene data analysis for the development of a multiplex polymerase chain reaction test capable of identifying biting midge vector species and haplotypes (Diptera: Ceratopogonidae) of the Culicoides subgenus Avaritia Fox, 1955. Genes 2024, 15, 323. [Google Scholar] [CrossRef]
  60. Kiehl, E.; Walldorf, V.; Klimpel, S.; Al-Quraishy, S.; Mehlhorn, H. The European vectors of bluetongue virus: Are there species complexes, single species or races in Culicoides obsoletus and C. pulicaris detectable by sequencing ITS-1, ITS-2 and 18S-rDNA? Parasitol. Res. 2009, 105, 331–336. [Google Scholar] [CrossRef]
  61. Meiswinkel, R.; De Bree, F.; Bossers-de Vries, R.; Elbers, A.R. An unrecognized species of the Culicoides obsoletus complex feeding on livestock in the Netherlands. Vet. Parasitol. 2015, 207, 324–328. [Google Scholar] [CrossRef]
  62. Wenk, C.E.; Kaufmann, C.; Schaffner, F.; Mathis, A. Molecular characterization of Swiss Ceratopogonidae (Diptera) and evaluation of real-time PCR assays for the identification of Culicoides biting midges. Vet. Parasitol. 2012, 184, 258–266. [Google Scholar] [CrossRef]
  63. Elbers, A.R.W.; Meiswinkel, R. Culicoides (Diptera: Ceratopogonidae) host preferences and biting rates in the Netherlands: Comparing cattle, sheep and the black-light suction trap. Vet. Parasitol. 2014, 205, 330–337. [Google Scholar] [CrossRef] [PubMed]
  64. Zimmer, J.Y.; Brostaux, Y.; Haubruge, E.; Francis, F. Larval development sites of the main Culicoides species (Diptera: Ceratopogonidae) in northern Europe and distribution of coprophilic species larvae in Belgian pastures. Vet. Parasitol. 2014, 205, 676–686. [Google Scholar] [CrossRef] [PubMed]
  65. Ninio, C.; Augot, D.; Delecolle, J.C.; Dufour, B.; Depaquit, J. Contribution to the knowledge of Culicoides (Diptera: Ceratopogonidae) host preferences in France. Parasitol. Res. 2011, 108, 657–663. [Google Scholar] [CrossRef] [PubMed]
  66. Garros, C.; Gardès, L.; Allène, X.; Rakotoarivony, I.; Viennet, E.; Rossi, S.; Balenghien, T. Adaptation of a species-specific multiplex PCR assay for the identification of blood meal source in Culicoides (Ceratopogonidae: Diptera): Applications on Palaearctic biting midge species, vectors of Orbiviruses. Infect. Genet. Evol. 2011, 11, 1103–1110. [Google Scholar] [CrossRef]
  67. Kettle, D.S.; Lawson, J.W.H. The early Stages of British biting midges Culicoides Latreille (Diptera: Ceratopogonidae) and allied genera. Bull. Entomol. Res. 1952, 43, 421–467. [Google Scholar] [CrossRef]
  68. Gethmann, J.; Probst, C.; Conraths, F.J. Economic impact of a bluetongue serotype 8 epidemic in Germany. Front. Vet. Sci. 2020, 7, 65. [Google Scholar] [CrossRef]
  69. Harrup, L.E.; Bellis, G.A.; Balenghien, T.; Garros, C. Culicoides Latreille (Diptera: Ceratopogonidae) taxonomy: Current challenges and future directions. Infect. Genet. Evol. 2015, 30, 249–266. [Google Scholar] [CrossRef]
  70. Pagès, N.; Sarto i Monteys, V. Differentiation of Culicoides obsoletus and Culicoides scoticus (Diptera: Ceratopogonidae) based on mitochondrial cytochrome oxidase subunit I. J. Med. Entomol. 2005, 42, 1026–1034. [Google Scholar] [CrossRef]
  71. Deblauwe, I.; de Witte, J.C.; de Deken, G.; de Deken, R.; Madder, M.; van Erk, S.; Hoza, F.A.; Lathouwers, D.; Geysen, D. A new tool for the molecular identification of Culicoides species of the Obsoletus group: The glass slide microarray approach. Med. Vet. Entomol. 2012, 26, 83–91. [Google Scholar] [CrossRef]
  72. Kaufmann, C.; Steinmann, I.C.; Hegglin, D.; Schaffner, F.; Mathis, A. Spatio-temporal occurrence of Culicoides biting midges in the climatic regions of Switzerland, along with large scale species identification by MALDI-TOF mass spectrometry. Parasit. Vectors 2012, 5, 246. [Google Scholar] [CrossRef]
  73. Mathieu, B.; Delecolle, J.-C.; Garros, C.; Balenghien, T.; Setier-Rio, M.-L.; Candolfi, E.; Cêtre-Sossah, C. Simultaneous quantification of the relative abundance of species complex members: Application to Culicoides obsoletus and Culicoides scoticus (Diptera: Ceratopogonidae), potential vectors of bluetongue virus. Vet. Parasitol. 2011, 182, 297–306. [Google Scholar] [CrossRef] [PubMed]
  74. Dähn, O.; Werner, D.; Mathieu, B.; Kampen, H. Development of conventional multiplex PCR assays for the identification of 21 west Palaearctic biting midge taxa (Diptera: Ceratopogonidae) belonging to the Culicoides subgenus Culicoides, including recently discovered species and genetic variants. Diversity 2023, 15, 699. [Google Scholar] [CrossRef]
  75. Vorsprach, B.; Meiser, C.K.; Werner, D.; Balczun, C.; Schaub, G.A. Monitoring of Ceratopogonidae in southwest Germany. Parasitol. Res. 2009, 105, 337–344. [Google Scholar] [CrossRef]
  76. Schwenkenbecher, J.M.; Mordue, A.J.; Switek, K.; Piertney, S.B. Discrimination of Culicoides midge larvae using multiplex polymerase chain reaction assays based on DNA sequence variation at the mitochondrial cytochrome c oxidase I gene. J. Med. Entomol. 2009, 46, 610–614. [Google Scholar] [CrossRef]
  77. Lehmann, K.; Werner, D.; Hoffmann, B.; Kampen, H. PCR identification of culicoid biting midges (Diptera, Ceratopogonidae) of the Obsoletus Complex including putative vectors of bluetongue and Schmallenberg viruses. Parasit. Vectors 2012, 5, 213. [Google Scholar] [CrossRef]
  78. Monaco, F.; Benedetto, L.; Di Marcello, V.; Lelli, R.; Goffredo, M. Development and preliminary evaluation of a real-time polymerase chain reaction for the identification of Culicoides obsoletus sensu stricto, C. scoticus and C. montanus in the Obsoletus Complex in Italy. Vet. Ital. 2010, 46, 215–220. [Google Scholar]
  79. Delécolle, J.-C. Nouvelle Contribution à L’étude Systématique et Iconographique des Espèces du Genre Culicoides (Diptera: Ceratopogonidae) du Nord-Est de la France. Ph.D. Thesis, University of Strasbourg, Strasbourg, France, 1985. [Google Scholar]
  80. Campbell, J.A.; Pelham-Clinton, E.C. A taxonomic review of the British species of Culicoides Latreille (Diptera, Ceratopogonidae). Proc. R. Soc. Edinb. Sect. B Biol. Sci. 1960, 67, 181–302. [Google Scholar] [CrossRef]
  81. Glukhova, V.M. Krovososuščie Mokrecy Rodov Culicoides i Forcipomyia (Ceratopogonidae); Nauka: Leningrad, Russia, 1989; ISBN 9785020257603. [Google Scholar]
  82. Mathieu, B.; Cêtre-Sossah, C.; Garros, C.; Chavernac, D.; Balenghien, T.; Carpenter, S.; Setier-Rio, M.L.; Vignes-Lebbe, R.; Ung, V.; Candolfi, E.; et al. Development and validation of IIKC: An interactive identification key for Culicoides (Diptera: Ceratopogonidae) females from the western Palaearctic region. Parasit. Vectors 2012, 5, 137. [Google Scholar] [CrossRef]
  83. Ries, C.; Sharav, T.; Tseren-Ochir, E.-O.; Beer, M.; Hoffmann, B. Putative novel serotypes ‘33’ and ‘35’ in clinically healthy small ruminants in Mongolia expand the group of atypical BTV. Viruses 2021, 13, 42. [Google Scholar] [CrossRef]
  84. Rodríguez, A.; Rodríguez, M.; Córdoba, J.J.; Andrade, M.J. Design of primers and probes for quantitative real-time PCR methods. Methods Mol. Biol. 2015, 1275, 31–56. [Google Scholar] [CrossRef]
  85. Bustin, S.A.; Mueller, R.; Nolan, T. Parameters for successful PCR primer design. Methods Mol. Biol. 2020, 2065, 5–22. [Google Scholar] [CrossRef] [PubMed]
  86. Borkent, A.; Dominiak, P. Catalog of the biting midges of the world (Diptera: Ceratopogonidae). Zootaxa 2020, 4787, 1–377. [Google Scholar] [CrossRef] [PubMed]
  87. Garros, C.; Balenghien, T.; Carpenter, S.; Delécolle, J.-C.; Meiswinkel, R.; Pédarrieu, A.; Rakotoarivony, I.; Gardès, L.; Golding, N.; Barber, J.; et al. Towards the PCR-based identification of Palaearctic Culicoides biting midges (Diptera: Ceratopogonidae): Results from an international ring trial targeting four species of the subgenus Avaritia. Parasit. Vectors 2014, 7, 223. [Google Scholar] [CrossRef] [PubMed]
  88. Nolan, D.V.; Carpenter, S.; Barber, J.; Mellor, P.S.; Dallas, J.F.; Mordue Luntz, A.J.; Piertney, S.B. Rapid diagnostic PCR assays for members of the Culicoides obsoletus and Culicoides pulicaris species complexes, implicated vectors of bluetongue virus in Europe. Vet. Microbiol. 2007, 124, 82–94. [Google Scholar] [CrossRef]
  89. Gomulski, L.M.; Meiswinkel, R.; Delécolle, J.C.; Goffredo, M.; Gasperi, G. Phylogenetic relationships of the subgenus Avaritia Fox, 1955 including Culicoides obsoletus (Diptera, Ceratopogonidae) in Italy based on internal transcribed spacer 2 ribosomal DNA sequences. Syst. Entomol. 2005, 30, 619–631. [Google Scholar] [CrossRef]
  90. New England Biolabs–Guidelines for PCR Optimization with Taq DNA Polymerase. Available online: https://www.neb.com/en/tools-and-resources/usage-guidelines/guidelines-for-pcr-optimization-with-taq-dna-polymerase (accessed on 21 March 2025).
  91. Garibyan, L.; Avashia, N. Polymerase chain reaction. J. Invest. Dermatol. 2013, 133, 1–4. [Google Scholar] [CrossRef]
  92. Life Technologies Real-Time PCR Handbook. Available online: https://www.gene-quantification.de/real-time-pcr-handbook-life-technologies-update-flr.pdf (accessed on 21 March 2025).
  93. Longo, M.C.; Berninger, M.S.; Hartley, J.L. Use of uracil DNA glycosylase to control carry-over contamination in polymerase chain reactions. Gene 1990, 93, 125–128. [Google Scholar] [CrossRef]
  94. Aguilar-Vega, C.; Rivera, B.; Lucientes, J.; Gutiérrez-Boada, I.; Sánchez-Vizcaíno, J.M. A study of the composition of the Obsoletus Complex and genetic diversity of Culicoides obsoletus populations in Spain. Parasit. Vectors. 2021, 14, 351. [Google Scholar] [CrossRef]
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Figure 1. Direct comparison of the PCR performance between the sixplex PCR approach (colored) and the taxon-specific singleplex PCRs (black) through the simultaneous measurement of serial dilutions of the different target DNAs: C. obsoletus clade O1 ((A) blue, Atto647N), C. obsoletus clade O2 ((B) green, Atto647N), C. obsoletus clade O3 ((C) red, Atto647N), C. scoticus clade 1 ((D) orange, FAM), C. chiopterus ((E) pink, TEX), and C. dewulfi ((F) purple, HEX). BT: Baseline threshold in the corresponding detection channel. Calculated standard curves are given for C. obsoletus clade O1, C. obsoletus clade O2, C. obsoletus clade O3, and C. scoticus clade 1.
Figure 1. Direct comparison of the PCR performance between the sixplex PCR approach (colored) and the taxon-specific singleplex PCRs (black) through the simultaneous measurement of serial dilutions of the different target DNAs: C. obsoletus clade O1 ((A) blue, Atto647N), C. obsoletus clade O2 ((B) green, Atto647N), C. obsoletus clade O3 ((C) red, Atto647N), C. scoticus clade 1 ((D) orange, FAM), C. chiopterus ((E) pink, TEX), and C. dewulfi ((F) purple, HEX). BT: Baseline threshold in the corresponding detection channel. Calculated standard curves are given for C. obsoletus clade O1, C. obsoletus clade O2, C. obsoletus clade O3, and C. scoticus clade 1.
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Figure 2. Determination of PCR cross-reactivity. Each singleplex PCR (A–F) was tested with 1 × 108 copies of DNA of the various taxa (a–f) to simulate pool testing of 100 biting midges each: C. obsoletus clade O1 (PCR A, HEX, DNA a), C. obsoletus clade O2 (PCR B, TEX, DNA b), C. obsoletus clade O3 (PCR C, Atto647N, DNA c), C. scoticus clade 1 (PCR D, FAM, DNA d), C. chiopterus (PCR E, TEX, DNA e), and C. dewulfi (PCR F, HEX, DNA f). BT: Baseline threshold in the corresponding detection channel. RFU: Relative fluorescence units.
Figure 2. Determination of PCR cross-reactivity. Each singleplex PCR (A–F) was tested with 1 × 108 copies of DNA of the various taxa (a–f) to simulate pool testing of 100 biting midges each: C. obsoletus clade O1 (PCR A, HEX, DNA a), C. obsoletus clade O2 (PCR B, TEX, DNA b), C. obsoletus clade O3 (PCR C, Atto647N, DNA c), C. scoticus clade 1 (PCR D, FAM, DNA d), C. chiopterus (PCR E, TEX, DNA e), and C. dewulfi (PCR F, HEX, DNA f). BT: Baseline threshold in the corresponding detection channel. RFU: Relative fluorescence units.
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Table 1. List of specific primers and probes designed for six taxa of the subgenus Avaritia. The PCR approach is designed in a modular manner: All PCRs can be either used alone (singleplex) or combined in different single-tube multiplex approaches (triplex, fourplex, sixplex). The sixplex PCR allows the differentiation of C. obsoletus (all three clades labeled with Atto647N probe), C. scoticus clade 1 (FAM), C. chiopterus (TEX), and C. dewulfi (HEX). For further identification of C. obsoletus positive samples to the clade level, a second PCR must be performed through specific signals of the following fluorophores: C. obsoletus clade O1 (HEX), C. obsoletus clade O2 (TEX), and C. obsoletus clade O3 (Atto647N). If there is no need to determine C. chiopterus and C. dewulfi, the samples can be directly analyzed in a one-step approach either for C. obsoletus clades O1 to O3 (triplex) alone or in combination with C. scoticus (fourplex).
Table 1. List of specific primers and probes designed for six taxa of the subgenus Avaritia. The PCR approach is designed in a modular manner: All PCRs can be either used alone (singleplex) or combined in different single-tube multiplex approaches (triplex, fourplex, sixplex). The sixplex PCR allows the differentiation of C. obsoletus (all three clades labeled with Atto647N probe), C. scoticus clade 1 (FAM), C. chiopterus (TEX), and C. dewulfi (HEX). For further identification of C. obsoletus positive samples to the clade level, a second PCR must be performed through specific signals of the following fluorophores: C. obsoletus clade O1 (HEX), C. obsoletus clade O2 (TEX), and C. obsoletus clade O3 (Atto647N). If there is no need to determine C. chiopterus and C. dewulfi, the samples can be directly analyzed in a one-step approach either for C. obsoletus clades O1 to O3 (triplex) alone or in combination with C. scoticus (fourplex).
TaxonOligo Name5′-ModificationSequence (5′ > 3′)3′-ModificationAmplicon Size (bp)SixplexFourplexTriplexSingleplex
C. obsoletus clade O1CobsO1_FnoneAGAAAAYGGRGCAGGAACCnone89xxxx
CobsO1_RnoneAAATAGCCAAATCTACAGAAGnonexxxx
CobsO1_P-AttoAtto647NCCCTCCCTTTCRTCTAATATCTCTBHQ-2x--x
CobsO1_P-HEXHEXCCCTCCCTTTCRTCTAATATCTCTBHQ-1-xxx
C. obsoletus clade O2CobsO2_FnoneAGCCGTTAATTTTATTACAACCnone127xxxx
CobsO2_RnoneTAATACAGGTAAAGATAGYAGGnonexxxx
CobsO2_P-AttoAtto647NACGATCATATGGAATAAMTTTCGATCBHQ-2x--x
CobsO2_P-TEXTEXACGATCATATGGAATAAMTTTCGATCBHQ-2-xxx
C. obsoletus clade O3CobsO3_FnoneGCTCTATTTTAGGTGCTGTTnone107xxxx
CobsO3_RnoneAGTAATTAATACAGATCATACGnonexxxx
CobsO3_P-AttoAtto647NTATTATCAATATRCGATCATACGGGABHQ-2xxxx
C. scoticus clade 1Csco_FnoneCACTTTATTATTAATTAGAAGTTTAGTTnone116xx-x
Csco_RnoneAAATTGCTAAGTCAACTGAGGnonexx-x
Csco_P-FAMFAMACCCTCCACTTTCAGCAAATGTCTBHQ-1xx-x
C. chiopterusCchi_FnoneCCCTGATATAGCTTTTCCAnone86x--x
Cchi_RnoneCTAAGCTACTTAYTAATAGTAGnonex--x
Cchi_P-TEXTEXTGAATACTRCCRCCCTCTATCACCBHQ-2x--x
C. dewulfiCdew_FnoneACAATCATTAATATACGACCAAnone118x--x
Cdew_RnoneTARCTCCTGCTAAAACTGGAnonex--x
Cdew_P-HEXHEXCACAGCTATTCTTTTACTTCTGTCACBHQ-1x--x
x: Applicable. -: Not applicable.
Table 2. Determination of the diagnostic sensitivity (dSe) and specificity (dSp) of the developed real-time PCR for the various biting midge taxa. A total of 91 genetically pre-identified (sequenced) DNA samples of single specimens were initially tested with the sixplex PCR. Samples detected with Atto647N-labeled probe (C. obsoletus) were further analyzed with a triplex PCR to differentiate the three C. obsoletus clades.
Table 2. Determination of the diagnostic sensitivity (dSe) and specificity (dSp) of the developed real-time PCR for the various biting midge taxa. A total of 91 genetically pre-identified (sequenced) DNA samples of single specimens were initially tested with the sixplex PCR. Samples detected with Atto647N-labeled probe (C. obsoletus) were further analyzed with a triplex PCR to differentiate the three C. obsoletus clades.
TaxonSamples Tested (n) 1Sixplex PCR (n) 2Triplex PCR (n) 2dSedSp
C. obsoletus clade O139703939/39 (100%)38/39 (97.4%)
C. obsoletus clade O22829 328/28 (100%)28/28 (100%)
C. obsoletus clade O3333/3 (100%)3/3 (100%)
C. scoticus clade 11010-10/10 (100%)10/10 (100%)
C. chiopterus44-4/4 (100%)4/4 (100%)
C. dewulfi77-7/7 (100%)7/7 (100%)
Total91917191/91 (100%)91/92 (98.9%)
1 Taxon-specific DNA samples of single specimens, pre-identified with COI barcoding. 2 Number of samples with a positive result. Cut-off value: Ct < 30 = positive. 3 OQ941557 was positive for C. obsoletus clade O2 (Ct 25.60) and C. obsoletus clade O1 (Ct 28.56). -: Not applicable.
Table 3. Cross-reactivity of the newly developed real-time PCR against other Culicoides taxa (n = 36). The DNAs of single specimens or 1 × 106 copies of the artificial COI gene of each taxon were tested with the sixplex PCR approach and, if necessary, the triplex PCR to determine the specific clade of C. obsoletus. The lowest Ct values are indicated in brackets and were specified by taxon-specific PCRs. Ct values > 30 (above the cut-off) were considered negative.
Table 3. Cross-reactivity of the newly developed real-time PCR against other Culicoides taxa (n = 36). The DNAs of single specimens or 1 × 106 copies of the artificial COI gene of each taxon were tested with the sixplex PCR approach and, if necessary, the triplex PCR to determine the specific clade of C. obsoletus. The lowest Ct values are indicated in brackets and were specified by taxon-specific PCRs. Ct values > 30 (above the cut-off) were considered negative.
SubgenusTaxonGenBank Accession No.Real-Time PCR Result
Avaritia Fox, 1955C. imicola2OQ789072negative (Ct 36.79) 4
C. montanus2OQ789074positive (Ct 19.54) 5
C. sanguisuga2MK760238negative
C. scoticus clade 2 2OQ789084positive (Ct 19.97) 6
C. sinanoensis2MK760244negative
Beltranmyia Vargas, 1953C. salinarius2OQ789083negative
Culicoides Latreille, 1809C. boyi3-negative
C. bysta3-negative
C. cryptipulicaris3-negative
C. delta2OQ789035negative
C. fagineus haplotype F1 3-negative
C. fagineus haplotype F2 2OQ789036negative
C. flavipulicaris3-negative
C. grisescens haplotype G1 2OQ789037negative (Ct 38.25) 7
C. grisescens haplotype G2 2OQ789038negative (Ct 36.27) 7
C. kalix3-negative
C. lupicaris haplotype L1 2OQ789039negative
C. lupicaris haplotype L2 2OQ789041negative
C. newsteadi s.s. 3-negative (Ct 32.40) 6
C. newsteadi haplotype N1 2OQ789045negative (Ct 36.09) 4
C. newsteadi haplotype N2 3-positive (Ct 27.12) 6
C. newsteadi haplotype N3 2OQ789048negative
C. pulicaris 2OQ789058negative
C. punctatus2OQ789064negative
C. selandicus2OQ789052negative
C. subfagineus3-negative
Monoculicoides Khalaf, 1954C. riethi2OQ789081negative
Sensiculicoides Shevchenko, 1977C. alazanicus2OQ789067negative
C. festivipennis2OQ789070negative (Ct 34.85) 7
C. griseidorsum2OQ789071negative
C. kibunensis2OQ789073negative
C. pictipennis2OQ789079negative (Ct 34.72) 7
C. poperinghensis2OQ789080negative
Silvaticulicoides Glukhova, 1977C. achrayi2OQ789066negative
Wirthomyia Vargas, 1973 1C. riouxi2OQ789082negative
UnplacedC. pallidicornis2OQ789078negative (Ct 34.63) 4
dSp (other Culicoides taxa):n = 3633/36 (91.7%)
1 According to the latest world catalog of biting midges [86], this species is unplaced, but—based on male and female morphology—should be assigned to the subgenus Wirthomyia. 2 DNA extracts of single specimens. 3 1 × 106 copies of the artificial COI gene [74]. 4 C. dewulfi (HEX) in sixplex PCR. 5 C. obsoletus clade O1 (HEX) in triplex PCR. 6 C. scoticus clade 1 (FAM) in sixplex PCR. 7 C. obsoletus (Atto647N) in sixplex PCR. -: Not applicable.
Table 4. Cross-reactivity of the newly developed real-time PCR against non-culicoid ‘dipteran species’ (n = 15). The DNA extract of one specimen each, representing either a species or a genus, was tested. All samples were analyzed with the sixplex PCR approach. Ct values are indicated in brackets and were specified by taxon-specific PCRs. Ct values > 30 (above the cut-off) were considered negative.
Table 4. Cross-reactivity of the newly developed real-time PCR against non-culicoid ‘dipteran species’ (n = 15). The DNA extract of one specimen each, representing either a species or a genus, was tested. All samples were analyzed with the sixplex PCR approach. Ct values are indicated in brackets and were specified by taxon-specific PCRs. Ct values > 30 (above the cut-off) were considered negative.
Taxon/GenusGenBank Accession No.Real-Time PCR Result
Alluaudomyia spec.PP110213negative
Camptocladius stercorariusPP110214negative
Chironomus lugubrisPP110215negative
Clogmia albipunctataPP110216negative (Ct 36.40) 1
Desmometopa sordidaPP110227negative
Forcipomyia spec.PP110217negative (Ct 35.83) 1
Nemotelus notatusPP110218negative
Nilotanypus dubiusPP110219negative
Physiphora alceaePP110220negative
Psychoda cinereaPP110221negative
Sepsis violaceaPP110222negative
Smittia spec.PP110223negative
Spelobia luteilabrisPP110224negative
Sphaerocera curvipesPP110225negative
Tephrochlamys rufiventrisPP110226negative (Ct 36.89) 2
dSp (other Diptera taxa):n = 1515/15 (100%)
1 C. dewulfi (HEX). 2 C. obsoletus (Atto647N).
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Dähn, O.; Hoffmann, B.; Werner, D.; Mathieu, B.; Kampen, H. Development of Quantitative Real-Time PCR Tests for the Identification of Biting Midge Species and Clades (Diptera: Ceratopogonidae) of the Obsoletus Group (Subgenus Avaritia), Including Important Viral Vectors in Europe. Insects 2025, 16, 500. https://doi.org/10.3390/insects16050500

AMA Style

Dähn O, Hoffmann B, Werner D, Mathieu B, Kampen H. Development of Quantitative Real-Time PCR Tests for the Identification of Biting Midge Species and Clades (Diptera: Ceratopogonidae) of the Obsoletus Group (Subgenus Avaritia), Including Important Viral Vectors in Europe. Insects. 2025; 16(5):500. https://doi.org/10.3390/insects16050500

Chicago/Turabian Style

Dähn, Oliver, Bernd Hoffmann, Doreen Werner, Bruno Mathieu, and Helge Kampen. 2025. "Development of Quantitative Real-Time PCR Tests for the Identification of Biting Midge Species and Clades (Diptera: Ceratopogonidae) of the Obsoletus Group (Subgenus Avaritia), Including Important Viral Vectors in Europe" Insects 16, no. 5: 500. https://doi.org/10.3390/insects16050500

APA Style

Dähn, O., Hoffmann, B., Werner, D., Mathieu, B., & Kampen, H. (2025). Development of Quantitative Real-Time PCR Tests for the Identification of Biting Midge Species and Clades (Diptera: Ceratopogonidae) of the Obsoletus Group (Subgenus Avaritia), Including Important Viral Vectors in Europe. Insects, 16(5), 500. https://doi.org/10.3390/insects16050500

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