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Real-time quantitative PCR assay development and application for assessment of agricultural surface water and various fecal matter for prevalence of Aliarcobacter faecis and Aliarcobacter lanthieri



Aliarcobacter faecis and Aliarcobacter lanthieri are recently identified as emerging human and animal pathogens. In this paper, we demonstrate the development and optimization of two direct DNA-based quantitative real-time PCR assays using species-specific oligonucleotide primer pairs derived from rpoB and gyrA genes for A. faecis and A. lanthieri, respectively. Initially, the specificity of primers and amplicon size of each target reference strain was verified and confirmed by melt curve analysis. Standard curves were developed with a minimum quantification limit of 100 cells mL− 1 or g− 1 obtained using known quantities of spiked A. faecis and A. lanthieri reference strains in autoclaved agricultural surface water and dairy cow manure samples.


Each species-specific qPCR assay was validated and applied to determine the rate of prevalence and quantify the total number of cells of each target species in natural surface waters of an agriculturally-dominant and non-agricultural reference watershed. In addition, the prevalence and densities were determined for human and various animal (e.g., dogs, cats, dairy cow, and poultry) fecal samples. Overall, the prevalence of A. faecis for surface water and feces was 21 and 28%, respectively. The maximum A. faecis concentration for water and feces was 2.3 × 107 cells 100 mL- 1 and 1.2 × 107 cells g− 1, respectively. A. lanthieri was detected at a lower frequency (2%) with a maximum concentration in surface water of 4.2 × 105 cells 100 mL− 1; fecal samples had a prevalence and maximum density of 10% and 2.0 × 106 cells g− 1, respectively.


The results indicate that the occurrence of these species in agricultural surface water is potentially due to fecal contamination of water from livestock, human, or wildlife as both species were detected in fecal samples. The new real-time qPCR assays can facilitate rapid and accurate detection in < 3 h to quantify total numbers of A. faecis and A. lanthieri cells present in various complex environmental samples.


  1. 1.

    Novel qPCR assays for A. faecis and A. lanthieri

  2. 2.

    Identifying prevalence of Aliarcobacter spp. in environmental samples

  3. 3.

    Quantitation of A. faecis and A. lanthieri in water and feces


Recently, the Arcobacter genus has been reclassified and divided into seven new genera where novel genus Aliarcobacter consists of eight species [1]. Of these Aliarcobacter species, A. faecis and A. lanthieri, isolated from human and animal fecal sources, have been identified as pathogenic bacteria [2,3,4]. Some of the most prevalent Aliarcobacter species including A. butzleri, A. cryaerophilus, and A. skirrowii have been identified as causative agents for human (e.g., gastroenteritis, bacteremia, and sepsis) and animal (e.g., mastitis, diarrhea, abortion, and reproductive disorders) infections [5]. These species have also  been isolated from a variety of food products, including chicken, beef, pork, and shellfish as well as various aquatic sources [6,7,8,9,10] and pose an important risk for human infection from contamination of water and food resulting from a variety of sources including livestock and poultry wastes, agricultural runoff, septic leakages, and direct or indirect inputs of wildlife fecal matter [11,12,13,14]. Therefore, there is a need to determine the degree of prevalence and identify potential sources of contamination of A. faecis and A. lanthieri in various fecal and aquatic niches.

Conventional culture-based methods for the identification and quantification of bacterial species from potentially contaminated environmental samples are traditionally slow and cannot be used for the identification of genus Aliarcobacter to the species-level [15]. Biochemical tests for the correct differentiation of Aliarcobacter spp., including A. faecis and A. lanthieri are difficult to differentiate using fastidious selective growth conditions, especially when they are present in low concentrations and in competition with other contaminants [5, 16]. Moreover, these methods may not be accurate enough to measure cell viability, as cells may enter into viable but non-culturable (VBNC) or non-viable and non-culturable (NVNC) states. For many situations, it is important to use techniques that can quantify the total number of cells, including VBNC and NVNC states, more accurately in various complex environmental niches. Non-viable or non-culturable cells of Gram-negative bacteria can potentially contaminate water by producing virulence-associated factors and toxins that can pose health risks to humans [17, 18].

Real-time quantitative PCR (qPCR) assays have provided more rapid and robust tools to detect and quantify Aliarcobacter spp. in pure culture, fecal, hide, food, and complex environmental samples [19,20,21,22]. None of these developed real-time qPCR assays were capable of differentiating and quantifying A. faecis and A. lanthieri directly from environmental matrices, partly because of their unknown status and/or low abundance in these niches.

Therefore, it is necessary to develop fast and accurate methods for detecting these species in complex environmental matrices, since conventional methods are not always accurate measures for recovery and may fail to detect these species when prevalent at low concentrations and when competing with other Aliarcobacter spp. The main objectives of this study  were to: i) develop and optimize species-specific direct real-time qPCR assays to quantitatively detect A. faecis and A. lanthieri in environmental niches; and ii) validate and apply these qPCR assays to detect, quantify, and assess the prevalence of A. faecis and A. lanthieri in agricultural surface water and fecal samples from human and animal sources.


Optimization of species-specific real-time qPCR assays and development of standard curves

Two novel real-time PCR assays were developed and optimized using A. faecis LMG 28519 and A. lanthieri LMG 28516 reference strains. The assays were further validated by applying to the field isolates of each target species (A. faecis: n = 29; and A. lanthieri: n = 10). The primers specifically amplified to their target sequences with expected melting peaks of 74 °C for A. lanthieri and 79 °C for A. faecis (Fig. 1A and B) and typical amplicon sizes 152 bp and 72 bp, respectively (Figure S1A and B). Moreover, no amplification signals were observed for any other Aliarcobacter spp. or other bacterial reference species and strains that could potentially occur in water and fecal matter (Table 1).

Fig. 1

The melting curves generated for the typical amplicons showed specific relative intensity of 79 °C for A. faecis (rpoB gene) (Panel A) and 74 °C for A. lanthieri (gyrA gene) (Panel B) reference strains and field isolates against -ve controls

Table 1 List of reference strains of target and other bacterial species and strains used in this study

The limit of detection for quantitative analysis of each optimized real-time PCR assay was determined by developing standard curves of reference strains of A. faecis and A. lanthieri DNA templates extracted from spiked water and dairy cow manure samples, in units of number of cells mL− 1 (Fig. 2A and B) and cells g− 1 (Fig. 3A and B). Although a minimum of 10 cells mL− 1 or 10 cells g− 1 were also used for the quantitative assay, amplification was observed at ≥40 Cq value; therefore, Cq value ≥42 thresholds were considered as negative or indeterminate.

Fig. 2

Real-time qPCR-based standard curves developed for (Panel A) A. faecis and (Panel B) A. lanthieri from 10-fold serially diluted spiked cells (from 102 to 108 mL− 1) in autoclaved agricultral surface water using species-specific qPCR assays. Each point representing the result of triplicate amplification of each dilution. The coefficients of determination R2 and the slopes of each regression curves are indicated. The standard curves obtained by correlation of the Quantification Cycle (Cq) values and log10 cell number from the amplification plot

Fig. 3

Standard curves developed from DNA extracted from 10-fold serially diluted spiked cells of A. faecis (Panel A) (ranging from 102 to 108 g− 1) and A. lanthieri (Panel B) (ranging from 102 to 107 g- 1) in cow manure using species-specific qPCR assays. The standard curves obtained from amplification plot by correlation of the Quantification Cycle (Cq) cycle values and number of cells g− 1 of feces. Each point is representing the result of duplicate amplification of each dilution where the correlation coefficients R2 and the slopes of the regression curve are shown

qPCR assay validation and application for detection and quantitation of A. faecis and A. lanthieri in agricultural surface water and fecal sources

The qPCR assays were further validated and applied by analysing a total of 804 environmental (fecal and surface water) samples. Of the total 588 agricultural surface water samples, A. faecis was detected at a frequency of 21% (n = 124), while A. lanthieri (n = 13) was only detected in 2% of samples. Similarly, of the total 216 (human, n = 19; animals, n = 197) fecal samples, A. faecis (n = 61) was detected more commonly (28%) than A. lanthieri (10%; n = 22).

Further comparative analyses showed that the rate of A. faecis prevalence was significantly higher (p < 0.05) than A. lanthieri in agricultural sites (Table 2). Interestingly, only A. faecis (19% frequency), not A. lanthieri, was detected at the reference sampling site 24 (Table 2). Overall, the frequency of co-occurrence of these two target species was low and observed in only one single surface water sample, which was collected from an agricultural drainage ditch with upstream proximity to dairy livestock operations. Moreover, among the 11 agriculturally impacted sites, A. faecis was detected at a significantly (p < 0.05) higher frequency (> 20%) at sampling sites 5, 6, 10, 18 and 20 as compared to sampling sites 1, 8, 9, 19, 21 and 253 (< 20%). There was no significance (p > 0.05) difference  in the occurrence of A. lanthieri among the sites.

Table 2 Total number (588) of surface water samples collected from the South Nation River watershed tested for detection and prevalence of A. faecis and A. lanthieri

Of the total 216 fecal samples collected from human and various animal fecal sources, 28% (n = 61) and 10% (n = 22) samples were positive for A. faecis and A. lanthieri, respectively. Among these different fecal samples, A. faecis was detected at higher frequencies in human, cat, cow, dog, and pig, compared to A. lanthieri which was detected at lower frequency (Table 3). Interestingly, only one fecal sample from chicken was positive for A. lanthieri whereas one fecal sample from sheep was positive for A. faecis . On the other hand, duck, goat, and pony fecal samples were negative for both target species. Similar to the water samples, a low frequency of co-occurrence of both species in only four (cow: n = 2; human: n = 1; pig: n = 1) fecal samples was observed. Additional comparative analysis showed, overall, no significant (p > 0.05) difference  in the rate of prevalence of A. faecis and A. lanthieri between human and animal fecal samples was observed. Similarly, no significant difference between the rate of prevalence of A. faecis and A. lanthieri was found among human, cat, and dog fecal samples. However, a significantly higher frequency of occurrence (p < 0.05) of A. faecis than A. lanthieri was observed between cow and pig fecal samples.

Table 3 Total number (216) of human and animal fecal samples tested and positive  for A. faecis (61) and A. lanthieri (22) using species-specific qPCR assays

Furthermore, the total cell concentrations of the 124 A. faecis and 13 A. lanthieri positive surface water samples ranged from 2.57 × 103 to 2.29 × 107 cells 100 mL− 1 and 1.15 × 104 to 4.16 × 105 cells 100 mL− 1, respectively. However, the 112 A. faecis positive surface water samples gave 103 to 105 cells 100 mL− 1, where only 12 of the positive samples had high concentrations (106 to 107 cells 100 mL− 1) compared to 13 A. lanthieri positive samples that had 104 to 105 cells 100 mL− 1. Two agricultural sampling sites (1 and 8) had similar maximal levels of A. faecis cell concentrations (1.9 × 107 and 2.3 × 107 100 mL- 1) (Fig. 4). Although A. lanthieri was not detected in all agricultural sites, a similar average level (1 × 105 cells 100 mL− 1) of cell concentrations was observed in sampling sites 1, 6, 9, 18, 20, 21, and 253. The cell concentrations of A. lanthieri were lower than A. faecis across all sites (Fig. 4). Results were further analyzed across sample sites: of the top 5% non-zero quantitative data from sites 1, 8, and 9, 1% of samples showed the cell concentrations above 4.1 × 105, 2.1 × 105, and 1.2 × 105 cells 100 mL− 1 of A. faecis as compared to A. lanthieri where 1% of samples of site 1 had cell concentration above 1.2 × 104 cells 100 mL− 1.

Fig. 4

Cell concentration (Log10 cells 100 mL− 1) of A. lanthieri and A. faecis in surface water samples

The cell concentrations of 61 A. faecis and 22 A. lanthieri positive  fecal samples ranged from 1.4 × 100 to 1.2 × 107 cells g− 1 and 3.8 × 101 to 2.0 × 106 cells g− 1, respectively. Of the total 83 positive fecal samples for both species, 45 (54%) A. faecis and 21 (25%) A. lanthieri positive samples had cell concentration ranging from 103 to 105 cells g− 1, while six (10%) A. faecis and one (4%) A. lanthieri positive samples had higher cell concentrations in the range of 106 and 107 cells g− 1. When examining cell concentrations further across each fecal source, the highest average cell concentrations of A. faecis was found in human (2.3 × 106 cells g− 1), cow (7.1 × 105 cells g− 1), and sheep (2.4 × 105 cells g− 1) compared to A. lanthieri where highest average levels were found in chicken (3.4 × 105 cell g-1) and pig (2.7 × 105 cells g− 1) fecal samples (Fig. 5). However, the highest cell concentration of A. faecis was observed in human (1.2 × 107 cells g-1) and cow (3.3 × 106 cells g− 1), while A. lanthieri had the highest cell concentrations in pig (2.0 × 106 cells g-1) and cow (7.8 × 105 cells g− 1) fecal samples. However, fecal samples from dog had the lowest cell concentrations for both A. faecis (4.8 × 102 cells g-1) and A. lanthieri (3.8 × 101 cells g− 1), respectively. Overall, of the 10% non-zero quantitative data, only 1% A. faecis and A. lanthieri positive samples had cell concentration above 2.9 × 106 and 7.1 × 105 cells g− 1, respectively.

Fig. 5

Cell concentration (Log10 cells g− 1) of A. lanthieri and A. faecis in humand and animal  fecal samples


Conventional culture-based multiplex PCR assays for the detection of A. faecis and A. lanthieri, along with four other closely related Aliarcobacter spp., were developed by Khan et al. [15]. In the present study, we further established species-specific direct DNA-based real-time quantitative PCR assays to improve the detection method for rapid identification and quantification of total number of (viable and non-viable) cells of A. faecis and A. lanthieri in surface water and fecal samples. Each species-specific qPCR assay is rapid, sensitive, and reliable for quantitative analysis of A. faecis and A. lanthieri DNA. The assay has a reproducible detection limit per reaction with linear amplification over a wide range of seven to eight orders of magnitude. qPCR assays are less time- and labor-intensive than culture-based methods, and have minimum potential for cross-contamination; therefore, the assays developed here are more robust and useful in diagnostic and analytical settings, especially when the cells of the target species are present at low concentrations [23, 24]. The other advantage is that these assays do not require post-PCR confirmation, and possess the ability to provide quick results which are more desirable for high-throughput studies [25, 26]. In addition, the fluorescent dye SYBR Green was used in the developed assays, which is more cost-effective than fluorogenic probes. qPCR assays can also detect and quantify total (viable and non-viable) number of cells, which is important as the non-viable cells can generate human immunological responses despite these cells being incapable of causing infection. Therefore, the present qPCR assays we have developed allow quantitative detection of these species from complex environmental samples even when they are present at low levels.

To validate the newly developed assays, this study analyzed 588 water samples from an agriculturally dominated watershed and 216 samples from various fecal sources, and A. lanthieri and A. faecis were detected and quantified. Overall, we found that A. faecis was more prevalent and occurred at higher levels than A. lanthieri in both fecal and water samples. Of the SNR water sites sampled, sites 1, 8, and 9 had significantly higher levels of A. faecis compared to sites 1 and 18 where higher relative levels of A. lanthieri were detected. Site 1, a drinking water intake source site, and site 8 are located on the main South Nation River stem, compared to sites 9 and 18 which are located on small stream orders that are closer to livestock operations. Although occurrence of A. lanthieri was not significantly different across the water sampling sites, it was most prevalent at site 21, a small agricultural drainage ditch [27] where potential fecal inputs from adjacent farm lands and wildlife  can occur readily due to tile drainage and surface runoff [28].

Levican et al. [29] found that cell counts for adhesion and invasion of different Aliarcobacter spp. were possible above the limits of 1.7 × 104 CFU mL− 1 and 1.7 × 102 CFU mL- 1, respectively. The cell concentrations of A. lanthieri and A. faecis that we detected here ranged as high as 107 cells 100 mL− 1. Our findings are in congruence with a previous study [30] where a comparable range of concentration (2.0 × 105 to 1.2 × 109 cells 100 mL− 1) of Arcobacter spp. in various water sources was reported.

In order to compare the rate of prevalence of these species in agriculture and non-agricultural surface waters, site 24 was chosen as a reference site, as it is not impacted by any known direct anthropogenic activity [31]. However, A. faecis was detected at this site which suggests that there may be alternate sources of water contamination, possibly from wildlife. However, in previous studies human-specific bacterial markers were detected at site 24 [32, 33]. Throughout the sampling period, among agriculturally dominated SNR sites, A. faecis was most frequently detected at sites 5, 6, 10, 18, and 20 that have dairy operations in the upstream vicinity (Table 3). Additionally, A. lanthieri was most frequently detected at site 21 (Table 3), where dairy-based farming operations occur along the drainage ditch.

The prevalence of other microbial species in the SNR watershed has previously been examined, which add value in our capacity to detect A. lanthieri and A. faecis in the same study area. For example, Lyautey et al. [34] investigated the prevalence of Listeria monocytogenes, and also found that occurrence was associated with proximity to dairy farming operations. The authors found that sites 9 and 18 had the highest prevalence of L. monocytogenes. However, our results showed high frequency of A. faecis in site 18 compared to site 9. Frey et al. [35] detected Campylobacter spp. and Salmonella spp. at the same SNR watershed sites where cattle fecal markers were detected. A. lanthieri and A. faecis were originally isolated from human and fecal sources [4, 5], and in this study both species were detected in human and livestock feces, as well as in agricultural surface water. This strongly indicates that contamination of water by fecal matter from livestock, particularly cattle, could be linked to the prevalence of A. lanthieri and A. faecis.


The qPCR assays designed here can accurately detect the prevalence and quantify the total number of cells of A. faecis and A. lanthieri in complex environmental niches. It is critical to develop alternative methods other than the widely-used culture-based techniques for the detection of gram-negative bacteria in environmental or clinical samples, as the presence of virulence, antibiotic resistance and toxin (VAT) genes can still pose a health risk even when cells are in a non-viable state. The study results suggest that routine quantitative testing of water sources for microbial contamination is important, especially in areas such as agricultural and urban communities where fecal contamination risks are higher. The developed assays could, therefore, provide rapid DNA-based tools for early and reliable detection of target species in field samples, which would help in improving water quality and intervention for reducing and eliminating the risk of contamination of A. faecis and A. lanthieri in aquatic sources.


qPCR assay development and optimization

Bacterial species and culture conditions

For testing the specificity and sensitivity of primers and real-time qPCR assays for the detection and identification of A. faecis and A. lanthieri, two reference strains of A. faecis LMG 28519 and A. lanthieri LMG 28516, were used as positive controls (Table 1). Six other Aliarcobacter spp., nine species from genus  Arcobacter, Haloarcobacter, Malacobacter and Pseudoarcobacter, and 50 other bacterial reference species and strains were used as negative controls (Table 1). In addition to the two LMG strains above, 29 A. faecis and 10 A. lanthieri cultures of our lab collection, isolated from various human and animal fecal and water samples, were used as positive controls. All control reference strains were grown on selective media according to appropriate aerobic and microaerophilic culture conditions. A. faecis and A. lanthieri strains were grown in Arcobacter media broth and incubated at 30 °C under microaerophilic (85% N2, 10% CO2 and 5% O2) conditions with continuous shaking at 125 rpm.

DNA extraction from pure cultures of reference strains and field isolates

The DNA from pure cultures of reference strains and field isolates was extracted using a boiling method [36] where a single colony was suspended in 75 μL TE (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) buffer, boiled for 10 min and centrifuged. The supernatant containing DNA was quantified using a Qubit 3.0 fluorometer (Thermo Fisher Scientific, Waltham, MA, USA), transferred to a sterile tube, and stored at − 20 °C for further PCR analysis.

Spiked assay for standard curve development and quantitation

A spiking experiment was carried out to develop standard curves using A. faecis LMG 28519 and A. lanthieri LMG 28516 reference strains to assess the purity of nucleic acid in terms of yield, concentration, reproducibility and removal of potential PCR-inhibitory compounds. The experiment also helped to quantify and measure the sensitivity (least number of cells mL− 1) of the qPCR assays. A. faecis and A. lanthieri cells were grown under microaerophilic conditions as described above. The cells were collected by centrifugation at room temperature and re-suspended in 1 mL TE buffer. The cell concentration mL− 1 of each target species was measured on modified Arcobacter Agar Medium (m-AAM; Oxoid) containing selective antimicrobial agents (cefoperazone, amphotericin B, and teicoplanin) and incubated under conditions as described above. The known quantity of A. faecis or A. lanthieri reference strain cells (108 cells mL− 1) was then simultaneously spiked and serially (10-fold) diluted from 108 to 101 cells mL− 1 in autoclaved agricultural watershed water and cow manure samples. Each spiked water sample with known cell concentration was filtered through a 0.22 μm sterile nitrocellulose filter.

Total genomic DNA was extracted from each spiked filter and 0.5 g manure sample with known cell concentration using DNeasy PowerSoil Kit (Qiagen; formerly MoBio PowerSoil DNA Isolation Kit) following the manufacturer’s instructions. The purity and concentration of DNA was measured by Qubit 3.0 fluorometer and 1% agarose gel electrophoresis using 1X TAE (0.04 M Tris-acetate, 0.001 M EDTA, pH 7.8) buffer.

Primer design and qPCR assay conditions

Real-time qPCR assays were developed and optimized for A. lanthieri by designing primer pairs from the variable region of the gyrase (gyrA) gene. The primers were designed based on alignment analysis of gyrA gene sequences of Aliarcobacter and other reference species and strains belong to other genera available in the GenBank database. On the other hand, the real-time PCR assay for A. faecis was optimized by using primers from the rpoB gene encoding β-subunit of RNA polymerase previously designed by Khan et al. [15].

For each target species, a SYBR Green-based species-specific monoplex real-time qPCR amplification protocol was developed and optimized with a 20 μL reaction mixture containing 10 μL SsoAdvanced EvaGreen Supermix (Bio-Rad, Hercules, CA, USA), 10–50 ng μL− 1 of purified DNA template of each target species, 0.01 uM forward and reverse primers (Table 4), 5% dimethyl sulfoxide (DMSO), and 0.1 uM Bovine Serum Albumin (BSA). The final volume was adjusted to 20 μL by adding sterile water.

Table 4 Oligonucleotide primer pairs and protocol used for optimized species-specific real-time qPCR-based assays

The reactions were run on a Lightcycler® 480 Instrument II (Roche, Indianapolis, IN, USA) with an initial denaturation at 98 °C for 3 min followed by 50 cycles of denaturation at 98 °C for 15 s, annealing temperatures of 58 °C for A. faecis and 55 °C for A. lanthieri for 30 s, and extension at 72 °C for 30 s. The amplified product obtained from each cell number was confirmed by melt curve analysis where the melting peak was 79 and 74 °C for A. faecis and A. lanthieri, respectively (Table 4). Due to expected small amplicon sizes, the amplified products were further confirmed on a 2% agarose gel matrix, stained (ethidium bromide 0.5 μg mL− 1) and visualized on a UV transilluminator using an Alpha Imager (Fisher Scientific) gel documentation system.

Validation and application of qPCR assays

Study site description, and surface water and fecal sample collection

The developed qPCR assays were further validated and applied to investigate the rate of prevalence and concentration of A. faecis or A. lanthieri cells in surface water and fecal samples. In order to assess the purity of total DNA in terms of removal of PCR inhibitors (such as humic acid, organic and inorganic compounds) and sensitivity of assays for quantitation of total number of cells, surface water samples were collected from the South Nation River (SNR) watershed, located near Ottawa, Ontario in eastern Canada [37]. The watershed covers an area of ~ 3900 km2, of which approximately 60% is used for agricultural purposes, primarily related to dairy farming. A detailed description of the watershed and sampling sites have been previously reported by Wilkes et al. [37, 38] and Lapen et al. [39] (Table 2). For this study, a total of 12 sites of varying stream orders were selected for sampling, based on their proximity to agriculturally-impacted areas. In addition, one site with no known upstream anthropogenic activity was selected as a reference site (Site 24; Edge et al.) [31] (Fig. 6). A total of 588 (from 2013 to 2018) surface water samples were collected on a bi-weekly basis between April and November. In addition, a total of 216 fecal samples from human (n = 19) and various animal (n = 197) sources including cat (n = 20); chicken (n = 8); cow (n = 68); dog (n = 18); duck (n = 1); goat (n = 4); pig (n = 75); pony (n = 2) and sheep (n = 1) were collected in the same region.

Fig. 6

South Nation River watershed area map showing twelve sites selected for the study

The surface water and fecal samples were collected in sterile polypropylene bottles and bags, placed in coolers and delivered to Agriculture and Agri-Food Canada-Ottawa, Ontario Laboratory where the samples were processed within 24 h of their collection for microbiological analysis. Water samples were filtered through 0.22 μm sterile nitrocellulose filters. The DNA from filters and fecal samples were extracted using DNeasy PowerSoil Kit and quantified by Qubit 3.0 fluorometer.

Quantitation of A. faecis and A. lanthieri cell concentration in environmental sources

The two optimized real-time qPCR assays described above were validated, using the developed standard curves, by detecting and quantifying the total number (viable and non-viable) of A. faecis and A. lanthieri cells 100 mL− 1 from agricultural surface water and fecal samples. The specificity and quality of amplified products were confirmed by analyzing and comparing the melting curves to the standard melting peaks obtained for A. faecis and A. lanthieri amplicons. In addition, the amplification quality was also validated by agarose gel electrophoresis using 100 bp DNA size marker (Thermo Fisher Scientific) (Fig. S2A&B). The gel was stained, visualized, and photographed as described in the preceding section.

Data analysis

McNemar Chi-square Contingency and Fisher’s Exact tests were applied to compare the rate of prevalence and identify significant differences (p < 0.05) of A. faecis and A. lanthieri among different agricultural and non-agricultural sites, surface water and fecal samples using STATISTICA (StatSoft, Inc., 2013) [40].

Availability of data and materials

The data generated and analyzed in this study are available upon request.



Dimethyl sulfoxide


Bovine Serum Albumin


modified Arcobacter Agar Medium


South Nation River


gyrase gene


  1. 1.

    Pérez-Cataluña A, Salas-Massó N, Diéguez AL, Balboa S, Lema A, Romalde JL, Figueras MJ. Revisiting the taxonomy of the genus Arcobacter: getting order from the chaos. Front Microbiol. 2018;9:2077.

  2. 2.

    Whiteduck-Léveillée K, Whiteduck-Léveillée J, Cloutier M, Tambong JT, Xu R, Topp E, Khan IUH. Arcobacter lanthieri sp. nov., isolated from pig and dairy cattle manure. Int J Syst Evol Microbiol. 2015;65:2709–16.

  3. 3.

    Whiteduck-Léveillée K, Whiteduck-Léveillée J, Cloutier M, Tambong JT, Xu R, Topp E, Khan IUH. Identification, characterization and description of Arcobacter faecis sp. nov., isolated from a human waste septic tank. Syst Appl Microbiol. 2016;39:93–9.

  4. 4.

    Zambri M, Cloutie M, Adam Z, Lapen DR, Wilkes G, Sunohara M, Topp E, Talbot G, Khan IU. Novel virulence, antibiotic resistance and toxin gene-specific PCR-based assays for rapid pathogenicity assessment of Arcobacter faecis and Arcobacter lanthieri. BMC Microbiol. 2019;19(11):1–15.

  5. 5.

    Ferreira S, Queiroz JA, Oleastro M, Domingues FC. Insights in the pathogenesis and resistance of Arcobacter: a review. Crit Rev Microbiol. 2016;42(3):364–83.

  6. 6.

    Atabay HI, Aydin F, Hou K, Sahin M, Vandamme P. The prevalence of Arcobacter spp. on chicken carcasses sold in retail markets in Turkey, and identification of the isolates using SDS-PAGE. Int J Food Microbiol. 2003;81:21–8.

  7. 7.

    Collado L, Guarro J, Figueras MJ. Prevalence of Arcobacter in meat and shellfish. J Food Prot. 2009;72(5):1102–6.

  8. 8.

    Villarruel-López A, Márquez-González M, Garay-Martinez LE, Zepeda H, Castillo A, de la Garza LM, Torres-Vitela R. Isolation of Arcobacter spp. from retail meats and cytotoxic effects of isolates against vero cells. J Food Prot. 2003;66(8):1374–8.

  9. 9.

    Fera MT, Maugeri TL, Gugliandolo C, Beninati C, Giannone M, La Camera E, Carbone M. Detection of Arcobacter spp. in the coastal environment of the Mediterranean Sea. Appl Environ Microbiol. 2004;70(3):1271–6.

  10. 10.

    Rice EW, Rodgers MR, Wesley IV, Johnson CH, Tanner SA. Isolation of Arcobacter butzleri from ground water. Lett Appl Microbiol. 1999;28:31–5.

  11. 11.

    Collado L, Levican A, Perez J, Figueras MJ. Arcobacter defluvii sp. nov., isolated from sewage samples. Int J Syst Evol Microbiol. 2010;61:2155–61.

  12. 12.

    Houf K, Stephan R. Isolation and characterization of the emerging foodborn pathogen Arcobacter from human stool. J Microbiol Methods. 2007;68:408–13.

  13. 13.

    van Driessche E, Houf K, van Hoof J, De Zutter L, Vandamme P. Isolation of Arcobacter species from animal feces. FEMS Microbiol Lett. 2003;229:243–8.

  14. 14.

    van Driessche E, Houf K, Vangroenweghe F, De Zutter L, Van Hoof J. Prevalence, enumeration and strain variation of Arcobacter species in the faeces of healthy cattle in Belgium. Vet Microbiol. 2005;105:149–54.

  15. 15.

    Khan IUH, Cloutier M, Libby M, Lapen DR, Wilkes G, Topp E. Enhanced single-tube multiplex PCR assay for detection and identification of six Arcobacter species. J Appl Microbiol. 2017;123(6):1522–32.

  16. 16.

    Diéguez AL, Balboa S, Magnesen T, Romalde JL. Arcobacter lekithochrous sp. nov., isolated from a molluscan hatchery. Int J Syst Evol Microbiol. 2017;67(5):1327–32.

  17. 17.

    Fera MT, Maugeri TL, Gugliandolo C, La Camera E, Lentini V, Favaloro A, Carbone M. Induction and resuscitation of viable nonculturable Arcobacter butzleri cells. Appl Environ Microbiol. 2008;74(10):3266–8.

  18. 18.

    Li L, Mendis N, Trigui H, Oliver JD, Faucher SP. The importance of the viable but non-culturable state in human bacterial pathogens. Front Microbiol. 2014;5:258.

  19. 19.

    Abdelbaqi K, Buissonnière A, Prouzet-Mauleon V, Gresser J, Wesley I, Mégraud F, Ménard A. Development of a real-time fluorescence resonance energy transfer PCR to detect Arcobacter species. J Clin Microbiol. 2007;45(9):3015–21.

  20. 20.

    Brightwell G, Mowat E, Clemens R, Boerema J, Pulford DJ, On SL. Development of a multiplex and real time PCR assay for the specific detection of Arcobacter butzleri and Arcobacter cryaerophilus. J Microbiol Methods. 2007;68:318–25.

  21. 21.

    de Boer RF, Ott A, Güren P, van Zanten E, van Belkum A, Kooistra-Smid AMD. Detection of Campylobacter species and Arcobacter butzleri in stool samples by use of real-time multiplex PCR. J Clin Microbiol. 2013;51(1):253–9.

  22. 22.

    González A, Suski J, Ferrús MA. Rapid and accurate detection of Arcobacter contamination in commercial chicken products and wastewater samples by real-time polymerase chain reaction. Foodborne Path Dis. 2010;7(3):327–38.

  23. 23.

    Girones R, Ferrús MA, Alonso JL, Rodriguez-Manzano J, Calgua B, de Abreu Corrêa A, Bofill-Mas S. Molecular detection of pathogens in water–the pros and cons of molecular techniques. Water Res. 2010;44(15):4325–39.

  24. 24.

    He J-W, Jiang S. Quantification of enterococci and human adenoviruses in environmental samples by real-time PCR. Appl Environ Microbiol. 2005;71(5):2250–5.

  25. 25.

    Bellin T, Pulz M, Matussek A, Hempen HG, Gunzer F. Rapid detection of enterohemorrhagic Escherichia coli by real-time PCR with fluorescent hybridization probes. J Clin Microbiol. 2001;39(1):370–4.

  26. 26.

    Sails AD, Fox AJ, Bolton FJ, Wareing DRA, Greenway DLA. A real-time PCR assay for the detection of Campylobacter jejuni in foods after enrichment culture. Appl Environ Microbiol. 2003;69(3):1383–90.

  27. 27.

    Wilkes G, Brassard J, Edge TA, Gannon V, Gottschall N, Jokinen CC, Jones TH, Khan IUH, Marti R, Sunohara MD, Topp E. Long-term monitoring of waterborne pathogens and microbial source tracking markers in paired agricultural watersheds under controlled and conventional tile drainage management. Appl Environ Microbiol. 2014;80(12):3708–20.

  28. 28.

    Chen W, Wilkes G, Khan IUH, Pintar KDM, Thomas JL, Lévesque CA, Lapen DR. Aquatic bacterial communities associated with land use and environmental factors in agricultural landscapes using a metabarcoding approach. Front Microbiol. 2018;9:2301.

  29. 29.

    Levican A, Alkeskas A, Günter C, Forsythe SJ, Figueras MJ. Adherence to and invasion of human intestinal cells by Arcobacter species and their virulence genotypes. Appl Environ Microbiol. 2013;79(16):4951–7.

  30. 30.

    Shrestha RG, Tanaka Y, Malla B, Tandukar S, Bhandari D, Inoue D, Sei K, Sherchand JB, Haramoto E. Development of a quantitative PCR assay for Arcobacter spp. and its application to environmental water samples. Microbes Environ. 2018;33(3):309–16.

  31. 31.

    Edge TA, El-Shaarawi A, Gannon V, Jokinen C, Kent R, Khan IUH, Koning W, Lapen D, Miller J, Neumann N, Phillips R. Investigation of an Escherichia coli environmental benchmark for waterborne pathogens in agricultural watersheds in Canada. J Environ Qual. 2012;41(1):21–30.

  32. 32.

    Lyautey E, Lu Z, Lapen DR, Wilkes G, Scott A, Berkers T, Topp E. Distribution and diversity of Escherichia coli populations in the South Nation River drainage basin, eastern Ontario, Canada. Appl Environ Microbiol. 2010;6(5):1486–96.

  33. 33.

    Marti R, Gannon VPJ, Jokinen C, Lanthier M, Lapen DR, Neumann NF, Topp E. Quantitative multi-year elucidation of fecal sources of waterborne pathogen contamination in the South Nation River basin using Bacteroidales microbial source tracking markers. Water Res. 2013;47:2315–24.

  34. 34.

    Lyautey E, Lapen DR, Wilkes G, McCleary K, Pagotto F, Tyler K, Topp E. Distribution and characteristics of Listeria monocytogenes isolates from surface waters of the south Nation River watershed, Ontario, Canada. Appl Environ Microbiol. 2007;73(17):5401–10.

  35. 35.

    Frey SK, Topp E, Edge T, Fall C, Gannon V, Jokinen C, Marti R, Neumann N, Ruecker N, Wilkes G, Lapen DR. Using SWAT, Bacteroidales microbial source tracking markers, and fecal indicator bacteria to predict waterborne pathogen occurrence in an agricultural watershed. Water Res. 2013;47(16):6326–37.

  36. 36.

    Khan IUH, Hill S, Nowak E, Palme ME, Jarjanazi H, Lee D-Y, Mueller M, Schop R, Weir S, Abbey A, Winter J, Edge TA. Investigation of the prevalence of thermophilic Campylobacter species at Lake Simcoe recreational beaches. Inland Waters. 2013;3(1):93–104.

  37. 37.

    Wilkes G, Ruecker NJ, Neumann NF, Gannon VP, Jokinen C, Sunohara M, Topp E, Pintar KD, Edge TA, Lapen DR. Spatiotemporal analysis of Cryptosporidium species/genotypes and relationships with other zoonotic pathogens in surface water from mixed-use watersheds. Appl Environ Microbiol. 2013;79(2):434–48.

  38. 38.

    Wilkes G, Edge TA, Gannon VPJ, Jokinen C, Lyautey E, Neumann NF, Lapen DR. Associations among pathogenic bacteria, parasites, and environmental and land use factors in multiple mixed-use watersheds. Water Res. 2011;45(18):5807–25.

  39. 39.

    Lapen DR, Schmidt PJ, Thomas JL, Edge TA, Flemming C, Keithlin J, Neumann N, Pollari F, Ruecker N, Simhon A, Topp E. Towards a more accurate quantitative assessment of seasonal Cryptosporidium infection risks in surface waters using species and genotype information. Water Res. 2016;105:625–37.

  40. 40.

    StatSoft, Inc. Electronic statistics textbook. Tulsa: StatSoft.; 2013.

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We thank the field crew and co-op students Mark Libby, Linda Liu, Ozair Islam and Ferdinand Miranda for sample collection and lab assistance. We also thank farmers for their cooperation in accessing farms for water and fecal sample collection.


The study funds were provided by Agriculture and Agri-Food Canada under project #s: J-000157 and J-001012. The funding source was not involved in study design, sample collection, data analysis and interpretation.

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MM, MC, and IK conceived research project, performed sequence analyses and designed the experiments. MM, MC and IK designed primers and qPCR protocols, performed laboratory experiments, analyzed results and drafted manuscript. EC, GW, DL and ET coordinated in selecting field sampling sites and sample collection protocols as well as contributed in interpreting results data and editing manuscript. All authors reviewed, edited and approved the final manuscript.

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Correspondence to Izhar U. H. Khan.

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Additional file 1: Supplementary Figure 1A&B. Real-time qPCR amplified product confirmation on 2% agarose gel for A. faecis LMG 28519 reference strain and field isolates (Panel A; Lanes 1–4) and A. lanthieri reference strain LMG 28516 and field isolates (Panel B; Lanes 1–4) with an expected 152 and 72 bp sizes, respectively. Lanes 5 and 11: A. butzleri, A. cryaerophilus, A. skirrowii, A. thereius, A. trophiarum, A. cibarius and no DNA template (PCR reaction mix) served as negative controls; M: 100 bp DNA size marker. Supplementary Figure 2A&B. Real-time qPCR amplified product confirmation on 2% agarose gel showing positive and negative field samples for A. faecis (Panel A) and A. lanthieri (Panel B) with an expected 152 and 72 bp sizes, respectively. Lane 1: A. faecis and A. lanthieri reference strains served as positive controls; M: 100 bp DNA size marker.

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Miltenburg, M.G., Cloutier, M., Craiovan, E. et al. Real-time quantitative PCR assay development and application for assessment of agricultural surface water and various fecal matter for prevalence of Aliarcobacter faecis and Aliarcobacter lanthieri. BMC Microbiol 20, 164 (2020).

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  • qPCR
  • Aliarcobacter faecis
  • Aliarcobacter lanthieri
  • Agricultural watershed
  • Surface water
  • Assay
  • Fecal matter